Electrophysiology Fundamentals, Membrane Potential and Electrophysiological Techniques

Electrophysiology is the measurement of the electrical activity, or “excitability”, of biological cells (be they muscle cells, neurons, or stem cells). This can be done at a single-cell level, or it can include simultaneous measurements from hundreds or thousands of cells.

This article focuses on single-cell electrophysiology and introduces the fundamentals of electrophysiological investigation by describing the ionic basis of action potentials and the different electrophysiological techniques used to explore and understand excitable tissues and organs.

Simply put, electrophysiology is the study of the electrical properties of biological circuits within cells, tissues, whole organs and systems.

At the simplest level, any electronic circuit (such as a radio or computer) contains a battery connected to components by copper wires, through which negatively charged electrons flow and do work, obeying Ohm’s law:  voltage (V) = current (I) x resistance (R).

In biology, excitable cells obey the same fundamental law, but the charge is carried by atoms in solution—that is ions (chloride (Cl^-), sodium (Na⁺), potassium (K⁺) and calcium (Ca²⁺)). The flow of these ions is regulated by ion channels—enzymes that catalyze the passive flow of ions down electrochemical gradients (described below). These ion channels are proteins encoded by different genes. The physics is the same as in an electronic circuit, but the biology adds a few layers of complexity.

Since Galvani animated frogs’ legs with metal electrodes in the late 1700s, scientists have worked at the intersection of physics and physiology to pioneer our understanding of what makes cells “excitable”.¹

Advances in electrophysiology theory, techniques and equipment over the last hundred years have vastly amplified our understanding of how the body works, including fundamental knowledge of the heart, brain (and other organs), and improved diagnosis and treatment of disease.

An early insight was the realization that charged ions in solution could move and generate voltage differences, a physical phenomenon explained by the Nernst equation (for a history read Archer, 1989). Then in the early 1950’s, Hodgkin and Huxley revealed the ionic basis of the neuronal action potential,²  instigating an intense period of investigation to understand molecular mechanisms underlying the action potential.³ Technological advancements in electrode design were made in parallel.⁴ In 1981, Sakmann and Neher published their paper on patch-clamp electrophysiology, pioneering the use of glass micropipettes capable of measuring ion flow through single ion channels or across a single cell membrane.⁵ This technology drove an explosion in our understanding of how excitable cells work and enabled scientists to investigate the fundamental mechanisms underlying excitability and brain function⁶ with the help of this

Membrane potential, action potential steps, and the action potential graph

Membrane potential

Membrane potential is the voltage or potential difference across the cell membrane. The potential difference is caused by the hydrophobic membrane separating charges, acting as both a capacitor and resistor to the movement of charged ions across it.

The predominant cellular ions that carry charge are Na⁺, K⁺, Ca²⁺ and Cl^-. These ions have different concentrations inside and outside the cell. So when ion channels open, the ions want to move to balance out their concentrations.

Generally, Na⁺, Ca²⁺ and Cl^- ions exist in greater concentrations extracellularly and K⁺ ions in greater concentrations intracellularly, with the inside of the cell having a net negative charge. This sets up an electrochemical gradient (or “driving force”) so that when an ion channel opens, the permeable ions flow down their electrochemical gradient carrying charge across the membrane and causing a voltage change. This results in either depolarization or hyperpolarization of the cell membrane potential.

The positively charged ions Na⁺ and Ca²⁺ have an electrochemical gradient favoring their movement into the negatively charged cell, so their positive charge depolarizes the electrochemical potential. On the other hand, K⁺ ions are highly concentrated inside the cell, so their driving force is in the opposite direction, out of the cell. As a result, they carry positive charge out of the cell and make the cell membrane potential more negative; this is called hyperpolarization.

Ions cannot freely diffuse through the hydrophobic lipid bilayer cell membrane, but they can move through the channels or pores set within it that extend across the membrane from the inside to the outside. Many of these channels are highly selective for only one species of ion and are often “gated,” opening only in response to a change in membrane voltage (voltage-gated ion channels) or on the binding of a small molecule or ligand (ligand-gated ion channels).

The flow of ions across the membrane sets its overall resistance (according to Ohm’s law). If there are a large number of channels open, the membrane resistance (Rm) will be low, as the more channels are open, the more ions can flow, ergo a greater conductance and smaller resistance are produced.

Another important property of the lipid bilayer is its capacitance (membrane capacitance, Cm), which arises from the physical properties of the bilayer; an insulator immersed in conducting solutions on each side. There is a separation of charge across the membrane associated with the potential difference across the membrane. The net effect is that the ion flow through the ion channels must charge the membrane capacitor first before changing the membrane potential. Thus, a cell with large capacitance and a small conductance will change potential slowly, while a cell with smaller capacitance will change quickly. The speed of this change is often referred to as the membrane time constant, Rm.Cm = Tau. This property of a neuron has a profound impact on the time interval over which synaptic potentials will summate (see below section on graded potentials).

Action potential steps and graph

Excitable cells commonly have resting membrane potentials of around -60 mV (negative inside when measured with respect to the outside of the cell). Skeletal muscle, cardiac muscle and neurons express voltage-gated Na⁺ channels and voltage-gated K⁺ channels.

During an action potential, voltage-gated Na⁺ channels open allowing the rapid influx of Na⁺ ions and the fast depolarization of the cell membrane potential. If this is all that happened, then the potential would go to ~+40 mV and stay there. But, this strong depolarization opens the voltage-gated K⁺ channels through which K⁺ ions flow in the opposite direction and pull the voltage back (repolarize) to around -60 mV. The specific duration and waveform of the action potential are determined by the type and density of the ion channels present in the excitable cell membrane, as shown for neurons and cardiac cells in Figures 1 and 2, respectively.

Figure 1: Neuronal action potential.

Neurons

1) Action potentials must be triggered by a small depolarization of the resting membrane potential (around -60 mV); this might be caused by a synaptic input, for example. When the depolarization reaches “threshold,” this triggers the opening of voltage-gated Na⁺ channels (-55 mV). 2) This causes a rapid influx of Na⁺ ions which depolarize the membrane towards +40 mV. 3) The Na⁺ channels inactivate (a form of closure) within a millisecond or so, and the voltage-gated K⁺ channels open. 4) This efflux of K⁺ ions repolarizes the membrane back to -60 mV or beyond to more negative potentials, causing an afterhyperpolarizing potential. 5) On a much slower timescale, the minuscule concentrations of ions exchanged across the membrane are pumped back by membrane-bound Na⁺/K⁺ exchangers (sometimes called ion pumps) that exchange K⁺ and Na⁺ ions to reset and maintain the resting potential.

Figure 2: Cardiac action potential.

Cardiac cells

1) Rapid Na⁺ influx through voltage-gated Na⁺ channels depolarizes the cell membrane. 2) Voltage-gated K⁺ channels open, beginning repolarization. 3) Voltage-gated Ca²⁺channels open and the influx of Ca²⁺ positive charge is balanced by efflux of K⁺ positive charges, producing a plateau phase. 4) Ca²⁺channels close and K⁺ channels remain open, repolarizing membrane potential to -90 mV.

An action potential is initiated when the cell membrane is depolarized to the threshold for Na⁺ channel activation. In neurons, action potentials are fundamental to neuronal communication.

Graded potentials are sub-threshold changes in the membrane potential in response to stimuli or incoming innervation from other cells. These postsynaptic potentials can be excitatory (depolarizing) or inhibitory (hyperpolarizing) depending on the type of channels that underlie them. The influence the sub-threshold response has on initiating an action potential is determined by many factors, including: the size of the input and its subsequent response, the distance from the site of action potential initiation, and the biophysical properties of the membrane, such as its membrane time constant tau. In practice, graded-potentials sum on top of each other in a process called sub-threshold integration. (Figure 3).

Figure 3: Graded potentials.

Electrophysiologists measure small cellular currents in the order of pA–nA. Therefore, they require sensitive equipment that can exclude vibration and electrical interference (noise) from the ambient surroundings. The standard rig setup required for in vitro and in vivo electrophysiology is pictured below (Figure 4).

Figure 4: Standard electrophysiology rig setup.

1) An air table to prevent physical vibrations adding movement artifacts to the experimental recording.

2) A faraday cage to shield the setup from electrical interference. Removing ambient electrical noise is crucial to obtaining clear, useful electrophysiological recordings.

3) A microscope and micromanipulator(s) to position the microelectrode(s). Depending on the experimental setup, the user might incorporate multiphoton imaging capabilities. For in vivo investigations, the user might opt to remove the microscope to make room for different apparatus, such as treadmills.

4) An amplifier is needed to collect and amplify the acquired signals from your electrodes. This is connected to a digitizer to convert analog signals into digital signals.

5) Data acquisition and analysis software is needed to set up the experiment, design and run protocols and extract meaningful results from the data collected.

In addition, the user may need to consider incorporating drug delivery systems and temperature control devices.

Patch clamp

Patch-clamp electrophysiology is a technique pioneered by Sakmann and Neher in the 1970s and 80s⁵ and for which they shared the Nobel Prize for Physiology and Medicine in 1991. It involves creating a series circuit with a cell or patch of cell membrane without puncturing the cell wall. Instead, a glass micropipette filled with ionic solution and containing a silver/silver-chloride wire attached to a patch-clamp amplifier forms a high resistance gigaohm seal between the patch and the glass in the mouth of the pipette. The different patch-clamp configurations: cell-attached, whole-cell, inside-out and outside-out; are shown in Figure 5 which is adapted from the original paper.⁵ The figure highlights the critical path used to carry out whole-cell in vitro patch-clamp recordings.

The whole-cell configuration is achieved by approaching the cell membrane with a capillary glass microelectrode which has been micro-forge heated and pulled so that the mouth diameter is ~1 µm, with a resistance of between 2 and 6 MΩ, and pipette offset zeroed.

A dimple is observed by filling the micropipette with an internal solution and applying positive pressure (< 2 ml) as the pipette comes into contact with the cell membrane. By commanding a 5 mV step pulse to the electrode, we can use this as a seal test to understand when we have a patch seal with the cell. Release of positive pressure allows the membrane and pipette mouth to become contiguous, forming a loose patch, increasing resistance and reducing the current amplitude observed in the 5 mV seal test. Light sucking advances a “patch” of membrane into the mouth of the pipette, this is known as the cell-attached configuration and generates a high resistance (GΩ) seal between the capillary glass and the membrane patch. This normally reduces the observed seal test current to a flat line with two fast pipette capacitance transients at the start and end of the voltage step. These are removed by circuitry in the patch-clamp amplifier.

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Before going “whole-cell”, the electrode is gently pulled out and up to the surface edge of the cell, removing the risk of the nucleus blocking the patch and increasing access resistance. The cell is held at a negative potential approximately equal to resting membrane potential (-60 mV) and negative pressure is applied to rupture the membrane. The rupture means that the pipette solution is now continuous with the cytoplasm, and the electrical properties of the whole cell are in series with the pipette electrode. This means that the excitability of the whole-cell membrane can now be measured and manipulated. 

Figure 5: How to achieve the different patch-clamp configurations.

In the whole-cell configuration, it is possible to investigate the electrical properties of the cell in current-clamp and voltage-clamp modes.

Current-clamp

In current-clamp mode, the user injects a known current amplitude to the inside of the cell through their setup and observes the change in cellular excitability in response to these current injections. It is a valuable technique because it can mimic physiological scenarios, like a synaptic input. In Figure 6 A, you can see the response of one cell to increasing step current injections from -20 pA to +20 pA in 10 pA steps, such that +10 pA of current injection was sufficient to bring the cell to threshold and fire action potentials.⁷ A summary plot of the mean current injection-firing rate relationship for multiple cells is shown in Figure 6 B. There is a positive relationship between current injection and the firing rate of the cells.

Figure 6: Current clamp investigation of mouse AgRP neurons in a brain slice preparation. Credit: Branco et al. 2016,⁷ reproduced under the Creative Commons Attribution 4.0 International license.

Voltage clamp

In voltage-clamp mode, the membrane potential is clamped at user-specified voltages. The clamp is enabled by the feedback amplifier and headstage, which inject current to hold the cell's membrane potential at different command voltages set by the user.

Ion channels open at the different command voltages, meaning membrane resistance changes as ionic currents flow across the membrane. The feedback amplifier instantaneously compensates for this by injecting the reciprocal current to maintain the cell at the command voltage, giving a read out of this injected current as the membrane current.

Although it is not a physiological measurement of cellular ionic properties, it has proved a very insightful method in investigating the conductances present in cell membranes and those that underlie cellular excitability.² It is especially helpful when combined with pharmacological blockers of different conductances. In Figure 7, neurons in the medial nucleus of the trapezoid body (MNTB) and lateral superior olive (LSO) in the mouse auditory brainstem were investigated by voltage clamp in the presence of Na⁺ channel blockers to isolate K⁺conductances.⁸ The aim of this experiment was to investigate the relative contribution of one such voltage-gated K⁺ channel (K_V3.1) to the total K⁺ conductance by recording currents at different voltage step commands in the absence and presence of a known blocker of K_V3.1 K⁺, tetraethylammonium (TEA). The figure shows that TEA reduced the overall amplitude of the peak outward K⁺ current by ~ 50% at +40 mV in both MNTB and LSO neurons. This suggests that K_V3.1 channels mediate 50% of the cells’ outward K⁺ conductance.

Figure 7: Current-voltage relationship of K⁺ currents measured in cells in the auditory brainstem. Current traces observed in cells from the MNTB (A) and LSO (B). Current-voltage relationships (IV curves) from MNTB (C) and LSO (D) show the amplitude of the peak and sustained current recorded at each command voltage. Voltage command steps are inset in (C). Peak current observed in the absence and presence of TEA (1 mM) in the MNTB (E) and LSO (F). Credit: Choudhury et al. 2020, reproduced under the Creative Commons Attribution 4.0 international license.

This technique involves placing wire electrodes or silicon probes directly into the subject in vivo or layering primary cells, cultured cells or tissue slices over electrodes in vitro.

For in vivo extracellular electrophysiology, the electrodes are generally thin in diameter and slide down into the tissue adjacent to the cells of interest. The advantage of this technique over intracellular electrophysiology is that it can be performed in awake, behaving subjects, providing greater insights to the neuronal activity that underlies behaviors. Electrodes can be arranged as:

• Single electrodes - one electrode, one recording site
• Tetrodes - four electrodes bundled together, enables better cell sorting
• Multielectrode arrays (MEAs) - arrays of multiple recording electrodes, records spiking across a greater area of tissue

The electrodes record the electronic field potentials produced by spiking or “firing” neurons. These waveforms are called “local field potentials” and can be composed of the spiking of multiple cells adjacent to the electrode. The number of neurons that an electrode or silicon probe can “listen” to is dependent on the impedance of the electrode and the number of recording sites (channels) available.

Single unit recording

Researchers must prepare their subjects carefully when performing in vivo electrophysiology experiments (Figure 8). Recordings from brain tissue requires a section of skull to be removed to access the brain. The electrode must be advanced slowly through the tissue at a rate of ~ 1 µm per second, to prevent mechanical stress and to avoid causing a bleed in the tissue. Using a micromanipulator facilitates this. Experimenters know their position in the brain by “zeroing” their micromanipulator’s coordinates when the electrode touches the brain’s surface. They can then compare their position to published brain atlas coordinates, such as the Allen Brain atlas. Once in their area of interest, experimenters can move the probe or electrode with fine control to improve recording fidelity from neurons of interest via the microdrive.

Signals from the electrode are amplified and observed in real time using an oscilloscope. Experimenters also “listen” to cell spiking activity by connecting an audio device that outputs a noise in response to a spike.⁹ Experimenters working with transgenic animal subjects expressing channelrhodopsin in cell types of interest may also combine in vivo electrophysiology with optogenetic stimulation by using a light source to irradiate their neurons of interest.^9,10

Spike sorting algorithms and software enables users to sort the local field potential into individual units, or neurons, for analysis after recording.¹¹ This step is critical as the data recorded during in vivo recordings can be extremely noisy.

Figure 8: In vivo electrophysiological recording set up. The microelectrode is lowered into the tissue using a micromanipulator. The microdrive enables fine positioning of the microelectrode once it is in the correct region. The microscope is used by the operator to enable correct guidance and tracking of the electrode. Signals from the tissue are amplified by the amplifier and activity is tracked on the oscilloscope. The computer enables recording, monitoring, analysis and control of the experiment via digital interface and the amplifier.

Multi-unit recording

Recently, technological advances in electrode design have led to the development of the Neuopixels probe.¹² These probes use complementary metal-oxide semiconductor (CMOS) technology to enable simultaneous recording from thousands of neurons.¹³ This number looks set to increase as advances continue to miniaturize the size of the technology.

The advantage of Neuropixels is the unprecedented resolution afforded by the many recording sites along the shank of the electrode. However, the drawback is that information can only be gleaned from neurons that are adjacent to the single shaft as it projects into the brain tissue. This can be overcome to some extent by combining multiple electrodes in different sites per recording session. But, there is a practical limit to the number of electrodes that can be placed in the brain.

Solutions like the CMOS-hosted in vivo multielectrode system (CHIME) enable recording from hundreds of electrodes spread in a 3D arrangement. The electrodes comprise hundreds of glass-ensheathed microelectrodes that project down from a CMOS amplifier array.¹⁴ These enable monitoring of neuronal activity over a larger spatial area.

Multielectrode arrays (MEA)

Arrays of multiple electrodes in a dish can be used for higher-throughput extracellular recordings from layers of cells in vitro, or from brain slices. MEA systems have been used in preclinical and drug discovery research for over 50 years.¹⁵

The electrodes are arranged on the bottom of the dish in grids like a chessboard. The cells are then cultured on top of them, or the tissue slice is laid on top. The cellular activity is measured extracellularly as local field potentials. By recording from multiple electrodes simultaneously, researchers can investigate network dynamics within the tissue or between cells, as well as gather data from multiple cells at once, increasing the throughput of these experiments.

In terms of MEA design, the number of electrodes per well has increased over the years with the advent of new technologies, such as CMOS arrays. This means the number of electrodes per dish has jumped from 64 to 1000+, improving the resolution scientists have to observe and record cellular activity.

Pre-clinical research

Both in vitro and in vivo electrophysiology is used extensively in pre-clinical and academic research. Use of these technologies has contributed greatly to our understanding of behavioral neuroscience, connectomics, neurophysiology and neuropharmacology, cardiology and toxicology. In addition, electrophysiology continues to be the “ground-truth” assay when measuring neuronal activity.¹⁶

Drug discovery

Medium to high-throughput electrophysiology systems are used for compound screening or toxicological assays in cells. Some systems can automatically “patch” cells, while others use electrode arrays to measure local field potentials from cells. Depending on the application, high-throughput electrophysiology systems can be used to measure cellular contraction, impedance and other assays.¹⁷

Clinical electrophysiology

Clinical electrophysiologists regularly perform tests on patients to assist medical diagnosis and patient monitoring. Such tests include 12 lead electrocardiogram (ECG), electroencephalogram (EEG), nerve conduction tests and auditory testing.¹⁸

About the authors

Dr. Adam Tozer

Adam undertook postgrad training in the Forsythe lab where his research focused on communication between neurons in an area of the brain important for processing the auditory environment. He then moved to the Heisler and subsequently Branco labs at the University of Cambridge, UK, and the MRC Laboratory of Molecular Biology, Cambridge UK, respectively. His postdoctoral research focused on neuronal communication in the hypothalamus, specifically in the neurons that drive feeding behavior. Adam is an avid science communicator and moved away from the lab into full-time science communication and marketing to help promote the great work done in labs across the World.

Prof. Ian Forsythe