Optimized Transfection Strategy for Expression and Electrophysiological Recording of Recombinant Voltage-Gated Ion Channels in HEK-293T Cells

Electrophysiology is commonly used to study the activity of recombinant voltage gated ion channels in cultured human embryonic kidney cells. Successful recording requires that the cells express channels at sufficient levels at the cell membrane for detection and that the cells be plated at a low enough cell density for isolation of individual cells. Performing cell recordings is frequently hampered by the long incubation periods at 28 degrees Celsius needed for such expression where there is typically a loss of cell adhesion and membrane stability.

Here, an optimized method for cell culture and transfection is demonstrated that circumvent these problems. The cells are transfected at a moderate co fluency, then incubated at 28 degrees Celsius until adequate ion channel expression is achieved. The transfected cells are then plated onto glass cover slips at low density and incubated at 37 degrees Celsius for several hours to allow for cell attachment and membrane stabilization.

Once appropriately plated cells are then identified by co-expressed EGFP and whole cell patch clamp. Recording can be performed or the cells can be once again incubated at 28 degrees Celsius. To further increase channel surface expression results from recording experiments on cells transfected with voltage-gated calcium channels such as L CAV one L CAV two, or L cav three indicate that the channels have been successfully transfected and expressed by the cells.

Although only representative results for L CAV three are shown In our laboratory, we have developed a method to enhance the expression of poorly expressing ion channel genes for patch clamp recording in human cell lines. Now the conventional approach is to first split the cells on the cover slips and transfect with plasma CDNAs containing the ion channel genes, and then incubating for many days at 28 degrees Celsius in order to achieve high efficient ion channel expression prior to electrophysiological recording. Our new approach greatly reduces the required incubation time at 28 degrees Celsius prior to recording.

Now by decreasing the 28 degree incubation time, we find that cell membranes are much more stable and the transfected cells adhere much more to the cover slips. This increases the number of recordable cells and significantly makes experimentation more efficient. Prior to transfection split a fully confluent flask of HEK 2 9 3 T cells, one to four into a new 25 centimeter squared flask and place the cells in a 37 degrees Celsius 5%carbon dioxide incubator.

Allow the cells to attach for three to four hours after four hours, examine the cells by differential interference contrast to verify attachment. Note that the cells no longer look round and are flattening. Extensions or pseudopodia are visible.

Less confluence cells would look the same but would be spaced further apart. Once the cells have attached, combine the necessary reagents to perform a standard calcium phosphate transfection. Prepare a total of 12 micrograms of DNA in 600 microliters of transfection solution per flask.

Here CDNAs from various ion channels are cloned into the P-I-R-E-S two EGFP plasmid Transfection of this expression vector into a mammalian cell line. Results in the production of enhanced green fluorescent protein from the same mRNA as the cloned insert through translation initiated at an internal ribosome entry site just downstream of the ion channel coating sequence and upstream of the EGFP coating sequence. This allows positively transfected cells to be identified without possible interference in protein function that may be caused by fusing the EGFP protein directly to the protein of interest.

Pipette the transfection solution over the cells dropwise, then gently swirl the flask and place it in the 37 degrees Celsius incubator overnight the next day. Check the transfection efficiency by viewing the cells under a fluorescence microscope. Positively transfected cells should appear green throughout the entire cell.

Once transfection has been verified, carefully pipette four to five milliliters of warm medium into the inverted flask. Wash the cells by slowly turning the flask over so that the cell surface is down and rocking the flask so that the wash solution gently moves over them. Remove the medium and repeat this wash.

Step two more times. Then add six milliliters of medium to the flask and incubate at 37 degrees Celsius for two hours. Expression of invertebrate ion channels presents some challenges.

First, they're often of large size and may therefore take a long time to translate at the endoplasmic reticulum. Also, they may not be efficiently targeted to the cell membrane in a mammalian cell line. Finally, since different species exhibit different codon usage frequencies, heterologous CDNAs may harbor high frequencies of codons that are uncommon in mammalian cells such as HEK 2 9 3 T to overcome some of these difficulties.

Following the two hour incubation at 37 degrees Celsius after washing transfect, transfer the flasks to a 28 degrees Celsius incubator for two to six days to extend the expression period. The incubation time must be determined empirically for each type of ion channel being expressed. Since incubation at 28 degrees Celsius for prolonged time periods leads to cell detachment and poor membrane stability.

Further cell preparation is needed prior to performing electrophysiological studies. First, remove the media from the transfected cells by aspiration pipette one milliliter of 37 degrees Celsius trypsin solution into an inverted flask. Then gently return the flask to the cell surface down position to allow the trypsin to run gently over the cells.

Turn the flask back over and remove the trypsin. Then add 250 microliters of warm trypsin directly over the cells and incubate at 37 degrees Celsius for three to five minutes until the cells detach. In the meantime, place three to eight poly L lysine coated cover slips into a 60 millimeter culture dish and add five milliliters of warm culture medium.

Once the cells have detached resus, suspend them in four milliliters of warm medium by pipetting up and down about six times next, using the tip of the micro pipette, push down on the cover slips to ensure that they're at the bottom of the dish. Then add 250 to 500 microliters of resuspended cells. Incubate the cells at 37 degrees Celsius for three hours to allow them to adhere to the cover slips.

Leaving the cells for too long at 37 degrees will result in cell division and dilution of the transfected plasmids. The transfected cells should be plated at a low enough density so that they are isolated from one another as shown here. This will facilitate electrophysiological recording.

For cells that require accessory subunits, it is often necessary to transfer the cells to 28 degrees Celsius and incubate them for an additional three to four days before electrophysiological. Recording is carried out for cells that do not require accessory subunits. Recording can be performed immediately following this incubation.

The electrophysiology rig used here includes an amplifier, motorized, dual pipette manipulators, and epi fluorescence microscope, and perfusion systems. The equipment has been mounted on a vibration isolation table and stray. Electrical noise is limited by a 40 inch tall type two ferrett cage Electronic control systems including a PC equipped with a digit data 1440, a analog to digital converter interface in conjunction with P clamp 10.1 software.

The computer monitor amplifier, motorized manipulator, and a valve link profusion control system are all mounted outside the Faraday cage. On a freestanding 19 inch metal electrical rack, all electrical equipment is grounded to a copper distributor and plugged into a medical grade isolation transformer, which provides line insulation, a consistent ground and surge suppression. Prepare for experiments by mounting fire polished micro pipettes with a resistance of two to five mega ohms on the micro pipette holder on the head stage.

The patch pipettes are filled with internal solution flick to remove air bubbles and finally mounted on the head stage. Next, using forceps transfer one cover slip with plated cells to a 35 millimeter plastic Petri dish containing three milliliters of external recording solution at room temperature. The volume of recording solution must be measured accurately when performing drug experiments during which known amounts of drug are added to the dish with a micro pipette, place the dish on the microscope stage.

Then fill a micro electrode holder half cell with three molar cesium chloride. Then insert the reference electrode into the half cell and place the ground electrode into the bath using the microm manipulator, lower the patch pipette into the bath solution. Visualize the cells in fluorescence mode and center an isolated positively transfected adherent green cell in the middle of the field of view.

While in bath mode, bring the patch pipette into close proximity of the cell. Zero the square wave trace observed on the computer screen. Adjust the position of the patch pipette so that it is just touching the cell and the square current trace on the screen begins to fluctuate.

Apply gentle suction with an attached syringe until a giga ohm seal is formed. Once a giga ohm seal is achieved, switch the program to patch mode. Apply a quick pulse of suction with the syringe in order to break through the cell membrane.

Then switch to cell mode and use whole cell parameters on the amplifier. To compensate for capacitive transient, it is necessary to equilibrate the patch so that the intracellular recording solution dialyze into the patched cell. Input the voltage clamp protocols written for this experiment here.

The experimenter has designed a protocol to measure ion currents through voltage gated ion channels that result when the voltage across the cell membrane is modulated. In this case, we are looking at an IV protocol that specifies that the software hold the cell at minus 110 millivolts and then SEP to minus 70 millivolts for 250 milliseconds. Then back down to minus 110 millivolts holding potential each time the step increases by 10 millivolts so that the steps are to minus 70 millivolts, minus 60 millivolts, et cetera, until the last step is to plus 20 millivolts.

Filter the recorded currents at 10 kilohertz using a Lowpass Bessel filter on the amplifier and digitize at a sampling frequency of two kilohertz using the clamp X 10.1 software use only recordings with minimal leak less than 10%For analysis, HEK 2 93 T cells were cultured and transfected with L Cav. Three calcium channels as shown here. Successful transfection of heterologous ion channel CDNAs was indicated by fluorescence generated from the P-I-R-E-S two eeg FP vector.

Here we show successful expression of an invertebrate cav three voltage calcium channel. When cells are not transfected, no calcium currents can be elicited. Whereas transfected cells generate large currents when whole cell voltage clamp protocols are applied to the cells.

Experimenters can use various electrophysiological conditions and data analysis regimens depending on their requirements. While attempting this procedure, it is important to remember to use your own judgment regarding the timing of experiments. Since different channels expressed with different efficiencies at the cell membrane, it's up to you to determine the optimal strategy to use to increase the viability of the cells as well as maximize the surface expression of the channels you're interested in.

After watching this video, we hope that you have gained a good understanding of how to culture and maintain a human cell line, as well as how to transfect heterologous ion channel CDNAs and subunit CDNAs into these cells. For electrophysiological recording.