Automated Intracellular Pharmacological Electrophysiology for Ligand-Gated Ionotropic Receptor and Pharmacology Screening.

Communication between neuronal cells, which is central to brain function, is performed by several classes of ligand-gated ionotropic receptors. The gold-standard technique for measuring rapid receptor response to agonist is manual patch-clamp electrophysiology, capable of the highest temporal resolution of any current electrophysiology technique. We report an automated high-precision patch-clamp system that substantially improves the throughput of these time-consuming pharmacological experiments. The patcherBotPharma enables recording from cells expressing receptors of interest and manipulation of them to enable millisecond solution exchange to activate ligand-gated ionotropic receptors. The solution-handling control allows for autonomous pharmacological concentration-response experimentation on adherent cells, lifted cells, or excised outside-out patches. The system can perform typical ligand-gated ionotropic receptor experimentation protocols autonomously, possessing a high success rate in completing experiments and up to a 10-fold reduction in research effort over the duration of the experiment. Using it, we could rapidly replicate previous data sets, reducing the time it took to produce an eight-point concentration-response curve of the effect of propofol on GABA type A receptor deactivation from likely weeks of recording to ∼13 hours of recording. On average, the rate of data collection of the patcherBotPharma was a data point every 2.1 minutes that the operator spent interacting with the patcherBotPharma The patcherBotPharma provides the ability to conduct complex and comprehensive experimentation that yields data sets not normally within reach of conventional systems that rely on constant human control. This technical advance can contribute to accelerating the examination of the complex function of ion channels and the pharmacological agents that act on them. SIGNIFICANCE STATEMENT: This work presents an automated intracellular pharmacological electrophysiology robot, patcherBotPharma, that substantially improves throughput and reduces human time requirement in pharmacological patch-clamp experiments. The robotic system includes millisecond fluid exchange handling and can perform highly efficient ligand-gated ionotropic receptor experiments. The patcherBotPharma is built using a conventional patch-clamp rig, and the technical advances shown in this work greatly accelerate the ability to conduct high-fidelity pharmacological electrophysiology.

Promises and Pitfalls of Parasite Patch-clamp.

Protozoan parasites acquire essential ions, nutrients, and other solutes from their insect and vertebrate hosts by transmembrane uptake. For intracellular stages, these solutes must cross additional membranous barriers. At each step, ion channels and transporters mediate not only this uptake but also the removal of waste products. These transport proteins are best isolated and studied with patch-clamp, but these methods remain accessible to only a few parasitologists due to specialized instrumentation and the required training in both theory and practice. Here, we provide an overview of patch-clamp, describing the advantages and limitations of the technology and highlighting issues that may lead to incorrect conclusions. We aim to help non-experts understand and critically assess patch-clamp data in basic research studies.

Advanced real-time recordings of neuronal activity with tailored patch pipettes, diamond multi-electrode arrays and electrochromic voltage-sensitive dyes.

To understand the working principles of the nervous system is key to figure out its electrical activity and how this activity spreads along the neuronal network. It is therefore crucial to develop advanced techniques aimed to record in real time the electrical activity, from compartments of single neurons to populations of neurons, to understand how higher functions emerge from coordinated activity. To record from single neurons, a technique will be presented to fabricate patch pipettes able to seal on any membrane with a single glass type and whose shanks can be widened as desired. This dramatically reduces access resistance during whole-cell recording allowing fast intracellular and, if required, extracellular perfusion. To simultaneously record from many neurons, biocompatible probes will be described employing multi-electrodes made with novel technologies, based on diamond substrates. These probes also allow to synchronously record exocytosis and neuronal excitability and to stimulate neurons. Finally, to achieve even higher spatial resolution, it will be shown how voltage imaging, employing fast voltage-sensitive dyes and two-photon microscopy, is able to sample voltage oscillations in the brain spatially resolved and voltage changes in dendrites of single neurons at millisecond and micrometre resolution in awake animals.

Single Ion-Channel Analysis in Droplet Interface Bilayer.

Droplet interface bilayer (DIB) is a method of fabricating lipid bilayer membrane by contacting two aqueous droplets coated with a monolayer of lipid molecules in oil media. Lipids coat the droplet surface either by vesicles fusing to the water-oil interface from the droplet side or diffusing toward the interface from the oil side, thereby forming a lipid monolayer. With the DIB technique, nanoliter amounts of aqueous solution is needed and one may obtain two different compositions of monolayers to form asymmetric bilayer which is difficult to replicate by other in vitro lipid membrane methods. Here, a DIB-based protocol is reported to fabricate a stable lipid bilayer membrane to perform single-channel electrophysiology on a pore-forming toxin.

Continuous and Rapid Solution Exchange in a Lipid Bilayer Perfusion System Based on Droplet-Interface Bilayer.

Because of the high sensitivity of lipid bilayers to external pressure fluctuations, a major challenge in functional studies of biological pores or ion channels is the difficulty in exchanging solutions rapidly while maintaining the stability of the lipid bilayer in a model membrane. Here we describe a droplet-interface bilayer-based perfusion system that has been routinely used in our research and is currently the most robust and stable perfusion system that provides prompt solution exchange surrounding a lipid bilayer. In this model membrane system, solutions can be completely exchanged within 1-2 s to obtain prompt responses of a lipid bilayer or membrane pores to the membrane environments. Also, our system is stable enough to sustain continuous perfusions up to at least dozens of minutes. To demonstrate, we show that acidification-induced protein channel insertion, substrate binding to protein channels, and pH gradient-driven protein translocation of anthrax toxin can be sequentially initiated by continuous perfusions in our system. Moreover, by rapidly switching the solutions, the protein translocation based on ratchet mechanisms can be paused and reinitiated iteratively in our system. Overall, this perfusion system provides a controllable and reliable solution exchange platform for investigations of pores and translocations on lipid bilayers.

An Introduction to Patch Clamp Recording.

Electrophysiology is an essential tool aiding the study of the functions and dysfunctions of electrically excitable cells and their networks. The patch clamp method is a refined electrophysiological technique that can directly measure the membrane potential and/or the amount of current passing across the cell membrane. The patch clamp technique is also incredibly versatile and can be used in a variety of different configurations to study a range of properties, from spontaneous cell firing activity in native tissue to the activation and/or deactivation kinetics of individual channels expressed in recombinant cell lines. In this chapter we give an overview of patch clamping and how the different configurations can be set up and applied to electrophysiological research.

Patch Clamp Technology in the Twenty-First Century.

In the almost four decades since its inception, the patch clamp technique has transitioned from a specialist skill to a method commonly used among many others in a lab. Development of patch clamp instrumentation has not been steady: A boost of product releases in rapid succession by multiple manufacturers in the 1990s had slowed to a trickle by the mid-2000s. In 2016, Sutter Instrument's entry into the market of turnkey patch clamp amplifier systems, defined as an amplifier with matching data acquisition hardware and software, caused a fresh breeze in a field in danger of going stale. Sutter has meanwhile completed the product line, culminating in the flagship dPatch® Ultra-fast, Low-noise Digital Amplifier. The dPatch System constitutes a contemporary, digital design that features many firsts, including digital signal compensation, an extremely high bandwidth and fully integrated dynamic clamp capability, paired with the increasingly popular SutterPatch® Software.This chapter compares feature sets of the new Sutter instrumentation with the established platforms by the other two providers of turnkey systems, Axon Instruments by Molecular Devices and HEKA Elektronik by Harvard Bioscience. A variety of products from other manufacturers, who rely on combination with components from other sources rather than offering turnkey systems, are listed, but for their conceptual diversity not compared at a great level of detail. The chapter further covers architectural considerations for patch clamp systems, headstage design, data acquisition strategies and efficient structuring of the recorded data, controlling and monitoring periphery, advanced technologies, such as software lock-in amplifier capability and dynamic clamp features, and application modules for efficient analysis of action potentials and postsynaptic events.

Perforated Whole-Cell Recordings in Automated Patch Clamp Electrophysiology.

The automated patch clamp (APC) technology is used for increasing the data throughput of electrophysiological measurements, especially in safety pharmacology and drug discovery. Typically, electrical access to the cells are obtained using standard whole-cell formation by rupturing the membrane, thereby causing a rapid washout of cytosolic components. In contrast the perforated whole-cell configuration provides electrical access to the cell interior while limiting intracellular wash-out. This method allows for recordings of ion channels that are gated by intracellular modulators (e.g., ATP, cyclic nucleotides, or Ca2+), prevents channel current "run down," and maintains a physiological membrane potential for action potential recordings. Here we present some practical approaches to the use of perforated patch clamp for APC recordings. Our findings from these high-throughput, data-rich measurements (e.g., defining optimized concentrations and practical recommendations for four different perforating agents) can be more broadly applied to perforated patch clamp experiments in general (automated and manual), improving success rates, experimental conditions, and applications.

Utilising Automated Electrophysiological Platforms in Epilepsy Research.

Genetic mutations have long been implicated in epilepsy, particularly in genes that encode ion channels and neurotransmitter receptors. Among some of those identified are voltage-gated sodium, potassium and calcium channels, and ligand-gated gamma-aminobutyric acid (GABA), neuronal nicotinic acetylcholine (CHRN), and glutamate receptors, making them key therapeutic targets. In this chapter we discuss the use of automated electrophysiological technologies to examine the impact of gene defects in two potassium channels associated with different epilepsy syndromes. The hKCNC1 gene encodes the voltage-gated potassium channel hKV3.1, and mutations in this gene cause progressive myoclonus epilepsy (PME) and ataxia due to a potassium channel mutation (MEAK). The hKCNT1 gene encodes the weakly voltage-dependent sodium-activated potassium channel hKCNT1, and mutations in this gene cause a wide spectrum of seizure disorders, including severe autosomal dominant sleep-related hypermotor epilepsy (ADSHE) and epilepsy of infancy with migrating focal seizures (EIMFS), both conditions associated with drug-resistance. Importantly, both of these potassium channels play vital roles in regulating neuronal excitability. Since its discovery in the late nineteen seventies, the patch-clamp technique has been regarded as the bench-mark technology for exploring ion channel characteristics. In more recent times, innovations in automated patch-clamp technologies, of which there are many, are enabling the study of ion channels with much greater productivity that manual systems are capable of. Here we describe aspects of Nanion NPC-16 Patchliner, examining the effects of temperature on stably and transiently transfected mammalian cells, the latter of which for most automated systems on the market is quite challenging. Remarkable breakthroughs in the development of other automated electrophysiological technologies, such as multielectrode arrays that support extracellular signal recordings, provide additional features to examine network activity in the area of ion channel research, particularly epilepsy. Both of these automated technologies enable the acquisition of consistent, robust, and reproducible data. Numerous systems have been developed with very similar capabilities, however, not all the systems on the market are adapted to work with primary cells, particularly neurons that can be problematic. This chapter also showcases methods that demonstrate the versatility of Nanion NPC-16 Patchliner and the Multi Channel Systems (MCS) multielectrode array (MEA) assay for acutely dissociated murine primary cortical neurons, enabling the study of potassium channel mutations implicated in severe refractory epilepsies.

Combining Whole-Cell Patch-Clamp Recordings with Single-Cell RNA Sequencing.

To understand how the brain functions we need to understand the properties of its constituent cells. Whole-cell patch-clamp recordings of neurons have enabled studies of their intrinsic electrical properties as well as their synaptic connectivity within neural circuits. Recent technological advances have now made it possible to combine this with a sampling of their transcriptional profile. Here we provide a detailed description how to combine whole-cell patch-clamp recordings of neurons in brain slices followed by extraction of their cytoplasm suitable for single-cell RNA sequencing and analysis.

Nucleated, Outside-Out, Somatic, Macropatch Recordings in Native Neurons.

Patch-clamp recordings are a powerful tool for the live measurement of the plasma membrane biophysical properties, with the ability to discriminate fast events such as fast inactivating Na+ currents (<1 ms c.a.). It can be used in virtually every cell-type, including cardiomyocytes, skeletal muscles, neurons, and even epithelial cells and fibroblasts. Voltage-clamp, patch-clamp recordings can be used to measure and characterize the pharmacological and biophysical profile of membrane conductances, including leak, voltage-gated, and ligand-gated ion channels. This technique is particularly useful in studies carried out in cell-lines transfected with the gene expressing the conductance under investigation. However, voltage-clamp measures conducted on the soma of a native, adult neuron, for example in an acute brain slice or in the brain of a live individual, are subject to three major limitations: (1) the branching structure of the neuron causes space-clamp errors, (2) ion channels are differentially expressed across different neuronal compartments (such as soma, dendrites, and axons), and (3) the complex geometry of neurons makes it challenging to calculate current densities. While not preventing the experimenter to conduct patch-clamp, voltage-clamp recordings in native neurons, these limitations make the measures poorly standardized and hence often unusable for testing specific hypotheses.To overcome the limitations outlined above, outside-out, patch-clamp recordings can be carried out instead (See Chap. 1, Sect. 3.5); however, the signal-to-noise ratio in outside-outs from native, adult neurons is usually too low for obtaining accurate measurements.Here we describe how to carry out nucleated, outside-out, somatic, macropatch recordings (from now on abbreviated into "macropatch recordings") to obtain accurate and standardized measures of the biophysical and pharmacological properties of somatic, neuronal membrane conductances.

Single or Double Patch-Clamp Recordings In Ex Vivo Slice Preparation: Functional Connectivity, Synapse Dynamics, and Optogenetics.

Patch-clamp recordings are the method of choice to define cell-type specific electrophysiological properties of single neurons and the synaptic connectivity between pairs of connected neurons in brain slices. In combination with optogenetic tools, patch-clamp recordings allow for the investigation of long-range afferent connectivity from identified distant brain areas. Here we describe the necessary equipment to carry out patch clamp recordings, surgical methods for dissection and preparation of horizontal brain slices containing the hippocampus, and a step-by-step guide for establishing patch clamp recordings in the whole-cell configuration. We provide protocols for single neuron stimulation via the patch pipette and for photostimulation experiments that activate axon terminals expressing light sensitive ion channels.

Electrophysiological Approaches to Studying the Suprachiasmatic Nucleus.

In mammals, the part of the nervous system responsible for most circadian behavior can be localized to a bilaterally paired structure in the hypothalamus known as the suprachiasmatic nucleus (SCN). Understanding the mammalian circadian system will require a detailed multilevel analysis of neural SCN circuits ex vivo and in vivo. Many of the techniques and approaches that are used for the analysis of the circuitry driving circadian oscillations in the SCN are similar to those employed in other brain regions. There is, however, one fundamental difference that needs to be taken into consideration, that is, the physiological, cell, and molecular properties of SCN neurons vary with the time of day. In this chapter, we will consider the preparations and electrophysiological techniques that we have used to analyze the SCN circuit focusing on the acute brain slice and intact, freely moving animal.

In vivo patch-clamp recordings reveal distinct subthreshold signatures and threshold dynamics of midbrain dopamine neurons.

The in vivo firing patterns of ventral midbrain dopamine neurons are controlled by afferent and intrinsic activity to generate sensory cue and prediction error signals that are essential for reward-based learning. Given the absence of in vivo intracellular recordings during the last three decades, the subthreshold membrane potential events that cause changes in dopamine neuron firing patterns remain unknown. To address this, we established in vivo whole-cell recordings and obtained over 100 spontaneously active, immunocytochemically-defined midbrain dopamine neurons in isoflurane-anaesthetized adult mice. We identified a repertoire of subthreshold membrane potential signatures associated with distinct in vivo firing patterns. Dopamine neuron activity in vivo deviated from single-spike pacemaking by phasic increases in firing rate via two qualitatively distinct biophysical mechanisms: 1) a prolonged hyperpolarization preceding rebound bursts, accompanied by a hyperpolarizing shift in action potential threshold; and 2) a transient depolarization leading to high-frequency plateau bursts, associated with a depolarizing shift in action potential threshold. Our findings define a mechanistic framework for the biophysical implementation of dopamine neuron firing patterns in the intact brain.

Optimal Pipette Resistance, Seal Resistance, and Zero-Current Membrane Potential for Loose Patch or Breakthrough Whole-Cell Recording in vivo.

In vivo loose patch and breakthrough whole-cell recordings are useful tools for investigating the intrinsic and synaptic properties of neurons. However, the correlation among pipette resistance, seal condition, and recording time is not thoroughly clear. Presently, we investigated the recording time of different pipette resistances and seal conditions in loose patch and breakthrough whole-cell recordings. The recording time did not change with pipette resistance for loose patch recording (Rp-loose) and first increased and then decreased as seal resistance for loose patch recording (Rs-loose) increased. For a high probability of a recording time ≥30 min, the low and high cutoff values of Rs-loose were 21.5 and 36 MΩ, respectively. For neurons with Rs-loose values of 21.5-36 MΩ, the action potential (AP) amplitudes changed slightly 30 min after the seal. The recording time increased as seal resistance for whole-cell recording (Rs-tight) increased and the zero-current membrane potential for breakthrough whole-cell recording (MPzero-current) decreased. For a high probability of a recording time ≥30 min, the cutoff values of Rs-tight and MPzero-current were 2.35 GΩ and -53.5 mV, respectively. The area under the curve (AUC) of the MPzero-current receiver operating characteristic (ROC) curve was larger than that of the Rs-tight ROC curve. For neurons with MPzero-current values ≤ -53.5 mV, the inhibitory or excitatory postsynaptic current amplitudes did not show significant changes 30 min after the seal. In neurons with Rs-tight values ≥2.35 GΩ, the recording time gradually increased and then decreased as the pipette resistance for whole-cell recording (Rp-tight) increased. For the high probability of a recording time ≥30 min, the low and high cutoff values of Rp-tight were 6.15 and 6.45 MΩ, respectively. Together, we concluded that the optimal Rs-loose range is 21.5-36 MΩ, the optimal Rp-tight range is 6.15-6.45 MΩ, and the optimal Rs-tight and MPzero-current values are ≥2.35 GΩ and ≤ -53.5 mV, respectively. Compared with Rs-tight, the MPzero-current value can more accurately discriminate recording times ≥30 min and <30 min.

Electrophysiological Studies of GABAA Receptors Using QPatch II, the Next Generation of Automated Patch-Clamp Instruments.

Ligand-gated ion channels (LGICs) are a group of diverse ion channels that are gated by ligands and play important roles in normal physiological and pathological conditions. Many of them are drug targets that have been pursued, are being pursued, and will likely be pursued in the future by pharmaceutical companies and academic groups for a variety of diseases. One of those LGICs is the GABAA receptor, a heterooligomeric chloride channel that can be blocked and modulated at various sites. In order to study the receptor's functional response to compounds, the manual patch-clamp method provides a detailed but low-throughput electrophysiological characterization. QPatch II, a next-generation automated patch clamp machine that was recently developed by Sophion Bioscience, provides an automated electrophysiological study of ion channels. In this article, we use the GABAA receptor as an example for studying LGICs and describe two detailed protocols for using QPatch II to carry out pharmacological studies on the receptor. © 2020 by John Wiley & Sons, Inc. Basic Protocol 1: Ligand concentration-response experiment (GABAA receptor) on QPatch II Alternate Protocol: Non-cumulative ligand concentration-response experiment (GABAA receptor) on QPatch II Support Protocol 1: Cell culture of HEK293-hGABAA (α5ß3γ2) Support Protocol 2: Data analysis for Basic Protocol 1 Support Protocol 3: Data analysis for Alternate Protocol Basic Protocol 2: Antagonist dose-response experiment (GABAA receptor) on QPatch II Support Protocol 3: Data analysis for Basic Protocol 2.

Automated Planar Patch-Clamp Recording of P2X Receptors.

P2X receptors are a structurally and functionally distinctive family of ligand-gated ion channels that play important roles in mediating extracellular adenosine 5'-triphosphate (ATP) signaling in diverse physiological and pathophysiological processes. For several decades, the "manual" patch-clamp technique was regarded as the gold standard assay for investigating ion channel properties. More recently, breakthroughs in the development of automated patch-clamp technologies are enabling the study of ion channels, with much greater throughput capacities. These automated platforms, of which there are many, generate consistent, reliable, high-fidelity data. This chapter demonstrates the versatility of one of these technologies for ligand-gated ion channels, with a particular emphasis on protocols that address some of the issues of receptor desensitization that are commonly associated with P2X receptor-mediated currents.

Structure, function and techniques of investigation of the P2X7 receptor (P2X7R) in mammalian cells.

The P2X7 receptor [P2X7R or P2RX7 in National Center for Biotechnology Information (NCBI) gene nomenclature] is a member of the P2X receptor (P2XR) subfamily of P2 receptors (P2Rs). The P2X7R is an extracellular ATP-gated ion channel with peculiar permeability properties expressed by most cell types, mainly in the immune system, where it has a leading role in cytokine release, oxygen radical generation, T lymphocyte differentiation and proliferation. A role in cancer cell growth and tumor progression has also been demonstrated. These features make the P2X7R an appealing target for drug development in inflammation and cancer. The functional P2X7R, recently (partially) crystallized and 3-D solved, is formed by the assembly of three identical subunits (homotrimer). The P2X7R is preferentially permeable to small cations (Ca2+, Na+, K+), and in most (but not all) cell types also to large positively charged molecules of molecular mass up to 900Da. Permeability to negatively charged species of comparable molecular mass (e.g., Lucifer yellow) is debated. Several highly selective P2X7R pharmacological blockers have been developed over the years, thus providing powerful tools for P2X7R studies. Biophysical properties and coupling to several different physiological responses make the P2X7R amenable to investigation by electrophysiology and cell biology techniques, which allow its identification and characterization in many different cell types and tissues. A careful description of the physiological features of the P2X7R is a prerequisite for an effective therapeutic development. Here we describe the most common techniques to asses P2X7R functions, including patch-clamp, intracellular calcium measurements, and membrane permeabilization to large fluorescent dyes in a selection of different cell types. In addition, we also describe common toxicity assays used to verify the effects of P2X7R stimulation on cell viability.

Development of an Electrophysiological Assay for Kv7 Modulators on IonWorks Barracuda.

Relief from chronic pain continues to represent a large unmet need. The voltage-gated potassium channel Kv7.2/7.3, also known as KCNQ2/3, is a key contributor to the control of resting membrane potential and excitability in nociceptive neurons and represents a promising target for potential therapeutics. In this study, we present a medium throughput electrophysiological assay for the identification and characterization of modulators of Kv7.2/7.3 channels, using the IonWorks Barracuda™ automated voltage clamp platform. The assay combines a family of voltage steps used to construct conductance curves with a unique analysis method. Kv7.2/7.3 modulators shift the activation voltage and/or change the maximal conductance of the current, and both parameters have been used to quantify compound mediated effects. Both effects are expected to modulate neuronal excitability in vivo. The analysis method described assigns a single potency value that combines changes in activation voltage and maximal conductance and is expected to predict compound mediated changes in excitability.

Head-mounted approaches for targeting single-cells in freely moving animals.

Neural network processing is usually studied using the spike times of many extracellularly recorded neurons. Elucidating the cellular-synaptic mechanisms underlying these firing patterns requires identifying and controlling single cells and assessing their inputs. Single cell glass electrode techniques (intracellular, patch and juxtacellular) are well suited to filling this gap, in terms of physiology, cell identity and behavior. However, they are typically limited to in vitro and immobilized in vivo experiments, primarily due to the necessity for mechanical stability and steep learning curves. Several approaches have been recently developed to extend these technologies to freely moving animals. Here we summarize the advantages and results for different methods of single neuron glass recordings in vivo. We further review three approaches used to date for single cell recording in freely moving animals: static anchor systems, manual mechanic drives and motorized drives. Finally, we highlight new technologies capable of expanding the utility of single neuron recording in freely moving animals.