Biophysics

Epithelial tissues form barriers to the flow of ions, nutrients, waste products, bacteria, and viruses. The conventional electrophysiology measurement of transepithelial resistance (TEER/TER) can quantify epithelial barrier integrity, but does not capture all the electrical behavior of the tissue or provide insight into membrane-specific properties. Electrochemical impedance spectroscopy, in addition to measurement of TER, enables measurement of transepithelial capacitance (TEC) and a ratio of electrical time constants for the tissue, which we term the membrane ratio. This protocol describes how to perform galvanostatic electrochemical impedance spectroscopy on epithelia using commercially available cell culture inserts and chambers, detailing the apparatus, electrical signal, fitting technique, and error quantification. The measurement can be performed in under 1 min on commercially available cell culture inserts and electrophysiology chambers using instrumentation capable of galvanostatic sinusoidal signal processing (4 μA amplitude, 2 Hz to 50 kHz). All fits to the model have less than 10 Ω mean absolute error, revealing repeatable values distinct for each cell type. On representative retinal pigment (n = 3) and bronchiolar epithelial samples (n = 4), TER measurements were 500–667 Ω·cm² and 955–1,034 Ω·cm² (within the expected range), TEC measurements were 3.65–4.10 μF/cm² and 1.07–1.10 μF/cm², and membrane ratio measurements were 18–22 and 1.9–2.2, respectively.

Changes in neuronal conduction are common in disease states affecting peripheral nerves. These alterations can significantly impact nerve function and lead to sensorimotor disabilities. In vivo electromyography recording is a well-established electrophysiological method that has been used for decades to assess sensory and motor functions in the nervous system. Nerve studies are challenging to conduct in vivo in rodents, and the involvement of muscle activity makes it difficult to isolate and assess nerve function independently. This protocol provides a comprehensive guide for accurate ex vivo sciatic nerve dissection and handling from mice. It includes the creation of a three-compartment chamber and the establishment of electrophysiological protocols, which enable differential recordings and the analysis of compound action potentials from various nerve fibers. This setup allows researchers to study the specific effects of drugs and pathologies on nerves from a mechanistic perspective. The setup is a stand-alone apparatus that does not require the use of suction electrodes and the maintenance of negative pressure, which can affect the signal-to-noise ratio and recording stability.

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.

The planar lipid bilayer (PLB) technique represents a highly effective method for the study of membrane protein properties in a controlled environment. The PLB method was employed to investigate the role of mitochondrial inner membrane protein 17 (MPV17), whose mutations are associated with a hepatocerebral form of mitochondrial DNA depletion syndrome (MDS). This protocol presents a comprehensive, step-by-step guide to the assembly and utilization of a PLB system. The procedure comprises the formation of a lipid bilayer over an aperture, the reconstitution of the target protein, and the utilization of electrophysiological recording techniques to monitor channel activity. Furthermore, recommendations are provided for optimizing experimental conditions and overcoming common challenges encountered in PLB experiments. Overall, this protocol highlights the versatility of the PLB technique in advancing our understanding of membrane protein function and its broad application in various fields of research.

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.

Understanding dendritic excitability is essential for a complete and precise characterization of neurons’ input-output relationships. Theoretical and experimental work demonstrates that the electrotonic and nonlinear properties of dendrites can alter the amplitude (e.g., through amplification) and latency of synaptic inputs as viewed in the axosomatic region where spike timing is determined. The gold-standard technique to study dendritic excitability is using dual-patch recordings with a high-resistance electrode used to patch a piece of distal dendrite in addition to a somatic patch electrode. However, this approach is often impractical when distal dendrites are too fine to patch. Therefore, we developed a technique that utilizes the expression of Channelrhodopsin-2 (ChR2) to study dendritic excitability in acute brain slices through the combination of a somatic patch electrode and optogenetic activation. The protocol describes how to prepare acute slices from mice that express ChR2 in specific cell types, and how to use two modes of light stimulation: proximal (which activates the soma and proximal dendrites in a ~100 µm diameter surrounding the soma) with the use of a high-magnification objective and full-field stimulation through a low-magnification objective (which activates the entire somato-dendritic field of the neuron). We use this technique in conjunction with various stimulation protocols to estimate model-based spectral components of dendritic filtering and the impact of dendrites on phase response curves, peri-stimulus time histograms, and entrainment of pacemaking neurons. This technique provides a novel use of optogenetics to study intrinsic dendritic excitability through the use of standard patch-clamp slice physiology.

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.