Electrophysiology is defined in the dictionary as 'The study of electric phenomena of living organisms' which isn't really much help to anyone. What an electrophysiologist can measure is a movement of charge, in short an electric current. In biological systems, electric currents are carried by ions therefore our current measurements reflect the movement of ions. All the fun in electrophysiology comes in determining which ion carries which current and across which barrier. Patch-Clamp is the most sophisticated electrophysiological technique available at present. Using the patch-clamp technique we can measure ion movements through individual ion channels or across the whole cell membrane, we can split currents into components carried by individual ions or even stop the movement of a particular ion altogether.
The application of the patch clamp technique has provided so many insights into cellular physiology that its originators, Bert Sakmann and Erwin Neher were awarded the Nobel Prize for Physiology and Medicine in 1991.
Conventional sharp microlectrodes impale the cell in order to measure across the cell membrane. Patch clamp electrodes are much too large to be inserted into a cell, how then can potential and current be measured across the cell membrane? A key innovation of the patch clamp technique is to stick the patch pipette on to the surface of a cell membrane instead of piercing it. If a patch pipette is placed onto the cell surface and gentle suction is applied an Ω shaped bubble of membrane is drawn into the patch pipette. The edges of this patch of membrane adhere tightly or 'seal' to the glass of the patch pipette. The electrical resistance of this seal between pipette glass and membrane is so high (>109 Ω, i.e. a giga ohm seal, or gigaseal) that the small patch of membrane underneath the patch pipette is by comparison a low resistance pathway and thus the favoured route for current flow. This small patch of membrane may be voltage clamped to a series of potentials and the conductance of the patch calculated from the amount of current required to move from one potential to another.
The patch of membrane under the pipette is very small. If the radius of a patch clamp pipette is 1 µm then the area of the patch under the pipette will be about 3 square picometres. In a patch this small, opening or closing of a single ion channel will cause a significant alteration in the overall conductance of the patch. Under most circumstances, ion channels are either open or closed and they switch very rapidly from one state to the other. Thus the effect of a single ion channel opening is to cause an abrupt increase in the conductance of the patch of membrane beneath the pipette which may be 'visualised' using the patch clamp technique as a step-like increase in current.
These data in show a patch clamp record made from a mouse submandibular acinar cell. The rectangular deflections of the trace represent openings of a single ion channel. At a given voltage and ionic environment, the size of the current deflection is directly proportional to the conductance of this channel; the larger the deflection the greater the conductance. If two channels open simultaneously then the current is exactly twice as large. There were at least three channels in this particular patch because there are three distinct current levels. Ion channels may be distinguished from one another on the basis of this characteristic unit conductance, on the duration of each opening (open time) and on the probability of the channel being open (open probability) under a given set of conditions.
It is also possible to remove the patch of membrane from the cell without breaking the gigaseal and thus measure ion channel openings in an isolated patch of membrane. As what was inside the cell is now outside the patch-pipette this configuration is called an isolated inside-out patch. Even more tricky is to get an isolated outside-out patch, but that can be achieved too.
In addition to the single channel recording modes, the patch clamp technique may be applied to measure the currents that result from ion movements across the membrane of the whole cell. This mode of operation is known as the whole cell configuration. The first step in achieving this configuration is to obtain a high resistance contact between the pipette and the cell membrane ('gigaseal'). However, the patch of membrane under the pipette, which was the focus of attention in the single channel experiments is, in whole cell experiments, ruptured by application of a short pulse of negative pressure. Just as in the single channel experiments, the tight seal (GΩ) between pipette glass and cell membrane persists and the low resistance route for current flow is now into the cell and across entire cell surface membrane. A second feature of the whole cell configuration is that, following disruption of the patch of membrane under the pipette, the interior of the patch pipette is continuous with the cell interior. Thus the solution filling the patch pipette will enter into and equilibrate with the cell interior. Small ions equilibrate within seconds of breaking through into the whole cell configuration. In the whole-cell recording mode then, the ionic composition of both the external (bathing) and internal (pipette) solutions may be controlled for the purposes of the experiment.
Ions move across the cell membrane through ion channels down their electrochemical gradients. In whole cell experiments, both the electrical and chemical elements of the driving force may be regulated; the electrical component by means of the patch clamp amplifier and the concentration gradient by adjusting the composition of the intra- and extra- cellular bathing solutions.
Control of the ionic composition of the intracellular bathing solution represents a significant advance over other microelectrode techniques because the currents that give rise to the membrane potential may now be resolved into their component parts.
For example, submandibular (and lacrimal) acinar cells have Ca2+-activated K+ and Cl- channels. With a high K+ intracellular solution and a high Na+ extracellular bathing solution, the reversal potential for K+ may be calculated to be -80 mV. At this potential, there is no chemical or electrical driving force for K+ movement and K+ will not move through K+ channels even if they are open. If any current is measured under these conditions, it cannot therefore, be carried by K+. Similarly, if the Cl- concentration is identical in the intra- and extra- cellular bathing solution, the reversal potential for Cl- will be 0 mV. Any current measured at 0 mV cannot be carried by Cl-. However, at the Cl- reversal potential, there is a large driving force for K+. Thus K+ currents may be measured at the Cl- reversal potential. Chloride currents may be measured at the K+ reversal potential where there is a large driving force for Cl-.
The patch clamp amplifier may be set to alternate rapidly between these two potentials. A double voltage pulse is imposed upon a holding potential of -40 mV. The first voltage step of +40 mV shifts the potential to 0 mV, the second voltage step of -40 mV shifts the potential to -80 mV. Each voltage pulse is 100 ms in duration and the entire sequence is repeated every 500 ms. The current returned at each potential (shown within the box) may be averaged to give a single current value per pulse and a pair of current values for each pulse protocol every 500 ms. These averaged currents may then be plotted against time.
In effect, you can measure K+ and Cl- currents separately and (almost) simultaneously with a time resolution determined by the rate of stimulus. In submandibular and lacrimal cells a time resolution of 4Hz is about the best you can manage because the K+ channel is voltage as well as Ca2+ activated and it may require 20-30 ms to stabilise after the voltage pulse. Nevertheless, changes in Ca2+-dependent ion channel activity has been shown to accurately reflect the changes in [Ca2+]i at the membrane
Localised changes in [Ca2+]i and ion current activation after stimulation by 50µM inositol 1,4,5 trisphosphate which was perfused into the cell via the patch pipette and was present for the duration of the experiment. A: Image maps of [Ca2+]i at 0.5 s intervals immediately after stimulation. B: bright-field image showing a 3 cell cluster and the borders and location of apical (a) and basolateral (b) regions of the fura-2 loaded cell. C & D: numbered arrows indicate time points at which the first and last images in A were sampled. C: average change in [Ca2+]i within the apical (a) and basal (b) regions. D: K+ and Cl- currents. The dashed line indicates the zero current point.
From: A. R. Harmer, P. M. Smith and D. V. Gallacher. Local and global calcium signals and fluid and electrolyte secretion in mouse submandibular acinar cells Am J Physiol Gastrointest Liver Physiol 288:118-124, 2005.