The study of ion channel electrophysiology requires the measurement of ion currents across a single cell membrane or many sets of ion channels. This can be carried out using the patch clamp technique.
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History of electrophysiology and the patch clamp technique
The study of electrophysiology began in the 18th century with Luigi Aloisio Galvani’s pioneering experimental discovery. The legs of a dead frog could be induced to twitch when a small electric shock was applied to a nerve. Galvani concluded that there was intrinsic electricity in animals that governed nerve conduction.
Du Bois-Reymond Bernstein took this further, producing an electrode capable of detecting the ionic currents in the frog’s legs. In the 19th century, Alan Hodgkin and Andrew Huxley implicated the role of ion channels in nerve conductance through their invention of the ‘voltage clamp’ technique, which earned them a Nobel Prize in 1963. This technique was limited to large cells, however, due to the large size of electrodes.
It was later refined by Erwin Neher and Bert Sakmann who subsequently won the 1991 Nobel Prize in Physiology or Medicine for their ‘patch clamp’ technique, capable of measuring single-channel conductance using microelectrodes. Clamping refers to the control of voltage at a value determined by the experimenter.
The patch clamp has been further revised with the invention of the gigaseal technique in 1980. This has improved the quality of measurements. The study of electrophysiology has expanded out of biophysical laboratories and is now routinely used in biochemical research.
Quantitative analysis of the ion channel activity at single-channel resolution has applications that range from measuring native channels expressed endogenously to recombinant ion channels expressed in non-native environments.
Principles of the patch clamp technique
This technique is applied to excitable cells, so-called due to their ability to generate currents in response to stimulation. Due to their small size and rapid ionic conductance (afforded by selective ion channels), the current generated is tiny, in the order of 10-9 amperes.
To detect this nanoscale current, a blunt-ended glass or quartz pipette filled with an electrolyte solution is attached via suction to the membrane, forming a seal.
The seal generates a high resistance – in the giga-ohm range- thus earning this seal the term gigaseal. This is termed the ‘cell-attached’ configuration, of which there are many variants. The seal serves to isolate a patch of the membrane from the surrounding membrane, allowing only the ionic movements across this section to flow into the pipette.
The current generated by the ions flowing into the pipette can be recorded by an electrode, which is connected to a differential amplifier. A second electrode is placed in the solution immersing the cell as a reference or ‘ground’ electrode, which is used to define the zero level.
The electrode generates a negative feedback mechanism. This is a compensatory current that is similar to the current being recorded. The effect of this is to stop any alterations in the membrane potential from occurring.
The membrane potential measured is compared to the reference electrode value, the command potential. A current is applied which is equal in size to that of the membrane. This provides a way of measuring the current through the membrane.
Configurations of the Patch Clamp Technique
The four ‘cell attached’ configurations possible for patch-clamp techniques are as follows:
Cell-attached mode
The pipette is placed close to the cell membrane, and suction is applied to generate a seal between the pipette and the membrane. Single channel currents can be recorded. This is the standard configuration. However, it is limited by the inability to alter the concentrations of ions on either side of the membrane.
Whole-cell mode
A pipette is used to apply a suction force strong enough to rupture the cell membrane. Consequently, the pipette has access to the cytoplasm. There is a larger opening generated at the tip of the electrode pipette, which generates less resistance, and therefore better access to the interior of the cell.
The main drawback is the electrode pipette solution eventually replaces the content of the cell due to its larger volume – this results in a dialysis-like effect in which the important factors in the cytoplasm are washed out.
Inside-out mode
The membrane patch is excised from the cell and immersed in the surrounding bath solution. In this configuration, the seal remains intact whilst the rest of the cell is ruptured. Since the ‘inside’ face of the membrane is no longer facing the cytoplasm, but rather the bathing solution, the patch is referred to as ‘inside-out’.
The inside-out technique allows the investigation into how cytosolic factors affect ion channels. It is limited by the potential loss of factors crucial to regulating components present in the cytoplasm
Perforated-cell method
The pipette perforates, rather than ruptures the membrane. Pore-forming agents are used to perforate the membrane. These are usually antibiotics, which allow small ions to permeate, whilst excluding larger ions.
As a result, this configuration maintains the integrity of the cell interior, and so offers advantages over the other configurations, which includes minimizing disruption to the cell interior and allowing recordings over a longer period.
The perforated-cell method takes much longer due to the time required for perforation, and has fewer applications as drugs cannot be applied through the pipette.
Applications of the patch clamp technique
Patch clamp techniques have historically been applied to investigate neurophysiology. Over the past 20 years, however, patch clamp measurements have been applied to medical research – particularly to diseases that are attributed to ion channel malfunction. In pharmacology, it is used as a screening tool for drug development.
Further still, it can be coupled with live cell imaging techniques, such as calcium imaging. Calcium is a notable ion in the body as it is implicated in many signaling events in the brain. Understanding how the brain controls levels of calcium provides a diagnostic insight into brain function.
In this set-up, a calcium-sensitive fluorescent dye is applied to the brain cell through the pipette sealed to the membrane. In this way, both the changes in membrane current and fluorescence can be recorded, which enables visualization of a single-cell calcium signal.
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