The overall goal of this experiment is to determine chloride selectivity of new light-gated ion channels. This method can answer key questions in the biophysics of channelrhodopsins such as photocurrent amplitudes, kinetics and ion selectivity. The main advantage of this technique is the direct readout of biophysical channel activity that is fast and easy to replicate.
Generally individuals new to this method will struggle because each step of the experimental procedure from the preparation of HEK cells to the recording of photocurrents has to be optimized. Seeding and transection of the HEK cells will be demonstrated by Altina Klein. To begin this procedure, weigh the solid components for 100 milliliters of the intracellular and 500 milliliters of the extracellular buffered solutions in separate beakers.
And then add the respective amount of the prepared stock solutions. Next add ultrapure water and carefully adjust the pH using acids and bases. Then adjust the osmolarity with glucose to 290 milliosmoles and 320 milliosmoles for the intracellular and extracellular solutions, respectively.
In this procedure place up to three lysine-coated glass cover slips side-by-side in a 35-millimeter Petri dish. Then seed 1.5 times 10 to the fifth HEK cells in two milliliters of standard DMEM supplemented with 10%FBS and one micromolar retinol in each dish. To transfect the cells for each dish, prepare a solution of two micrograms of plasmid DNA in 250 microliters of DMEM without FBS and six microliters of transfection reagent.
After 15 minutes of incubation, gently add the solution to the cells. Before measuring, pull some low resistance patch pipettes using a micropipette puller and fire polish them. Store the pipettes in a dust-free container for the day of measurement.
30 minutes prior to recordings, turn on all the setup components to heat up to the working temperature. Ensure that the buffers are at room temperature. To begin, seal the measuring chamber with silicon to prevent leakage of the external buffer.
Then place one coverslip into the chamber and close it. Fill the chamber carefully with extracellular buffer to prevent the cells from detaching. Store the Petri dish with other cover slips in an incubator for the next set of experiments.
Then place the measuring chamber under the microscope using the 40X objective for cell visualization. Next put an Auger bridge over the bath electrode and place it into the chamber together with the fluid level sensor and the perfusion outlet of the bath handler. Exchange the extracellular solution twice with one milliliter of fresh external solution to remove the residual culture medium and the detached cells.
Bring the cells into focus and search for a transfected cell which needs to be isolated from other cells. Then use a triple band filter set and orange light to excite and visualize mCherry. Next, fill one of the pulled pipettes with the intracellular solution.
And remove the air bubbles by flipping the pipette several times. Afterward mount the pipette on the pipette holder and apply some positive pressure to prevent tip clogging. Locate the tip of the patch pipette under the microscope and navigate it close to the cell using the micromanipulator.
Within the data acquisition software start the membrane test in Bath mode and apply a voltage step. Check if the pipette resistance is in the desired range of 1.3 to 3.0 megaohms. Then zero the offset currents and adjust the pipette potential by turning the pipette offset knob on the amplifier.
To establish a patch in the whole-cell configuration, slowly approach the cell with the patch pipette from the top and release the positive pressure right before touching the cell. Sometimes the cell-attached configuration forms itself after that. After approaching the cell, change the membrane test from Bath mode to Patch mode.
Compensate the pipette capacitance by turning the pipette capacitance compensation knob to get a flat response of the test pulse. Switch the membrane test to Cell mode. Then rupture the patch without destroying the seal by applying short pulses of negative pressure or negative pressure with increasing strength to obtain whole-cell confirmation.
Start series resistance compensation by setting the two whole-cell parameters, cell capacitance and series resistance. In the series resistance compensation panel turn on prediction and correction up to 90%Then turn the whole-cell compensation switch on and adjust the whole-cell parameters in an iterative way to obtain an ideally flat test seal response. After establishing whole-cell configuration, wait at least two minutes before recording to ensure sufficient replacement of intracellular solution through the patch pipette.
Record light-induced currents at different holding potentials. Next exchange the extracellular buffer to low chloride in the chamber at least five times without destroying the patch and record another current voltage relation at the new chloride concentration. This figure shows that during the illumination with green light, PsACR1 features a fast transient current which rapidly decays to a stationary current level.
After the light is switched off, photocurrents decay to zero within milliseconds. Reducing the extracellular chloride concentration completely abolishes outward directed photo currents. Exchanging the extracellular chloride concentration causes a shift of the reversal potential that can be deduced from a current voltage plot.
The evaluation of reversal potentials from multiple measurements quantifies the dramatic reversal potential shift caused by the variation of external chloride concentration. Strikingly, the measured reversal potentials directly correspond to the calculated Nernst potentials for chloride which confirms the high chloride selectivity of PsACR1. After watching this video, you should have a good understanding of how to perform electrophysiological recordings of light-induced photocurrents on HEK cells expressing channelrhodopsin.
Once mastered, the whole procedure can be done in four days while the recording of photocurrents take several hours. Complementing this procedure, site-directed mutagenesis and spectroscopic methods can be performed to further elucidate structural features and cellular mechanisms of channelrhodopsins. The characterization and molecular engineering of channelrhodopsin paved the way for researchers in neuroscience to manipulate the excitability of selected neurons by light.
