Electroporation temporarily increases cell membrane permeability by means of high-voltage electric pulses. There are several rapidly-developing clinical treatments based on electroporation that target muscle and neuronal cells. It is thus crucial to investigate how electroporation affects the ability of these excitable cells to trigger action potentials.
In-vitro experiments are very important to understand the effects of electroporation on excitable cells. However, primary myocytes and neurons have a very complex ion-channel expression profile that makes it extremely challenging to disentangle the interplay between excitation and electroporation. We developed a protocol to monitor electroporation-induced changes in action potential generation using tet-on spiking HEK cells.
These cells express a minimum complement of sodium and potassium channels that make them excitable. Therefore, this cell simplified the experiments and the theoretical analysis of the obtained results. Using our protocol, we found that very mild electroporation can already dramatically affect cell acceptability.
These results will pave the way for developing theoretical models of electroporation and cell acceptability, and this will help the community to improve the rapidly emerging electroporation-based therapies in the heart, the brain, and skeletal muscles. To begin, pipette one milliliter of PBS into one well of the two-well cover-glass chamber slides. Add 10 microliters of sterile, one-milligram-per-milliliter poly-L-lysine solution, and mix well with a pipette.
Incubate for 1.5 hours at room temperature under sterile conditions in a laminar flow hood, then remove the PBS with poly-L-lysine without any further washing. To seed the cells, prepare a one-to-10 dilution of doxycycline stock solution in a culture medium in a 1.5-milliliter tube. Now, remove the culture medium from the T-25 flask.
Trypsinize the cells with 2.5 milliliters of trypsin EDTA solution in a carbon-dioxide incubator at 37 degrees Celsius for two minutes. Add 2.5 milliliters of fresh culture medium and gently pipette the cells to detach them from the surface. Calculate the volume of the trypsinized cell suspension needed to establish the culture.
Subtract the previously-calculated cell suspension volume from 1, 470 microliters to calculate the volume of the culture medium. To prepare samples with excitable S HEK cells, pipette the calculated volume of the culture medium and trypsinized cell suspension into one well. Add 30 microliters of the doxycycline dilution.
Mix well by gentle pipette-ing just before placing the chambers into the carbon dioxide incubator. For preparing samples with non-excitable NS HEK cells, pipette the calculated volume of culture medium into a well, and add 90%of the calculated volume of cell suspension for S HEK cells without doxycycline. Incubate the chambers in a 5%carbon-dioxide incubator at 37 degrees Celsius for two to three days prior to the experiment.
Before the experiment, pipette three microliters of potentiometric dye stock solution into a 1.5-milliliter tube. Leave the open tube at room temperature in a laminar flow hood for approximately 15 minutes to allow the ethanol to evaporate and the potentiometric dye to dry, then add 997 microliters of cold culture medium at four degrees Celsius to the tube to dissolve the dye to a final concentration of 12 micromolar. Next, remove the culture medium from the well.
Add one milliliter of potentiometric dye solution to the cells. Refrigerate the cells at four degrees Celsius and incubate them for 20 minutes. Afterward, remove the potentiometric dye solution and save it in a tube for further reuse for up to six sequential experiments in one day.
Gently wash the cells three times with tyrode solution. After the last wash, pipette one milliliter of low-potassium tyrode solution into the well. To begin, prepare the HEK cells for electroporation, then configure synchronization of the pulse delivery with image acquisition using a TTL signal from the microscope system that triggers the pulse generator.
At the bottom of the chamber, place two parallel platinum-iridium wire electrodes with a diameter of 0.8 millimeters and a five-millimeter distance between the inner edges. Fix the chamber under the microscope, then connect the electrodes to the pulse generator. Using a voltage and a current probe, connect the output of the pulse generator to an oscilloscope to enable verification of the correct pulse delivery during the experiment.
Focus the cells on the bright field and image the cells located in the middle between the electrodes. Acquire 80 images in time-lapse mode at a rate of at least 25 frames per second. Illuminate the cells with an excitation wavelength of 635 nanometers, set the exposure time to 10 milliseconds, and detect the emission at approximately 700 nanometers.
After the acquisition, wait for two minutes. Adjust the voltage on the pulse generator interface to 63 volts. Record a time-lapse in the same manner every two minutes.
During these acquisitions, apply a single 100-microsecond pulse at the time of acquisition of the 10th image. Acquire images with pulse voltages of 63, 75, 88, 100, 125, 150, 175, and 200 volts. Estimate the electric field to which the cells are exposed as the applied voltage-to-electrode-distance ratio.
For data analysis, export the time-lapse recordings to TIFF format, then place all time-lapse recordings of the applied voltages from an individual experiment into a single folder. Rename each time-lapse recording so that MATLAB can recognize the electric field. For the acquisition at the beginning of the experiment, where no pulse is applied, rename it using the phrase 000Vcm.
Next, open the application downloaded from GitHub. Click on Select data folder"to choose the folder containing the time-lapse recordings of one experiment. To select the pixels, use two sliders to determine the low threshold for a clear image of the membranes and the high threshold for eliminating debris from the analysis.
Now, click on Analyze data. In the Analysis"tab, a table with extracted parameters will appear. On the right side of the table, graphs of relative fluorescence over time for each electric field used will be displayed.
Select the graphs for a given electric field from the dropdown menu on the left side above the table. Examine all the graphs and perform manual correction in case of false peak detection. select Clear AUTO peaks"at the bottom of the Analysis"tab to clear the automatically-detected peaks.
Click on the graph to manually determine a new peak time point. If necessary, select Rerun AUTO peak detection"at the bottom of the Analysis"tab to perform automatic peak detection again. The extracted parameters will now be adjusted to the new peak.
Afterward, select Export Data"at the bottom-right of the Analysis"tab to export data to the same folder as a dotmap file, a spreadsheet file, and PNG images for each electric field used. Electric pulses of 126 volts per centimeter triggered multiple fluorescence peaks in excitable cells, as shown by the distinct oscillations in the fluorescence change over time. With increasing electric field strength, the response changed and exhibited sustained depolarization at 400 volts per centimeter.
Non-excitable cells exhibited no significant peaks in response to the lowest electric-field strengths, indicating no excitation. At higher electric-field strengths, the non-excitable cells also exhibited sustained depolarization, indicating electroporation. The maximum number of peaks in excitable cells was observed at 150 volts per centimeter, with a decreasing trend in the number of peaks as the voltage increased beyond this point.
