生物物理学

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.

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.

The Substantia Nigra pars compacta (SNc) is a midbrain dopaminergic nucleus that plays a key role in modulating motor and cognitive functions. It is crucially involved in several disorders, particularly Parkinson’s disease, which is characterized by a progressive loss of SNc dopaminergic cells. Electrophysiological studies on SNc neurons are of paramount importance to understand the role of dopaminergic transmission in health and disease. Here, we provide an extensive protocol to prepare SNc-containing mouse brain slices and record the electrical activity of dopaminergic cells. We describe all the necessary steps, including mouse transcardiac perfusion, brain extraction, slice cutting, and patch-clamp recordings.

The whole-cell patch-clamp method is a gold standard for single-cell analysis of electrical activity, cellular morphology, and gene expression. Prior to our discovery that patch-clamp pipettes could be cleaned and reused, experimental throughput and automation were limited by the need to replace pipettes manually after each experiment. This article presents an optimized protocol for pipette cleaning, which enables it to be performed quickly (< 30 s), resulting in a high yield of whole-cell recording success rate (> 90%) for over 100 reuses of a single pipette. For most patch-clamp experiments (< 30 whole-cell recordings per day), this method enables a single pipette to be used for an entire day of experiments. In addition, we describe easily implementable hardware and software as well as troubleshooting tips to help other labs implement this method in their own experiments. Pipette cleaning enables patch-clamp experiments to be performed with higher throughput, whether manually or in an automated fashion, by eliminating the tedious and skillful task of replacing pipettes. From our experience with numerous electrophysiology laboratories, pipette cleaning can be integrated into existing patch-clamp setups in approximately one day using the hardware and software described in this article.

Graphic abstract:

Rapid enzymatic cleaning for reuse of patch-clamp pipettes

Characterization of an electrically active cell, such as a neuron, demands measurement of its electrical properties. Due to differences in gene activation, location, innervation patterns, and functions, the millions of neurons in the mammalian brain are tremendously diverse in their membrane characteristics and abilities to generate action potentials. These features can be measured with a patch-clamp technique in whole-cell current-clamp configuration followed by detailed post-hoc analysis of firing patterns. This analysis can be time-consuming, and different laboratories have their own methods to perform it, either manually or with custom-written scripts. Here, we describe in detail a protocol for firing-pattern registration in neurons of the ventral tegmental area (VTA) as an example and introduce a software for its fast and convenient analysis. With the help of this article, other research groups can easily apply this method and generate unified types of data that are comparable between brain regions and various studies.

Graphic abstract:

Workflow of the Protocol

Cardiac pacemaker cells of the sino-atrial node are responsible for the initiation of the heart beat and express an array of ion channels. The patch-clamp technique is the gold standard method for investigating the function of ion channels expressed in electrically active cells. Conventional whole-cell and perforated patch-clamp techniques can be used to investigate ionic currents in the voltage-clamp mode and changes in membrane potential (e.g., action potential) in the current-clamp mode. Here, we provide details of protocols used to measure spontaneous and triggered action potentials and whole-cell funny current If (HCN4) in single cardiomyocytes isolated from the mouse sino-atrial node (SAN).

In multi-cellular organisms, cells communicate with each other utilizing chemical messengers. For many of these messenger molecules, the membrane is an insurmountable barrier. Yet, they act by binding to surface proteins often triggering a cascade of reactions inside the cell. Accordingly, studying ligand-receptor interactions at the cellular surface is key to understanding important aspects of membrane biology. However, despite a multitude of approaches to study membrane features, there is a need for developing techniques that can measure ligand binding with high temporal resolution and on a single cellular level. We recently developed a label-free approach to study ligand binding in real time. This methodology capitalizes on changes of the membrane’s surface potential induced by the adsorption of a charged ligand. The resulting apparent alteration of membrane capacitance is measurable by capacitance recordings. Herein, we describe the implementation of the same using recordings obtained from HEK293 cells stably expressing the human serotonin transporter (SERT), which were challenged with the inhibitor cocaine.