Label-free Isolation and Enrichment of Cells Through Contactless Dielectrophoresis

The overall goal of this procedure is to induce translational motion of cells suspended in a pressure driven flow by using dye electrophoresis without contacting electrodes to the sample. This is accomplished by first suspending the cells of interest in a low conductivity buffer. The second step is to load the microfluidic device with the cell suspension and introduce highly conductive fluid into the electrode channels.

Next, the prepared device is connected to high voltage electronics and the frequency response of the cell motion is recorded by video fed from a camera attached to an inverted microscope. The final step is to analyze the video images to quantitatively predict electrical properties of the cells. Ultimately, contactless die electrophoresis is used to show that cells with different electrical properties can be sorted in a label-free non-damaging manner, using inexpensive and simple to fabricate microfluidic devices.

So this method is useful for applications such as sorting live cells from dead, isolating tumor, initiating cells, separating out cancer cells based on stage and looking at the effects of of drugs on different cells in their dielectric properties. The main advantages of this technique over existing methods such as fluorescence activated cell sorting, is that cells can be sorted and characterized based on their intrinsic biophysical properties without the need for labeling based on biomarkers and also that sample sterility can be maintained by eliminating contact between the electrodes and the sample. Well, we first had this idea when we wanted to develop a technique that avoided several of the issues such as sample contamination, electrolysis expensive fabrication, and that's associated with traditional di electrophoretic devices.

For isolating mammalian cells. Follow previously reported procedures using photoresist and deep reactive ion etching to etch the desired channel design into a silicon wafer, depositing a thin layer of Teflon to improve the release of the micro device from the wafer. Wrap the wafer with aluminum foil to prevent spillover.

Mix polymethyl alane or PDMS in a 10 to one ratio of elastomer to curing agent DGAs for approximately 20 minutes and pour onto the wafer. Then heat the wafer for 45 minutes at 100 degrees Celsius to avoid damaging the Teflon coating of the stamp after cooling. Remove the aluminum foil.

Trim the PDMS using a surgical blade and punch access holes at the channel inlet outlet. Using a blunt puncher suited to the size of the tubing here, a 1.5 millimeter blunt puncher is used. Next, clean a glass microscope slide as directed in the text protocol.

Clean the PDMS device using Scotch Magic tape. Examine under a microscope to be sure channels are clean. Then expose the slide and PDMS device to air plasma for two minutes firmly press the channel side of the device to the glass slide, avoiding formation of air bubbles between the device and the slide.

Prepare a low conductivity buffer as described in the text protocol, which will be referred to as DEP buffer. Suspend the cells in DEP buffer at 3 million cells per milliliter here cancerous, late stage mouse, ovarian surface epithelial, or moss L cells are used after dying the cells using a fluorescent membrane. Permeable dye measure the conductivity of the dy cell suspension.

The conductivity should be approximately 100 microsiemens per centimeter. If the conductivity measurement is too high, spin the sample down. Resus suspend in depth buffer, and repeat the conductivity measurement.

The most difficult aspect of this protocol is avoiding bubbles in the electrode channels and the sample channel bubbles in the electrode channels can prevent electro hole connections and bubbles in the sample channel can prevent steady flow. Be sure to perform the demonstrated steps to avoid bubbles and ensure steady flow Before loading the C dip microfluidic chip. Place it under vacuum for 30 minutes.

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 six inch length of tubing is required. Fit the tubing to the needle tip on the syringe.

Then estimate the required length for outlet tubing by dividing the target collected sample volume by the cross-sectional area of the tube. Cut the required length of tubing. Next, draw the cell suspension into a syringe.

Check that no bubbles are located in the tubing or syringe. Upon removing the chip from the 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 insulating barrier to prime the electrode channels quickly place empty pipette tips in one outlet of each fluidic electrode channel pipette 200 microliters of PBS containing a small amount of bromine B.And gently tap to introduce the fluid into the opposite end of the electrode channel. Maintain gentle pressure until PBS with bromine B visibly progresses up the tip of the empty pipette tip, repeat for each electrode. Channel R domine B is used only to fluorescently.

Visualize the fluidic electrode channels after priming. Fill each pipette tip reservoir with PBS, with R domine 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. Collect the target volume. Position the chip on the stage of an inverted microscope equipped with the digital camera.

Mount the syringe on the syringe pump and insert the wire electrodes into the fluidic electrode channel reservoirs pump the fluid through at one milliliter per hour 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. Then reduce the flow rate to five microliters per hour.

For safety. Test the electronics by powering on the function generator, high voltage amplifier and oscilloscope, and adjusting to the desired voltage and frequency. Here these parameters are 200 volts and 20 kilohertz.

Upon verification of acceptable performance, shut the power off. Then connect the electrodes to the high voltage amplifier. Upon achieving steady flow, apply the desired voltage at the desired frequency In this experiment, voltage is 200 volts RMS at frequencies between five and 60 kilohertz.

Record the video files of the cell response with image processing techniques 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, which is the distribution of the cells without applying any electrical field.

After setting a new frequency wait, the amount of time required for cells to flow from the inlet to the monitored location and then begin recording, proceed to perform the image processing for characterizing crossover frequency as described in the text protocol, suspend the desired mixture in DEP buffer at a total concentration of 3 million particles per milliliter. Here this mixture is moss L cells with four micrometer fluorescent beads in DEP buffer perform the previously described steps. To prepare a CEP device, operate the experiment at a frequency between the two determined crossover frequencies of the target cells and the background particles so that the two populations of particles experience DEP forces in opposite directions.

To collect the contents of assorted sample, remove the outlet tubing upon collection of 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. For mammalian cancer cells, such as the most L cells used here, negative DEP was observed at five kilohertz and strong positive DEP.

At 60 kilohertz MOS L cells have a diameter of 17.7 plus or minus 3.3 micrometers, and the beads have a four micrometer nominal diameter. In addition to determining the crossover frequency of cells, this technique can be used to sort a mixture illustrated here by a mixture of MO'S L cells and beads. This example takes advantage of the negative dielectrophoresis exhibited by four micrometer beads at frequencies where MO'S L cells experience positive dielectrophoresis.

When the device was operated at 200 volts RMS and 10 kilohertz, both cells shown here in green and beads shown here in red were mixed as they experienced negative DEP at 50 kilohertz. Movement of MOS L cells to the top of the channel was observed while beads were restricted to the lower portion of the channel, enabling separation of the mixture into two components. So following this procedure, other techniques can be conducted downstream, such as cell culturing and immunofluorescence imaging or biochemical assays to to answer additional questions such as how the dielectric properties correlate to their physical characteristics of the cells.

So after its development, this technique really paved the way for biologists to study how metabolite induced changes in the cytoskeleton of the cell can be correlated to their dielectric properties using a mouse ovarian cancer model. After watching this video, you should have a good understanding of how to enrich and sort cells using contactless electrophoresis.