This work supported has been supported in part by the National Science Foundation under Grant No. EFRI 0938047, and by the Virginia Tech Institute for Critical Technology and Applied Science (ICTAS). The authors would like to express appreciation to Dr. Eva Schmelz and Dr. Paul Roberts for their kind gift of MOSE-L cells. The authors acknowledge Angela Anderson for her assistance with cell culture, Caitlan Swaffar for her help with editing this document and preparing experiments, and all Bioelectromechanical Systems lab members.

1. Fabricating a Prototype cDEP Microfluidic Device: Overview
<ol>
	<li>Follow previously reported procedures22 using photoresist and Deep Reactive Ion Etching (DRIE) to etch the desired channel design (Figure 1) into a silicon wafer (Figure 2). Deposit a thin layer of Teflon to improve the release of the microdevice from the wafer.</li>
</ol>
<img alt="Figure 1" src="/files/ftp_upload/50634/50634fig1.jpg" /> 
Figure 1. Schematic of a low frequency continuous sorting device. The sample channel runs left to right and is 500 &#956;m wide with sawtooth constrictions to 100 &#956;m. The two pairs of fluidic electrode channels compose the source and sink electrodes, respectively, and are separated from the sample channel by a 20 &#956;m thick PDMS barrier.
<img alt="Figure 2" src="/files/ftp_upload/50634/50634fig2.jpg" /> 
Figure 2. The fabrication process of a cDEP device. (a) For 60 sec, UV light selectively reacts with photoresist exposed through a mask pattern of the cDEP device geometry. (b) Photoresist is removed using developer. (c) Deep Reactive Ion Etching (DRIE) is used to etch 50 &#956;m deep features, and 5 min of wet etching by tetramethylammonium hydroxide (TMAH) 25% at 70 &#176;C is used to reduce roughness on the side walls. (d) A Teflon coating is added to improve device release from the wafer. (e) PDMS is poured onto the re-usable wafer stamp after degassing and cured for 45 min at 100 &#176;F. (f) PDMS is removed from the wafer. PDMS and a clean glass slide are exposed to air plasma for 2 min and bonded together.
<ol start="2">
	<li>Wrap the wafer with aluminum foil to prevent spill-over. Mix polydimethylsiloxane (PDMS) in 10:1 ratio of elastomer to curing agent, de-gas for approximately 20 min, and pour onto the wafer. Heat for 45 min at 100 &#176;C to avoid damaging the Teflon coating of the stamp. Allow the device to cool down.</li>
	<li>After cooling, remove aluminum foil, trim PDMS using a surgical blade, and punch access holes at channel inlet/outlet using a blunt puncher suited to the size of tubing. Here, a 1.5 mm blunt puncher is used.</li>
	<li>Clean a glass microscope slide using a rinse of soap and water, ethanol, DI water, and drying with air, respectively. Clean the PDMS device using Scotch Magic tape. Examine under microscope to be sure channels are clean. Expose the slide and PDMS device to air plasma for 2 min. Firmly press channel side of device to glass slide, avoiding formation of air bubbles between the device and the slide (Figure 3).</li>
</ol>
<img alt="Figure 3" fo:content-width="6in" fo:src="/files/ftp_upload/50634/50634fig3highres.jpg" src="/files/ftp_upload/50634/50634fig3.jpg" /> 
Figure 3. (a) The mask used for photolithography during the silicon wafer stamp fabrication, and (b) the finished PDMS-glass device with the sample and fluid electrode channels filled with green and red food coloring, respectively. See Figure 1 for dimensions. <a href="https://www.jove.com/files/ftp_upload/50634/50634fig3highres.jpg" target="_blank">Click here to view larger figure</a>.
2. Preparing a Cell Suspension in DEP Buffer
<ol>
	<li>Prepare a low conductivity (100 &#956;S/cm) buffer, which will be referred to as &#34;DEP buffer&#34; (8.5% sucrose [w/v], 0.3% glucose [w/v], 0.725% RPMI [v/v]).16</li>
	<li>Suspend cells in DEP buffer at 3 x 106 cells/ml. Here, cancerous late-stage mouse ovarian surface epithelial (MOSE-L) cells are used.</li>
</ol>
Note: Cell viability analysis by the trypan blue dye exclusion method showed a slight decrease in viability (from 95.1-91.6%) of early-stage MOSE (MOSE-E) cells suspended in DEP buffer at room temperature after 5 hr. It is predicted that a similar slight decrease in viability occurs for MOSE-L cells, indicating that cells should not be stored in DEP buffer long-term.
<ol start="3">
	<li>Dye cells using a fluorescent membrane-permeable dye, such as the enzymatically-activated Calcein AM.</li>
	<li>Measure conductivity of the dyed cell suspension. The conductivity should be approximately 100 &#956;S/cm. If the conductivity measurement is too high, spin the sample down, resuspend in DEP buffer at 3 x 106 cells/ml and repeat the conductivity measurement.</li>
</ol>
3. Loading the cDEP Microfluidic Chip
<ol>
	<li>Place the microfluidic chip under vacuum for 30 min.</li>
	<li>Meanwhile, cut a piece of flexible tubing to span the distance from the syringe pump to the microscope stage where the microfluidic chip will be located. Here, a 6 inch length of tubing is required. Fit tubing to needle tip on syringe.</li>
	<li>Estimate the required length for outlet tubing by dividing the target collected sample volume by the cross-sectional area of the tube. Cut the require length of tubing.</li>
	<li>Draw cell suspension into syringe. Check that no bubbles are located in the tubing or syringe.</li>
	<li>Upon removing chip from vacuum, insert the outlet tubing and prime the sample channel quickly to avoid trapped air bubbles in the channels. Gently tap the syringe when priming to avoid rupturing the thin (20 &#956;m) insulating barrier, though it has been observed that the barrier can withstand a constant high throughput near 5 ml/hr without rupturing.</li>
	<li>To prime the electrode channels, quickly place empty pipette tips in one outlet of each fluidic electrode channel. Pipette 200 &#956;l of PBS containing a small amount of rhodamine B and gently tap to introduce the fluid into the opposite end of the electrode channel. Maintain gentle pressure until PBS with rhodamine B visibly progresses up the tip of the empty pipette tip. Repeat for each electrode channel. Rhodamine B is used only to fluorescently visualize the fluidic electrode channels.</li>
	<li>After priming, fill each pipette tip reservoir with PBS with rhodamine B. Avoid bubble formation in the pipette tip reservoirs, and remove any visible bubble by pipetting gently up and down. Detach the outlet tubing and discard appropriately, then insert a fresh outlet tubing reservoir to collect the target volume.</li>
</ol>
4. Characterizing Crossover Frequency of Cells using cDEP Chip
<ol>
	<li>Position the chip on the stage of an inverted microscope equipped with digital camera (Figure 4).</li>
</ol>
<img alt="Figure 4" fo:content-width="5in" fo:src="/files/ftp_upload/50634/50634fig4highres.jpg" src="/files/ftp_upload/50634/50634fig4.jpg" /> 
Figure 4. (a) Overall experimental setup for cDEP experiments. The cDEP chip is located on the platform of the inverted microscope. A camera sends images to the computer for recording. The syringe pump maintains a constant inlet flow. The function generator generates a signal which is stepped up by the high voltage amplifier and connected to the cDEP chip by wire electrodes. The oscilloscope monitors the signal sent to the chip. (b) Detail view of the cDEP chip and fluidic and electronic connections. <a href="https://www.jove.com/files/ftp_upload/50634/50634fig4highres.jpg" target="_blank">Click here to view larger figure</a>.
<ol start="2">
	<li>Mount syringe on syringe pump and insert wire electrodes into fluidic electrode channel reservoirs.</li>
	<li>Pump fluid through at 1 ml/hr to ensure good contact between the syringe pump and the syringe. Visually inspect the channel length to ensure unimpeded flow of cells and the absence of bubbles or leaks. Reduce flow rate to 0.005 ml/hr. 
	Note: Throughput as high as 1 ml/hr31,35 has been achieved in devices of other geometries.</li>
	<li>For safety, test the electronics by powering on the function generator, high voltage amplifier, and oscilloscope, and adjusting to desired voltage and frequency. Here these parameters are 200 V and 20 kHz. Upon verification of acceptable performance, power off. Connect electrodes to high voltage amplifier. 
	Note: Observe appropriate electrical safety procedures when operating the electronics at the high voltages used for these experiments.</li>
	<li>Upon achieving steady flow, apply the desired voltage at the desired frequency. In this experiment, voltage is 200 VRMS at frequencies between 5-60 kHz.</li>
	<li>Record video files of the cell response with image processing techniques (step 5) to quantify cell distribution in the channel and to characterize the crossover frequency. To do this, maintain a constant voltage and vary the frequency systematically. At the start and end of the experiment, as well as randomly between experimental runs, record the control cell distribution, the distribution of the cells without applying any electrical field. Estimate the amount of time required for cells to flow from the inlet to the monitored location (here, 5 min). After setting a new frequency, wait this amount of time, then begin recording (here, a 2 min video of cell distribution).</li>
</ol>
5. Image Processing for Characterizing Crossover Frequency
<ol>
	<li>Using a computational script, analyze each video frame by frame to determine the relative y-position (where the z-direction is the vertical depth of the channel, the y-direction is the channel width, and x-direction is the channel length) of each fluorescing cell or particle at a specified x-coordinate downstream from the region of high DEP force in the channel. Create a graph of the relative distribution.</li>
	<li>Compare the centerline of the distribution at each voltage and frequency with the centerline of the control distribution. For a given voltage and varying frequencies, determine the crossover frequency, where the centerline matches the y-position of the control.</li>
</ol>
6. Varying the Experiment to Perform Cell Sorting or Enrichment
<ol>
	<li>Suspend desired mixture in DEP buffer at a total concentration of 3 x 106 particles/ml. Here, this mixture is MOSE-L cells with 4 &#956;m fluorescing beads in DEP buffer. Perform the previously described steps to prepare a cDEP device.</li>
	<li>To sort or enrich cells from a mixture, determine the crossover frequencies of the target cells and the background particles as previously described. Operate the experiment at a frequency between the two crossover frequencies of the target cells and the background particles so that the two populations of particles experience DEP forces in opposite directions. To sort MOSE-L cells and 4 &#956;m beads, operate the microdevice above 11.90&#177;0.63 kHz. the first crossover frequency of MOSE-L cells recently reported by Salmanzadeh et al.33</li>
	<li>To collect the contents of a sorted sample, remove the outlet tubing upon collecting the target volume, by placing a glove finger over the outlet opening and gently tugging on the tubing. Pour the collected target volume into a reservoir for further analysis. Note: Other sample recovery methods could include attaching a permanent reservoir to the chip and removing the sample by simple pipetting.</li>
</ol>

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Dr. Rafael Davalos has a patent pending for contactless dielectrophoresis.

DEP is a powerful technique for determining dielectric properties of particles and directing particle motion towards sorting, isolation, or enrichment applications. Due to the detrimental effect of direct electrode contact with a sample, others have taken approaches similar to the previously presented method to avoid contact. For example, Bashir et al. developed contact-free DEP devices by using a microfluidic PDMS device separated from printed circuit board electrodes by a thin glass coverslip, and this technique is also made available in video format.23,36
Here, we have shown the fabrication of a cDEP chip and fluidic electrodes using a single PDMS substrate, and the experimental protocol for separating ovarian cancer cells from a mixture of cells and fluorescent beads. The presented technique has been successfully used for a variety of more complex and physiologically relevant applications, including sorting live and dead cells,28 tumor initiating cells from prostate cancer cells,30 cancer cells from dilute red blood cells,31,32 and differentiating among stages of breast cancer37 and ovarian cancer.29 cDEP has also been used for mixing particles.38 These applications suggest that by using the simple technique presented, diverse purposes can be accomplished simply by altering the design of the channel geometry.
DEP is useful on the microscale for manipulation of particles.13 For spherical particles, the translational dielectrophoretic force depends upon the size and electrical properties of the particle and its suspending medium, as well as the gradient of the electric field squared:
<img alt="Equation 1" src="/files/ftp_upload/50634/50634eq1.jpg" /> 
where &epsilon;m is the permittivity of the suspending medium, r is the radius of the particle, and Rc[k(&omega;)]is the real part of the Clausius-Mossotti (CM) factor. The CM factor is a measure of the relative polarizability of the particle compared to the suspending medium and determines the direction of the dielectrophoretic force. It is described as
<img alt="Equation 2" src="/files/ftp_upload/50634/50634eq2.jpg" /> 
where <img alt="Equation 3" src="/files/ftp_upload/50634/50634eq3.jpg" /> and <img alt="Equation 4" src="/files/ftp_upload/50634/50634eq4.jpg" /> are the complex permittivities of the particle and medium, respectively. The complex permittivity, <img alt="Equation 5" src="/files/ftp_upload/50634/50634eq5.jpg" />, depends upon conductivity (&sigma;) and frequency (&omega;). The CM factor of spherical particles is theoretically bound between -0.5 and 1. If the CM factor is negative, the particles experience nDEP because the medium is more polarizable than the particle, so particles move away from regions of high electric field gradients. If the CM factor is positive, the particles are more polarizable than the medium and they experience pDEP in which they move toward regions of high electric field gradients.
For biological particles that are nonhomogeneous in structure, such as cells, the Clausius-Mossotti factor can be determined from an effective value for the particle permittivity:
<img alt="Equation 6" src="/files/ftp_upload/50634/50634eq6.jpg" /> 
where <img alt="Equation 7" src="/files/ftp_upload/50634/50634eq7.jpg" /> and <img alt="Equation 8" src="/files/ftp_upload/50634/50634eq8.jpg" /> are the complex permittivity of the effective complex permittivity of the interior of the cell, such as the cytoplasm, and the plasma membrane, respectively; r is the radius of the cell, and d is the thickness of the plasma membrane.26
When DEP occurs with particles suspended in a fluid, the motion of the particle relative to the fluid will generate a drag force on the particle. This drag force must be considered when determining the overall force acting on the particle. For the situations of interest here, viscous forces dominate and particles are assumed spherical, small, and moving with relatively low velocity, so Stokes' drag law provides a good approximation for the drag force:
<img alt="Equation 9" src="/files/ftp_upload/50634/50634eq9.jpg" /> 
where &eta; is the viscosity of the fluid, up is the velocity of the particle, and uf is the velocity of the fluid, which may also be moving. Given the dielectrophoretic force, known fluid and particle properties, and a known flow velocity, the balance between the drag force and the dielectrophoretic force can be solved to estimate the particle velocity. The shear rate that the cells experience should be below the threshold at which cell lysing can occur.
Characterization of the electrical properties of particles is necessary to predict and control how they will respond under DEP them. In this work, we specifically utilized low frequency cDEP with MOSE-L cells to demonstrate the protocol for determining first crossover frequency of cells, and then showed continuous sorting of polystyrene beads and MOSE-L cells based on their opposing DEP responses.
Altering the geometry of the cDEP device will change the spatial gradients of the electric field, allowing devices to be designed for high frequency or low frequency operation, and for high selectivity and efficiency of sorting for a specified cell type. Additionally, high throughput devices can be developed by fabricating wider channels,30 channels in parallel,30 or by multilayer fabrication35, in which the electrode channels are vertically stacked above and below a relatively deep sample channel. Thin membranes form the barrier between layers. Preliminary testing with devices fabricated in polymethylmethacrylate (PMMA) and polycarbonate (PC) thin films have demonstrated DEP response of MOSE-L cells. Current efforts are underway to refine multi-layer high-throughput devices and to improve the peripheral system towards an eventual plug-and-play platform. To expand from the basic experimental technique presented, the applications and specifications of the device can be tailored to fit particular demands, such as sorting versus characterization, or adding sample reservoirs and a semi-automated system to the device.

Biological sample enrichment and particle sorting is often necessary for subsequent analysis.1 For instance, isolation of rare cells from body fluids has important applications in cancer detection and individualized medicine. 2,3 The most commonly used enrichment techniques are fluorescent activated cell sorting (FACS)4 and magnetic activated cell sorting (MACS),5 which rely upon expressed surface markers to differentiate cells. Other strategies include hydrodynamic6 or inertial7,8 sorting, optical tweezers,9 acoustophoresis,10 and dielectrophoresis.11,12 Dielectrophoresis is the movement of a polarized particle in the presence of a non-uniform electric field.13 DEP has been used for a wide range of applications,2,14 including sorting cells based on viability,15 characterizing bioelectrical properties of cells,16 and sorting upon induced changes in biophysical properties of cells.17,18 Traditional DEP utilizes planar electrodes patterned within a microfluidic channel to apply a voltage and induce a non-uniform electric field.13 While this is a powerful technique, challenges can arise, such as electrode delamination and electrolysis. Insulator-based dielectrophoresis (iDEP)19 has addressed the challenges of fouling, electrode delamination, and spatial degradation of the electrode field through patterning insulating structures that induce non-uniformities in a DC electric field. iDEP has been used to selectively separate live and dead bacteria cells,19 isolate bacterial spores,20 and manipulate DNA,21 among other applications. Joule heating can be a challenge because it can occur as a result of the high DC voltages often required. To ameliorate these challenges, contact-free DEP microdevices have been developed.22-24
The technique presented here utilizes contactless dielectrophoresis (cDEP), so named for the lack of direct contact between metallic electrodes and the sample channel.22 Unique to this strategy is the replacement of metallic electrodes with fluid electrode channels filled with a highly conductive solution. These fluid electrodes are capacitively coupled across a thin insulating barrier to the sample channel via an AC voltage. Eliminating sample contact with electrodes reduces issues associated with DEP-based methods such as electrolysis and bubble formation, sample contamination, and electrode delamination. As a result, cDEP is especially useful for biological samples because it supports viability of the cells in the sample. Importantly, cDEP can maintain sample sterility. A chip can be prepared in a cell culture hood and the experiment can be conducted without requiring sample contact to metal electrodes or requiring that the sample be open to the environment. A simple reservoir can be fixed to the chip outlet to facilitate sterile sample recovery. Additionally, the fabrication of fluid electrodes from the same biocompatible polymer material (PDMS) as the sample channel reduces the high cost incurred with custom patterning of metal electrodes, the time required for fabrication, and limits the need for specialized cleanroom equipment to the initial patterning of the reusable silicon wafer stamp.
Movement of particles due to DEP depends on the characteristics of the particle and the medium, as well as the spatial gradients of the electric field. A particle- and frequency-dependent factor, called the Clausius-Mossotti (CM) factor, takes a value in the range -0.5 to 1, and determines the direction of the DEP force. The frequency at which the CM factor is exactly zero is called the crossover frequency. This is the point at which no dielectrophoretic force is exerted on a particle and the CM factor changes sign. A single crossover frequency for inert solid microspheres occurs when the CM factor changes from negative to positive. 25 For mammalian cells in low conductivity buffer on the order of 0.01 S/m, a first crossover frequency indicating a transition from nDEP to pDEP exists near 10-100 kHz, and is influenced by the size, shape, cytoskeleton, and membrane properties of the cell.26,27 A second crossover frequency at a shift from the pDEP to nDEP regime is on the order of 10 MHz, and is influenced by the nucleus-cytoplasm ratio, cytoplasm conductivity, and endoplasmic reticulum.27 DEP force can be applied without the presence of fluid flow, but here we utilize a flowing fluid to achieve continuous sorting of suspended particles. The combined influence of the dielectrophoretic force and the Stokes' drag force dictate the translational motion of a particle.
We have developed devices for operation in two frequency ranges. Higher frequency devices (100-600 kHz) have operated using pDEP and achieved batch sorting of cells, such as prostate tumor initiating cells (TICs), murine ovarian surface epithelial (MOSE) cells, MDA-MB-231 breast cancer cells, or live THP-1 cells by selectively trapping cells of interest on insulating posts located in the sample channel.28-31 Lower frequency (5-100 kHz) devices operate continuously, and when operated at a frequency at which one population experiences pDEP while the background population experiences nDEP, can redirect particle trajectories to achieve sorting.32-34 These low frequency devices have been used to sort cancer cells from red blood cells, determine the changes in dielectric properties of a progressive MOSE cell line, and to elucidate the effects of non-toxic sphingolipid treatments on reversing aggressive characteristics of aggressive MOSE cells. Additionally, cDEP microdevices can be designed to operate at increased throughput, currently up to 1 ml/hr.31,35
As described, the flexibility and low cost of the fabrication process enables custom-designed device geometries, which allow the presented experimental procedure to be relevant for a wide range of applications. The long-term goal of cDEP is to realize label-free cell sorting and enrichment on a clinical level, with sample recovery for subsequent culture or processing. The technique presented here is a simple and inexpensive method, from fabrication to experimentation, which increases the accessibility of DEP. We demonstrate the preparation of a cDEP chip and the experimental protocol to achieve characterization and enrichment of ovarian cancer cells from fluorescent microspheres.

<table border="1">
	<tbody>
		<tr>
			<td>Name of Reagent/Material</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Polydimethylsiloxane (PDMS)</td>
			<td>Dow Corning, Midland, MI,USA</td>
			<td>Sylgard 184</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Glass slides</td>
			<td>The Microscope Depot</td>
			<td>76079</td>
			<td>2x3 inch-ground edges</td>
		</tr>
		<tr>
			<td>Microbore PTFE Tubing</td>
			<td>Cole-Parmer Instrument Co, Vernon Hills, IL, USA</td>
			<td>EW-06417-31</td>
			<td>Thin walled 20 gauge, 0.032"ID x 0.056"OD, 100 ft/roll</td>
		</tr>
		<tr>
			<td>Luer-slip plastic syringes</td>
			<td>National Scientific company</td>
			<td>S7510-1</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Needle tip</td>
			<td>Howard Electronic instruments</td>
			<td>JG20-1.0</td>
			<td>20 Gauge 1.0", ID=0.025" OD=0.036"</td>
		</tr>
		<tr>
			<td>D(+)-Sucrose</td>
			<td>Fisher Scientific, Fair Lawn, NJ</td>
			<td>S3-500</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>D(+)-glucose, reagent ACS, anhydrous</td>
			<td>Acros Organics N.V., Fair Lawn, NJ</td>
			<td>AC410955000</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>RPMI-1640 Medium</td>
			<td>Quality Biological Inc.</td>
			<td>112-025-101</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Calcein AM, Molecular Probes</td>
			<td>Invitrogen Corp. (life technologies), Carlsbad, CA, USA</td>
			<td>C3100MP</td>
			<td>excitation wavelength 488/emission wavelength 516</td>
		</tr>
		<tr>
			<td>Rhodamine B, O</td>
			<td>Science Lab</td>
			<td>SLR1465-100G</td>
			<td>excitation wavelength 540/emission wavelength 625</td>
		</tr>
		<tr>
			<td>Phosphate buffered saline (10X)</td>
			<td>Gbiosciences, St. Louis, MO</td>
			<td>RC-147</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Leica, inverted light microscope</td>
			<td>Leica Microsystems, Bannockburn, IL, USA</td>
			<td>Leica DMI 6000B</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Leica DFC420, color camera</td>
			<td>Leica Microsystems, Bannockburn, IL, USA</td>
			<td>Leica DFC420</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Function generator</td>
			<td>GW Instek, Taipei, Taiwan</td>
			<td>GFG-3015</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Wideband power amplifier</td>
			<td>Amp-Line Corp., Oakland Gardens, NY, USA</td>
			<td>AL-50HF-A</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>HFHV Output Transformer</td>
			<td>&nbsp;</td>
			<td>AL-T50-V25/300-F100K-600K</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>High voltage amplifier</td>
			<td>Trek</td>
			<td>Model 2205</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>USB Modular Oscilloscope, 100 MHz</td>
			<td>AgilentTechnologies</td>
			<td>U2701A</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Expanded Plasma Cleaner</td>
			<td>Harrick Plasma</td>
			<td>PDC-001/002 (115/230V)</td>
			<td>air plasma</td>
		</tr>
		<tr>
			<td>Scotch Magic tape</td>
			<td>3M</td>
			<td>&nbsp;</td>
			<td>any available width is sufficient</td>
		</tr>
		<tr>
			<td>1.5 mm puncher</td>
			<td>Harris Uni-Core</td>
			<td>Z708836-25EA</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>.25% Trypsin-EDTA</td>
			<td>Invitrogen</td>
			<td>25200-056</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>FluoSpheres Sulfate Microspheres</td>
			<td>Invitrogen</td>
			<td>F8858</td>
			<td>4.0 &mu;m, red fluorescent (excitation wavelength 580/emission wavelength 605)</td>
		</tr>
		<tr>
			<td>AZ 9260 photoresist</td>
			<td>AZ Electronic Materials</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>AZ 400 K developer</td>
			<td>AZ Electronic Materials</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Tetramethylammonium hydroxide (TMAH) 25%</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>provided by Virginia Tech cleanroom</td>
		</tr>
		<tr>
			<td>Teflon coating</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>applied using DRIE machine</td>
		</tr>
		<tr>
			<td>Silicon wafer</td>
			<td>University Wafer</td>
			<td>452</td>
			<td>100 mm diameter, 500 &mu;m thickness, one side polished (SSP)</td>
		</tr>
		<tr>
			<td>Deep Reactive Ion Etching (DRIE)</td>
			<td>Alcatrel</td>
			<td>AMS SDE 100</td>
			<td>&nbsp;</td>
		</tr>
	</tbody>
</table>

label-free isolation and enrichment of cells through contactless dielectrophoresis

Dielectrophoresis (DEP) is the phenomenon by which polarized particles in a non-uniform electric field undergo translational motion, and can be used to direct the motion of microparticles in a surface marker-independent manner. Traditionally, DEP devices include planar metallic electrodes patterned in the sample channel. This approach can be expensive and requires a specialized cleanroom environment. Recently, a contact-free approach called contactless dielectrophoresis (cDEP) has been developed. This method utilizes the classic principle of DEP while avoiding direct contact between electrodes and sample by patterning fluidic electrodes and a sample channel from a single polydimethylsiloxane (PDMS) substrate, and has application as a rapid microfluidic strategy designed to sort and enrich microparticles. Unique to this method is that the electric field is generated via fluidic electrode channels containing a highly conductive fluid, which are separated from the sample channel by a thin insulating barrier. Because metal electrodes do not directly contact the sample, electrolysis, electrode delamination, and sample contamination are avoided. Additionally, this enables an inexpensive and simple fabrication process.
cDEP is thus well-suited for manipulating sensitive biological particles. The dielectrophoretic force acting upon the particles depends not only upon spatial gradients of the electric field generated by customizable design of the device geometry, but the intrinsic biophysical properties of the cell. As such, cDEP is a label-free technique that avoids depending upon surface-expressed molecular biomarkers that may be variably expressed within a population, while still allowing characterization, enrichment, and sorting of bioparticles.
Here, we demonstrate the basics of fabrication and experimentation using cDEP. We explain the simple preparation of a cDEP chip using soft lithography techniques. We discuss the experimental procedure for characterizing crossover frequency of a particle or cell, the frequency at which the dielectrophoretic force is zero. Finally, we demonstrate the use of this technique for sorting a mixture of ovarian cancer cells and fluorescing microspheres (beads).

DEP should be observed qualitatively in the sample channel to confirm the device is operational when the electric field is present. One method to test for the presence of DEP is to adjust the frequency to values far from the crossover frequency where strong pDEP and nDEP are predicted (roughly &gt;40 kHz to observe pDEP and &lt;10 kHz to observe nDEP for most mammalian cancer cells in a buffer with conductivity of 100 &mu;S/cm). It should be noted that inert microspheres exhibit pDEP below their crossover frequency and nDEP above their crossover frequency. For the sawtooth feature geometry presented here, pDEP should be seen for cells at the higher test frequency, indicated by the occurrence of pearl chaining and focusing of cells or particles at the sawtooth feature edge of the channel, while at the lower test frequency, nDEP should be apparent as cells are restricted to the region nearer the straight edge of the channel. At these low frequencies, cells may lyse due to electroporation, so the sample may appear to contain fewer cells, and those that are visible may appear enlarged or blurry if using an enzymatically-activated fluorescing dye. Verifying the frequencies at which pDEP and nDEP occur allows the determination of the crossover frequency as described in the protocol. For mammalian cancer cells, such as MOSE-L used here, we observed nDEP at 5 kHz (Figure 5a) and strong pDEP at 60 kHz (Figure 5b). MOSE-L cells had a diameter of 17.7&plusmn;3.3 &mu;m (n = 268 cells) and the beads have a 4 &mu;m nominal diameter. A full analysis of the dielectric properties of MOSE-L cells compared to MOSE cells in earlier stages of progression can be found in the recent work by Salmanzadeh et al.33
If pDEP and nDEP are not observed at these extremes, various problems could be occurring. The sample conductivity may be too high due to contaminants or cell lysing. Insufficient electric field may be generated due to bubbles trapped in the fluid electrode channels, which prevent conduction of current to the portion of the channels bordering the sample channel. Unsteady flow may be caused by a bubble trapped in the syringe or inlet tubing, bubbles resulting from not keeping the device under vacuum for a sufficient time or not quickly filling the device upon removing from vacuum, leakage due to delamination of poorly bonded PDMS to the glass slide, tears in the barrier membrane due to excessive force exerted when filling the channels, or flow rates at the extremes of the pump operating range.
In addition to determining the crossover frequency of cells, this technique can be used to sort a mixture, illustrated here by a mixture of MOSE-L cells and beads. This example takes advantage of the negative dielectrophoresis (nDEP) exhibited by 4 &mu;m beads at frequencies where MOSE-L cells experience positive dielectrophoresis (pDEP). We operated the device at 200 VRMS and 10 kHz (Figure 6a) and observed that both cells and beads were mixed as they experienced nDEP. At 50 kHz we observed movement of MOSE-L cells to the top of the channel, while beads were restricted to the lower portion of the channel, enabling separation of the mixture into two components (Figure 6b).
<img alt="Figure 5" src="/files/ftp_upload/50634/50634fig5.jpg" fo:content-width="6in" fo:src="/files/ftp_upload/50634/50634fig5highres.jpg"/> 
Figure 5. The position of MOSE-L cells is manipulated by changing the frequency of the applied electric field. The photos are taken at the final sawtooth feature in the sample channel and the fluidic electrodes are located at the corners on the right side. They are filled with PBS containing rhodamine B, which fluoresces to visualize the channels. (a) MOSE-L cells experience nDEP at 200 VRMS and 5 kHz. (b) MOSE-L cells experience pearl chaining and pDEP at 200 VRMS and 60 kHz. <a href="https://www.jove.com/files/ftp_upload/50634/50634fig5highres.jpg" target="_blank">Click here to view larger figure</a>.
<img alt="Figure 6" src="/files/ftp_upload/50634/50634fig6.jpg" fo:content-width="6in" fo:src="/files/ftp_upload/50634/50634fig6highres.jpg"/> 
Figure 6. A mixture of MOSE-L cells (green) and 4 &mu;m beads (red) flow through the final sawtooth feature in the sample channel of a low frequency cDEP device. (a) At 200 V and 10 kHz cells and beads are mixed. Arrows point to faintly appearing cells that likely have died due to the low frequency conditions. (b) At 200 V and 50 kHz cells experience pDEP and move to the feature edge of the channel, while beads experience nDEP and remain confined to the region nearer the straight edge. <a href="https://www.jove.com/files/ftp_upload/50634/50634fig6highres.jpg" target="_blank">Click here to view larger figure</a>.

Watch this Scientific Journal Video about Label-free Isolation and Enrichment of Cells Through Contactless Dielectrophoresis at JoVE.com

Label-free Isolation and Enrichment of Cells Through Contactless Dielectrophoresis

Contactless dielectrophoresis (cDEP) achieves sorting and enrichment of particles via their intrinsic dielectric properties. Fluidic electrode channels replace metallic electrodes traditional to DEP, suiting cDEP to non-damaging sterile characterization and sorting of biological particles. We demonstrate how to prepare a cDEP microdevice and conduct cell characterization and sorting experiments.

Dielectrophoresis (DEP) is the phenomenon by which polarized particles in a non-uniform electric field undergo translational motion, and can be used to ...

label-free-isolation-enrichment-cells-through-contactless

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Medicine

15.9K Aufrufe.  Virginia Tech.  Kontaktlose Dielektrophorese (cDEP) erreicht Sortierung und Anreicherung der Partikel über ihren inneren dielektrischen Eigenschaften. Fluidic Elektrodenkanäle ersetzen traditionelle metallische Elektroden, um DEP, passend cDEP auf nicht-sterile beschädigen Charakterisierung und Sortierung von biologischen Partikeln. Wir zeigen, wie man eine cDEP Kleinst vorzubereiten und durchzuführen Zellcharakterisierung und Sortierversuche. 

Serie: Label-free Isolation and Enrichment of Cells Through Contactless Dielectrophoresis

Isolation, Characterization and Comparative Differentiation of Human Dental Pulp Stem Cells Derived from Permanent Teeth by Using Two Different Methods

Developing wisdom teeth are easy-accessible source of stem cells during the adulthood which could be obtained by routine orthodontic treatments. Human pulp-derived stem cells (hDPSCs) possess high proliferation potential with multi-lineage differentiation capacity compare to the ordinary source of adult stem cells1-8; therefore, hDPSCs could be the good candidates for autologous transplantation in tissue engineering and regenerative medicine. Along with these benefits, possessing the mesenchymal stem cells (MSC) features, such as immunolodulatory effect, make hDPSCs more valuable, even in the case of allograft transplantation6,9,10. Therefore, the primary step for using this source of stem cells is to select the best protocol for isolating hDPSCs from pulp tissue. In order to achieve this goal, it is crucial to investigate the effect of various isolation conditions on different cellular behaviors, such as their common surface markers &amp; also their differentiation capacity.
Thus, here we separate human pulp tissue from impacted third molar teeth, and then used both existing protocols based on literature, for isolating hDPSCs,11-13 i.e. enzymatic dissociation of pulp tissue (DPSC-ED) or outgrowth from tissue explants (DPSC-OG). In this regards, we tried to facilitate the isolation methods by using dental diamond disk. Then, these cells characterized in terms of stromal-associated Markers (CD73, CD90, CD105 &amp; CD44), hematopoietic/endothelial Markers (CD34, CD45 &amp; CD11b), perivascular marker, like CD146 and also STRO-1. Afterwards, these two protocols were compared based on the differentiation potency into odontoblasts by both quantitative polymerase chain reaction (QPCR) &amp; Alizarin Red Staining. QPCR were used for the assessment of the expression of the mineralization-related genes (alkaline phosphatase; ALP, matrix extracellular phosphoglycoprotein; MEPE &amp; dentin sialophosphoprotein; DSPP).14

The method described isolation and characterization of human Dental Pulp Stem Cells (hDPSCs) by using either enzymatic dissociation of pulp (DPSC-ED) or direct outgrowth of stem cells from pulp tissue explants (DPSC-OG). Then followed by in vitro comparative differentiation of both types of hDPSCs into odontoblasts.

Developing wisdom teeth are easy-accessible source of stem cells during the adulthood which could be obtained by routine orthodontic treatments. Human ...

Enrichment for Chemoresistant Ovarian Cancer Stem Cells from Human Cell Lines

Cancer stem cells (CSCs) are defined as a subset of slow cycling and undifferentiated cells that divide asymmetrically to generate highly proliferative, invasive, and chemoresistant tumor cells. Therefore, CSCs are an attractive population of cells to target therapeutically. CSCs are predicted to contribute to a number of types of malignancies including those in the blood, brain, lung, gastrointestinal tract, prostate, and ovary. Isolating and enriching a tumor cell population for CSCs will enable researchers to study the properties, genetics, and therapeutic response of CSCs. We generated a protocol that reproducibly enriches for ovarian cancer CSCs from ovarian cancer cell lines (SKOV3 and OVCA429). Cell lines are treated with 20 &#181;M cisplatin for 3 days. Surviving cells are isolated and cultured in a serum-free stem cell media containing cytokines and growth factors. We demonstrate an enrichment of these purified CSCs by analyzing the isolated cells for known stem cell markers Oct4, Nanog, and Prom1 (CD133) and cell surface expression of CD177 and CD133. The CSCs exhibit increased chemoresistance. This method for isolation of CSCs is a useful tool for studying the role of CSCs in chemoresistance and tumor relapse.

Cancer stem cells (CSCs) are&#160;implicated in tumor relapse due to chemoresistance. We have optimized a protocol for selection and expansion of CSCs from ovarian cancer cell lines. By treating cells with the chemotherapeutic cisplatin and culturing&#160;in a stem cell promoting media we enrich for non-adherent CSC cultures.

Cancer stem cells (CSCs) are defined as a subset of slow cycling and undifferentiated cells that divide asymmetrically to generate highly proliferative, ...

Isolation, Cryopreservation and Culture of Human Amnion Epithelial Cells for Clinical Applications

Human amnion epithelial cells (hAECs) derived from term or pre-term amnion membranes have attracted attention from researchers and clinicians as a potential source of cells for regenerative medicine. The reason for this interest is evidence that these cells have highly multipotent differentiation ability, low immunogenicity, and anti-inflammatory functions. These properties have prompted researchers to investigate the potential of hAECs to be used to treat a variety of diseases and disorders in pre-clinical animal studies with much success.
hAECs have found widespread application for the treatment of a range of diseases and disorders. Potential clinical applications of hAECs include the treatment of stroke, multiple sclerosis, liver disease, diabetes and chronic and acute lung diseases. Progressing from pre-clinical animal studies into clinical trials requires a higher standard of quality control and safety for cell therapy products. For safety and quality control considerations, it is preferred that cell isolation protocols use animal product-free reagents. 
We have developed protocols to allow researchers to isolate, cryopreserve and culture hAECs using animal product-free reagents. The advantage of this method is that these cells can be isolated, characterized, cryopreserved and cultured without the risk of delivering potentially harmful animal pathogens to humans, while maintaining suitable cell yields, viabilities and growth potential. For researchers moving from pre-clinical animal studies to clinical trials, these methodologies will greatly accelerate regulatory approval, decrease risks and improve the quality of their therapeutic cell population.

We describe a protocol to isolate and culture human amnion epithelial cells (hAECs) using animal product-free reagents in accordance with current good manufacturing practices (cGMP) guidelines.

Human amnion epithelial cells (hAECs) derived from term or pre-term amnion membranes have attracted attention from researchers and clinicians as a ...

In vitro Enrichment of Ovarian Cancer Tumor-initiating Cells

Evidence suggests that small subpopulations of tumor cells maintain a unique self-renewing and differentiation capacity and may be responsible for tumor initiation and/or relapse. Clarifying the mechanisms by which these tumor-initiating cells (TICs) support tumor formation and progression could lead to the development of clinically favorable therapies. Ovarian cancer is a heterogeneous and highly recurrent disease. Recent studies suggest TICs may play an important role in disease biology. We have identified culture conditions that enrich for TICs from ovarian cancer cell lines. Growing either adherent cells or non-adherent &#8216;floater&#8217; cells in a low attachment plate with serum free media in the presence of growth factors supports the propagation of ovarian cancer TICs with stem cell markers (CD133 and ALDH activity) and increased tumorigenicity without the need to physically separate the TICs from other cell types within the culture. Although the presence of floater cells is not common for all cell lines, this population of cells with innate low adherence may have high tumorigenic potential.Compared to adherent cells grown in the presence of serum, TICs readily form spheres, are significantly more tumorigenic in mice, and express putative stem cell markers. The conditions are easy to establish in a timely manner and can be used to study signaling pathways important for maintaining stem characteristics, and to identify drugs or combinations of drugs targeting TICs. The culture conditions described herein are applicable for a variety of ovarian cancer cells of epithelial origin and will be critical in providing new information about the role of TICs in tumor initiation, progression, and relapse.

In vitro Enrichment of Ovarian Cancer Tumor-initiating Cells

Tumor-initiating cells (TICs) may represent a viable therapeutic target for the treatment of ovarian cancer, a highly recurrent and fatal disease. We present a protocol for culture conditions that enrich for this highly tumorigenic population of cells.

Evidence suggests that small subpopulations of tumor cells maintain a unique self-renewing and differentiation capacity and may be responsible for tumor ...

Isolation of Human Lymphatic Endothelial Cells by Multi-parameter Fluorescence-activated Cell Sorting

Lymphatic system disorders such as primary lymphedema, lymphatic malformations and lymphatic tumors are rare conditions that cause significant morbidity but little is known about their biology. Isolating highly pure human lymphatic endothelial cells (LECs) from diseased and healthy tissue would facilitate studies of the lymphatic endothelium at genetic, molecular and cellular levels. It is anticipated that these investigations may reveal targets for new therapies that may change the clinical management of these conditions. A protocol describing the isolation of human foreskin LECs and lymphatic malformation lymphatic endothelial cells (LM LECs) is presented. To obtain a single cell suspension tissue was minced and enzymatically treated using dispase II and collagenase II. The resulting single cell suspension was then labelled with antibodies to cluster of differentiation (CD) markers CD34, CD31, Vascular Endothelial Growth Factor-3 (VEGFR-3) and PODOPLANIN. Stained viable cells were sorted on a fluorescently activated cell sorter (FACS) to separate the CD34LowCD31PosVEGFR-3PosPODOPLANINPos LM LEC population from other endothelial and non-endothelial cells. The sorted LM LECs were cultured and expanded on fibronectin-coated flasks for further experimental use.

The goal of this protocol is to isolate lymphatic endothelial cells lining human lymphatic malformation cyst-like vessels and foreskins using fluorescence-activated cell sorting (FACS). Subsequent cell culturing and expansion of these cells permits a new level of experimental sophistication for genetic, proteomic, functional and cell differentiation studies.

Lymphatic system disorders such as primary lymphedema, lymphatic malformations and lymphatic tumors are rare conditions that cause significant morbidity ...

Studying Pancreatic Cancer Stem Cell Characteristics for Developing New Treatment Strategies

Pancreatic ductal adenocarcinoma (PDAC) contains a subset of exclusively tumorigenic cancer stem cells (CSCs) which have been shown to drive tumor initiation, metastasis and resistance to radio- and chemotherapy. Here we describe a specific methodology for culturing primary human pancreatic CSCs as tumor spheres in anchorage-independent conditions. Cells are grown in serum-free, non-adherent conditions in order to enrich for CSCs while their more differentiated progenies do not survive and proliferate during the initial phase following seeding of single cells. This assay can be used to estimate the percentage of CSCs present in a population of tumor cells. Both size (which can range from 35 to 250 micrometers) and number of tumor spheres formed represents CSC activity harbored in either bulk populations of cultured cancer cells or freshly harvested and digested tumors 1,2. Using this assay, we recently found that metformin selectively ablates pancreatic CSCs; a finding that was subsequently further corroborated by demonstrating diminished expression of pluripotency-associated genes/surface markers and reduced in vivo tumorigenicity of metformin-treated cells. As the final step for preclinical development we treated mice bearing established tumors with metformin and found significantly prolonged survival. Clinical studies testing the use of metformin in patients with PDAC are currently underway (e.g., NCT01210911, NCT01167738, and NCT01488552). Mechanistically, we found that metformin induces a fatal energy crisis in CSCs by enhancing reactive oxygen species (ROS) production and reducing mitochondrial transmembrane potential. In contrast, non-CSCs were not eliminated by metformin treatment, but rather underwent reversible cell cycle arrest. Therefore, our study serves as a successful example for the potential of in vitro sphere formation as a screening tool to identify compounds that potentially target CSCs, but this technique will require further in vitro and in vivo validation to eliminate false discoveries.

Pancreatic cancer stem cells (CSCs) can be expanded in vitro using the anchorage-independent sphere culture technique, which represents a powerful tool to study CSC biology and can serve as the first step to develop novel CSC-targeting therapies. Here the methodology for expanding, analyzing and targeting of pancreatic CSCs is provided.

Pancreatic ductal adenocarcinoma (PDAC) contains a subset of exclusively tumorigenic cancer stem cells (CSCs) which have been shown to drive tumor ...

Isolation and Propagation of Circulating Tumor Cells from a Mouse Cancer Model

Cancer metastasis is the foremost cause of cancer-associated deaths. Recent studies have shown that circulating tumor cells (CTCs) are important in cancer metastasis. Indeed, the number of CTCs correlates with tumor size. Here, a detailed description is provided of a methodology for isolation and propagation of CTCs from a syngeneic mouse model of hepatocellular carcinoma (HCC) which allows for downstream analysis of potentially important molecular mechanisms of solid organ tumor metastasis. This method is efficient and reproducible. It is a non-invasive technique and, therefore, has potential to replace the invasive biopsy of tissues from humans which may be associated with complications. Therefore, the method discussed here allows for the isolation and propagation of CTCs from whole blood samples such that they can be examined and characterized. This has potential for future adaptation for clinical applications such as diagnosis, and personalized targeted therapy.

Circulating tumor cells (CTCs) have been shown to play an important role in tumor metastasis. Here, a method for the isolation and propagation of CTCs from the whole blood of a syngeneic mouse tumor model of hepatocellular carcinoma (HCC) metastasis is described.

Cancer metastasis is the foremost cause of cancer-associated deaths. Recent studies have shown that circulating tumor cells (CTCs) are important in cancer ...

Isolation and Culture Expansion of Tumor-specific Endothelial Cells

Freshly isolated tumor-specific endothelial cells (TEC) can be used to explore molecular mechanisms of tumor angiogenesis and serve as an in vitro model for developing new angiogenesis inhibitors for cancer. However, long-term in vitro expansion of murine endothelial cells (EC) is challenging due to phenotypic drift in culture (endothelial-to-mesenchymal transition) and contamination with non-EC. This is especially true for TEC which are readily outcompeted by co-purified fibroblasts or tumor cells in culture. Here, a high fidelity isolation method that takes advantage of immunomagnetic enrichment coupled with colony selection and in vitro expansion is described. This approach generates pure EC fractions that are entirely free of contaminating stromal or tumor cells. It is also shown that lineage-traced Cdh5cre:ZsGreenl/s/l reporter mice, used with the protocol described herein, are a valuable tool to verify cell purity as the isolated EC colonies from these mice show durable and brilliant ZsGreen fluorescence in culture.

We report a reliable method to isolate and culture primary tumor-specific endothelial cells from genetically engineered mouse models.

Freshly isolated tumor-specific endothelial cells (TEC) can be used to explore molecular mechanisms of tumor angiogenesis and serve as an in vitro model ...

Isolation and Cellular Phenotyping of Mesenchymal Stem Cells Derived from Synovial Fluid and Bone Marrow of Minipigs

Mesenchymal stem cells (MSCs) have been established after isolation from various tissue sources, including bone marrow and synovial fluid. Recently, synovial-fluid-derived MSCs were reported to have multi-lineage differentiation potential and immunomodulatory features, which indicates that these cells can be used for tissue engineering and systemic treatments. This study presents a protocol for simple and non-invasive isolation of MSCs derived from the bone marrow and synovial fluid of minipigs to analyze cell surface markers for cell phenotyping and in vitro culturing. Using sexually mature six-month-old minipigs, bone marrow was extracted from the iliac crest bone using a bone marrow extractor, and the synovial fluid was aspirated from the femorotibial joint. Procedures for the collection of samples from both sources were non-invasive. The protocols for effective isolation of MSCs from harvested cell sources and for creating in vitro culture conditions to expand stable MSCs from minipigs and the application of systemic autologous treatments are provided. For cell phenotyping, the cell surface markers of both cells were analyzed using flow cytometry. In the results, the MSCs were isolated from the synovial fluid of the minipigs and showed that synovial-fluid-derived MSCs have a similar morphology and cell phenotype to bone-marrow-derived MSCs. Therefore, non-invasively obtained synovial fluid is a valuable source of MSCs.

A protocol establishing mesenchymal stem cells (MSCs) isolated from the synovial fluid and bone marrow of minipigs and non-invasively collected using syringe aspiration is presented. Cellular phenotyping was performed using flow cytometry after cell isolation and in vitro cultivation of bone marrow and synovial fluids derived from MSCs.

Mesenchymal stem cells (MSCs) have been established after isolation from various tissue sources, including bone marrow and synovial fluid. Recently, ...

Isolation and Culture of Primary Endothelial Cells from Canine Arteries and Veins

Cardiovascular disease is studied in both human and veterinary medicine. Endothelial cells have been used extensively as an in vitro model to study vasculogenesis, (tumor) angiogenesis, and atherosclerosis. The current standard for in vitro research on human endothelial cells (ECs) is the use of Human Umbilical Vein Endothelial Cells (HUVECs) and Human Umbilical Artery Endothelial Cells (HUAECs). For canine endothelial research, only one cell line (CnAOEC) is available, which is derived from canine aortic endothelium. Although currently not completely understood, there is a difference between ECs originating from either arteries or veins. For a more direct approach to in vitro functionality studies on ECs, we describe a new method for isolating Canine Primary Endothelial Cells (CaPECs) from a variety of vessels. This technique reduces the chance of contamination with fast-growing cells such as fibroblasts and smooth muscle cells, a problem that is common in standard isolation methods such as flushing the vessel with enzymatic solutions or mincing the vessel prior to digestion of the tissue containing all cells. The technique we describe was optimized for the canine model, but can easily be utilized in other species such as human.

Novel isolation methods of primary endothelial cells from blood vessels are needed. This protocol describes a new technique that completely inverts blood vessels of interest, exposing only the endothelial side to enzymatic digestion. The resulting pure endothelial cell culture can be used to study cardiovascular diseases, disease modelling, and angiogenesis.

Cardiovascular disease is studied in both human and veterinary medicine. Endothelial cells have been used extensively as an in vitro model to study ...