Biophysics

Voltage clamp fluorometry (VCF) is a powerful technique in which the voltage of a cell’s membrane is clamped to control voltage-sensitive membrane proteins while simultaneously measuring fluorescent signals from a protein of interest. By combining fluorescence measurements with electrophysiology, VCF provides real-time measurement of a protein’s motions, which gives insight into its function. This protocol describes the use of VCF to study a membrane protein, the voltage-sensing phosphatase (VSP). VSP is a 3 and 5 phosphatidylinositol phosphate (PIP) phosphatase coupled to a voltage sensing domain (VSD). The VSD of VSP is homologous to the VSD of ion channels, with four transmembrane helices (S1–S4). The S4 contains the gating charge arginine residues that sense the membrane’s electric field. Membrane depolarization moves the S4 into a state that activates the cytosolic phosphatase domain. To monitor the movement of S4, the environmentally sensitive fluorophore tetramethylrhodamine-6-maleimide (TMRM) is attached extracellularly to the S3-S4 loop. Using VCF, the resulting fluorescence signals from the S4 movement measure the kinetics of activation and repolarization, as well as the voltage dependence of the VSD. This protocol details the steps to express VSP in Xenopus laevis oocytes and then acquire and analyze the resulting VCF data. VCF is advantageous as it provides voltage control of VSP in a native membrane while quantitatively assessing the functional properties of the VSD.

Gap junctions are transmembrane protein channels that enable the exchange of small molecules such as ions, second messengers, and metabolites between adjacent cells. Gap junctions are found in various mammalian organs, including skin, endothelium, liver, pancreas, muscle, and central nervous system (CNS). In the CNS, they mediate coupling between neural cells including glial cells, and the resulting panglial networks are vital for brain homeostasis. Tracers of sufficiently small molecular mass can diffuse across gap junctions and are used to visualize the extent of cell-to-cell coupling in situ by delivering them to a single cell through sharp electrodes or patch-clamp micropipettes. Here, we describe a protocol for pre-labeling and identification of astrocytes in acute mouse forebrain slices using Sulforhodamine 101 (SR101). Fluorescent cells can then be targeted for whole-cell patch-clamp, which allows for further confirmation of astroglial identity by assessing their electrophysiological properties, as well as for passive dialysis with a tracer such as biocytin. Slices can then be subjected to chemical fixation and immunostaining to detect dye-coupled networks. This protocol provides a method for the identification of astrocytes in live tissue through SR101 labeling. Alternatively, transgenic reporter mice can also be used to identify astrocytes. While we illustrate the use of this protocol for the study of glial networks in the mouse brain, the general principles are applicable to other species, tissues, and cell types.

During neuronal synaptic transmission, the exocytotic release of neurotransmitters from synaptic vesicles in the presynaptic neuron evokes a change in conductance for one or more types of ligand-gated ion channels in the postsynaptic neuron. The standard method of investigation uses electrophysiological recordings of the postsynaptic response. However, electrophysiological recordings can directly quantify the presynaptic release of neurotransmitters with high temporal resolution by measuring the membrane capacitance before and after exocytosis, as fusion of the membrane of presynaptic vesicles with the plasma membrane increases the total capacitance. While the standard technique for capacitance measurement assumes that the presynaptic cell is unbranched and can be represented as a simple resistance-capacitance (RC) circuit, neuronal exocytosis typically occurs at a distance from the soma. Even in such cases, however, it can be possible to detect a depolarization-evoked increase in capacitance. Here, we provide a detailed, step-by-step protocol that describes how "Sine + DC" (direct current) capacitance measurements can quantify the exocytotic release of neurotransmitters from AII amacrine cells in rat retinal slices. The AII is an important inhibitory interneuron of the mammalian retina that plays an important role in integrating rod and cone pathway signals. AII amacrines release glycine from their presynaptic dendrites, and capacitance measurements have been important for understanding the release properties of these dendrites. When the goal is to directly quantify the presynaptic release, there is currently no other competing method available. This protocol includes procedures for measuring depolarization-evoked exocytosis, using both standard square-wave pulses, arbitrary stimulus waveforms, and synaptic input.

Despite playing diverse physiological roles, the area surrounding the central canal, lamina X, remains one of the least studied spinal cord regions. Technical challenges and limitations of the commonly used experimental approaches are the main difficulties that hamper lamina X research. In the current protocol, we describe a reliable method for functional investigation of lamina X neurons that requires neither time-consuming slicing nor sophisticated in vivo experiments. Our approach relies on ex vivo hemisected spinal cord preparation that preserves the rostrocaudal and mediolateral spinal architecture as well as the dorsal roots, and infrared LED oblique illumination for visually guided patch clamp in thick blocks of tissue. When coupled with electric stimulation of the spared dorsal roots, electrophysiological recordings provide information on primary afferent inputs to lamina X neurons from myelinated and non-myelinated fibers and allow estimating primary afferent–driven presynaptic inhibition. Overall, we describe a simple, time-efficient, inexpensive, and versatile approach for lamina X research.

Measuring the action potential (AP) propagation velocity in axons is critical for understanding neuronal computation. This protocol describes the measurement of propagation velocity using a combination of somatic whole cell and axonal loose patch recordings in brain slice preparations. The axons of neurons filled with fluorescent dye via somatic whole-cell pipette can be targeted under direct optical control using the fluorophore-filled pipette. The propagation delays between the soma and 5–7 axonal locations can be obtained by analyzing the ensemble averages of 500–600 sweeps of somatic APs aligned at times of maximal rate-of-rise (dV/dtmax) and axonal action currents from these locations. By plotting the propagation delays against the distance, the location of the AP initiation zone becomes evident as the site exhibiting the greatest delay relative to the soma. Performing linear fitting of the delays obtained from sites both proximal and distal from the trigger zone allows the determination of the velocities of AP backward and forward propagation, respectively.

Key features

• Ultra-thin axons in cortical slices are targeted under direct optical control using the SBFI-filled pipette.

• Dual somatic whole cell and axonal loose patch recordings from 5–7 axonal locations.

• Ensemble averaging of 500–600 sweeps of somatic APs and axonal action currents.

• Plotting the propagation delays against the distance enables the determination of the trigger zone's position and velocities of AP backward and forward propagation.

Intracellular signaling pathways directly and indirectly regulate neuronal activity. In cellular electrophysiological measurements with sharp electrodes or whole-cell patch clamp recordings, there is a great risk that these signaling pathways are disturbed, significantly altering the electrophysiological properties of the measured neurons. Perforated-patch clamp recordings circumvent this issue, allowing long-term electrophysiological recordings with minimized impairment of the intracellular milieu. Based on previous studies, we describe a superstition-free protocol that can be used to routinely perform perforated patch clamp recordings for current and voltage measurements.

Synapses provide the main route of signal transduction within neuronal networks. Many factors regulate critical synaptic functions. These include presynaptic calcium channels, triggering neurotransmitter release, and postsynaptic ionotropic receptors, mediating excitatory and inhibitory postsynaptic potentials. The key features of synaptic transmission and plasticity can be studied in primary cultured hippocampal neurons. Here, we describe a protocol for the preparation and electrophysiological analysis of paired hippocampal neurons. This model system allows the selective genetic manipulation of one neuron in a simple neuronal network formed by only two hippocampal neurons. Bi-directionally analyzing synaptic transmission and short-term synaptic plasticity allows the analysis of both pre- and postsynaptic effects on synaptic transmission. For example, with one single paired network synaptic responses induced by both, a wild-type neuron and a genetically modified neuron can be directly compared. Ultimately, this protocol allows experimental modulation and hence investigation of synaptic mechanisms and thereby improves previously developed methods of studying synaptic transmission and plasticity in ex vivo cultured neurons.

Key features

• Preparation of ex vivo paired cultured hippocampal neurons.

• Bi-directional electrophysiological recordings of synaptic transmission and plasticity.

• Genetic modulation of synaptic network formation (demonstrated by presynaptic viral overexpression of the auxiliary calcium channel α₂δ-2 subunit).

Graphical overview

Spiral ganglion neurons (SGN) are the primary neuronal pathway for transmitting sensory information from the inner ear to the brainstem. Recent studies have revealed significant biophysical and molecular diversity indicating that auditory neurons are comprised of sub-groups whose intrinsic properties contribute to their diverse functions. Previous approaches for studying the intrinsic biophysical properties of spiral ganglion neurons relied on patch-clamp and molecular analysis of cultured somata that were disconnected from their pre-synaptic hair cell partners. In the absence of the information provided by cell-to-cell connectivity, such studies could not associate biophysical diversity with functional sub-groups. Here we describe a protocol for preparing, recording, and labeling spiral ganglion neurons in a semi-intact ex-vivo preparation. In these preparations, the cell bodies of spiral ganglion neurons remain connected to their hair cell partners. The recordings are completed within 4 hours of euthanasia, alleviating concerns about whether long culture times and culture conditions change the intrinsic properties of neurons.

Prokaryotic ion channels have been instrumental in furthering our understanding of many fundamental aspects of ion channels’ structure and function. However, characterizing the biophysical properties of a prokaryotic ion channel in a native membrane system using patch-clamp electrophysiology is technically challenging. Patch-clamp is regarded as a gold standard technique to study ion channel properties in both native and heterologous expression systems. The presence of a cell wall and the small size of bacterial cells makes it impossible to directly patch clamp using microelectrodes. Here, we describe a method for the preparation of giant E. coli spheroplasts in order to investigate the electrophysiological properties of bacterial cell membranes. Spheroplasts are formed by first inhibiting bacterial cell wall synthesis, followed by enzymatic digestion of the outer cell wall in the presence of a permeabilizing agent. This protocol can be used to characterize the function of any heterologous ion channels or ion transporters expressed in E. coli membranes.

PC-1 and PC-2 form an ion channel complex called the polycystin complex, which predominantly localizes to a small hair-like organelle called the primary cilium. The polycystin complex permeates cations, K⁺, Na⁺, and Ca²⁺, and has an unusual 1:3 stoichiometry that combines one PC-1 subunit with three PC-2 subunits. However, the small size and shape of primary cilia impose technical challenges to study the polycystin complex in its native environment. In this paper, we describe the methodology to directly record ion channel activity in primary cilia. This method will allow a detailed functional characterization of how mutations within the polycystin complex cause Autosomal Dominant Polycystic Kidney Disease (ADPKD), essential to develop novel therapeutics for this ciliopathy.