This work was funded by the National Heart, Lung, and Blood Institute (R01 HL073965, IL) and (T32 HL007829-24, ISF). The authors would also like to acknowledge the Scientific Machine Shop at the University of Illinois at Chicago for generating our latest MPP flow chambers.

The use of animals in our studies is approved by the University of Illinois at Chicago Animal Care Committee (#16-183).
1. Assembly of the Modified Parallel Plate Flow Chamber
NOTE: Please refer to Table 1 and Figure 1 for MPP flow chamber piece IDs. Please refer to Figure 1 for a schematic detailing the orientation of chamber pieces for assembly.
<ol>
	<li>To adhere the rectangular cover glass, piece D, to the bottom of piece C, first make the silicone elastomer solution by thoroughly mixing 500 &#181;L silicone elastomer curing agent into 5 mL of silicone elastomer base.</li>
	<li>Apply a thin layer of the silicone elastomer solution around the edges of the rectangular space of piece C and gently place the rectangular cover glass piece D directly on the elastomer solution such that piece D completely covers the open rectangular space of piece C. Carefully wipe away excess silicone elastomer solution.</li>
	<li>Repeat step 1.2 for adhering the rectangular cover glass, piece F, to the bottom of piece E and allow the silicone elastomer solution to cure over night at room temperature. 
	NOTE: Once cured, the rectangular cover glass will remain adhered for up to six months before needing to be replaced.</li>
	<li>Beginning with the bottom chamber piece, piece E, assemble the MPP flow chamber by sequentially placing each piece on top of the previous in the following order: piece E (bottom), piece C, piece B, piece A (top).</li>
	<li>Align the screw holes of each piece at the corners and tightly screw the pieces together to prevent leaks from occurring while administering flow to the MPP flow chamber.</li>
</ol>
2. Cell Preparation and Seeding into the MPP Flow Chamber
NOTE: Follow steps 2.1&#8722;2.7 for cultured endothelial cells. Follow the method detailed in steps 2.8&#8722;2.14 for isolating endothelial cells from the mouse mesenteric arterial arcade and preparation of freshly isolated endothelial cells.
<ol>
	<li>In a 6-well plate, place four to five 12 mm cover glass circles/well and seed cells between 10% and 30% confluency such that single cells can be accessed for electrophysiological recordings.</li>
	<li>Incubate cells under standard culture conditions (5% CO2, 37 &#176;C) for no less than 2 h to allow cells to adhere and no more than 24 h as endothelial cells in authors&#8217; experience become very flat and difficult to patch when seeded at sub-confluency for more than 24 h.</li>
	<li>Remove a cover glass containing adhered cells from a well of the 6-well plate, quickly rinse in phosphate-buffered saline (PBS), and transfer to a 35 mm Petri dish containing 2 mL electrophysiological bath solution (Table 2) prior to transfer to the MPP flow chamber.</li>
	<li>Transfer the cover glass circle to the rectangular cover glass, piece D, which is adhered to piece C of the MPP flow chamber being sure that adequate solution stays on the cover glass so that cells to not become exposed to air. Add the desired bath solution (~250 &#181;L) to the cells so that the cover glass circle and cells are completely submerged in solution.</li>
	<li>Position the cover glass circle such that it rests in the half closest to the vacuum reservoir side so that cells will be in line with the slit openings of piece B. Ensure that the glass cover circle adheres to piece D through solution-glass adhesion so that application of flow to the chamber does not disrupt the position of the cover glass circle.</li>
	<li>Assemble the MPP flow chamber by screwing the pieces together in the appropriate order, as described in steps 1.4 and 1.5 and as shown in Figure 1. Transfer the chamber to the microscope stage and immediately perfuse the chamber with bath solution such that solution reaches the vacuum reservoir for aspiration (~10 mL).</li>
	<li>Identify a healthy cell for the experiment by identifying a cell with a dark border and obvious nucleus. Avoid cells that appear to be blebbing or cells that are in contact with other cells. 
	NOTE: In the authors&#8217; laboratory, human aortic endothelial cells and primary mouse mesenteric endothelial cells in culture are used. However, any other type of adherent cell that is of interest to specific research needs can be used in the same way.</li>
	<li>Wash the isolated arterial arcade in dissociation solution (Table 2). Transfer the arcade to a 2 mL centrifuge tube containing 2 mL of pre-warmed (37 &#176;C) dissociation solution (recipe for dissociation solution shown in Table 2) containing neutral protease (0.5 mg/mL) and elastase (0.5 mg/mL). Incubate for 1 h at 37 &#176;C with gentle shaking every 10 min.</li>
	<li>Remove 1 mL of the neutral protease/elastase dissociation solution and add 1 mg/mL collagenase type 1. Return the collagenase solution to the solution containing the arteries for a final collagenase type 1 concentration of 0.5 mg/mL. Incubate for 2&#8722;3 min at 37 &#176;C.</li>
	<li>Using 5 grade forceps, quickly move the arcade onto a 35 mm diameter Petri dish containing a 750 &#181;L drop of fresh, chilled dissociation solution. Further dissociate the tissue by using two 20 G syringe needles to mechanically liberate endothelial cells from enzymatically digested arteries.</li>
	<li>Using a 9&#8221; disposable Pasteur glass pipet, triturate the cell solution 10x before transferring the cells to a new 1.5 mL centrifuge tube using the glass pipet.</li>
	<li>Wash the Petri dish with another 750 &#181;L of dissociation solution and transfer to the same tube. Using the glass pipette, further mechanically disperse the cells by pipetting at least 10x to create a single cell suspension being careful not to introduce bubbles that may damage endothelial cell integrity.</li>
	<li>Add 750 &#181;L of the endothelial cell suspension (~500&#8722;1,000 cells) directly to piece D of the MPP flow chamber on the half closest to the reservoir vacuum side. Allow the endothelial cells to adhere between 45 min and 1 h at room temperature.</li>
	<li>Assemble the MPP flow chamber by screwing the pieces together in the appropriate order as described in steps 1.4 and 1.5 and as shown in Figure 1. Transfer the chamber to the microscope stage and immediately perfuse the chamber with bath solution such that solution reaches the vacuum aspiration (~10 mL). Identify accessible endothelial cells by their rough and round phenotype19,20. 
	NOTE: A variety of digestion methods and enzyme cocktails have been used to isolate endothelial cells from different arterial beds. See Table 3 for detailed descriptions of protocols that have been used by a variety of investigators to isolate endothelial cells for patch clamp electrophysiology of mechanosensitive ion channels. These methods are likely suitable for use in combination with the MPP flow chamber.</li>
</ol>
3. Controlling Shear Stress to the MPP Flow Chamber for Electrophysiological Recordings of Shear-activated Mechanosensitive Ion Channels
<ol>
	<li>Set-up a gravity perfusion system by connecting a 30 mL graduated syringe cylinder to a 3-way luer lock fitted with microbore tubing (internal diameter: 0.05 inch, outer diameter: 0.09 inch) suited for insertion into the 3 mm diameter inlet hole of piece A of the MPP chamber.</li>
	<li>Attach the graduated cylinder to the outer face of the Faraday cage surrounding the electrophysiology rig (Figure 2) using double-sided tape. Prior to inserting the tubing in the MPP chamber, pre-fill the syringe and tubing with bath solution (see Table 2 for bath solution used for investigating inwardly rectifying K+ channels in endothelial cells). Insert the tubing into the MPP flow chamber inlet hole of piece A.</li>
	<li>Pre-fill the MPP flow chamber with solution such that solution is being removed in the vacuum reservoir. Stop flow to the chamber and refill the graduated cylinder to the top mark. Calculate flow rates manually by allowing the solution to flow through the chamber and using a stop-watch to calculate mL/s at a given syringe cylinder height.</li>
	<li>Raise or lower the syringe to alter flow, and thus shear in the chamber, and continue this process until a desired level of shear stress is found.</li>
	<li>Calculate shear stress in a parallel chamber using the following equation21: 
	&#964; = 6&#181;Q/h2w 
	where &#181; = fluid viscosity (g/cm&#183;s), Q = flow rate (mL/s), and parallel plate chamber width (w = 2.2 cm) and height (h = 0.1 cm). 
	NOTE: In the current gravity perfusion system, at a syringe cylinder height (as measured from the top of the cylinder) of 57 cm above the microscope stage, the flow rate is 0.3 mL/s. The shear calculated in the chamber at this syringe cylinder height and flow rate is 0.7 dyn/cm2. It should also be noted that other perfusion systems, such as a peristaltic pump, can be used to control flow to the MPP flow chamber. However, these devices may add unwanted turbulence and influence stability of the electrophysiology measurements under flow, therefore, using the gravity perfusion system described here is recommended.</li>
	<li>Transfer an assembled chamber containing adhered cells to the microscope stage of the electrophysiology rig and insert the tubing pre-filled with bath solution into the hole of piece A. Simultaneously fill the chamber and wash the cells with 10 mL of bath solution by turning the 3-way luer lock such that solution flows to the chamber.</li>
	<li>Once the desired patch configuration is successfully obtained allow channel currents to stabilize in a static bath at room temperature. Once currents have stabilized, apply shear in a step-wise fashion allowing increases in current to stabilize prior to the next step increase in shear stress. 
	NOTE: The authors find the most success with the perforated patch configuration when studying mechanoactivated ion channels in endothelial cells. To perform perforated whole-cell patch configurations, add 5 &#181;L of a 60 mg/mL stock amphotericin B in dimethyl sulfoxide (DMSO) to 1 mL of 0.2 &#181;m sterile filtered pipette solution. After generating a giga-ohm seal in the cell-attached configuration, perforated whole-cell patches form within 2&#8722;5 min.</li>
	<li>Remove shear exposure to the cells by stopping flow to the chamber allowing mechanosensitive channel currents to return to baseline currents observed in the static bath.</li>
	<li>Isolate mechanosensitive ion channel currents of interest by altering solution valence (e.g., 60 mM K+ in bath solution with 0 Ca2+ in pipette solution to study inwardly rectifying K+ channels. Table 2 shows example solution recipes) and/or pharmacological inhibition of potentially contaminating current sources.</li>
</ol>

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The authors have nothing to disclose.

The vascular system is constantly exposed to active hemodynamic forces, which activate mechanosensitive ion channels3,22 but the physiological roles of these channels in shear stress-induced mechanotransduction is only starting to emerge4,6,8. The mechanisms responsible for the mechanosensitivity of shear stress-activated channels remain unknown. The protocol detailed he...

Hemodynamic forces generated by the blood flow are well known to play major roles in endothelial and vascular function1,2. It is also well known that several types of ion channels acutely respond to changes in shear stress3,4,5 leading to the hypothesis that ion channels can be primary shear stress sensors. More recently, we and others showed that mechanosensitive ion channels play critical roles in several shear-stress sensitive vascular functions, including the vasoactive response to shear stress6,7,8, and developmental angiogenesis9. The mechanisms of the shear-stress sensitivity of ion channels, however, are almost totally unknown. This gap of knowledge is likely to be due to the technical difficulty of performing electrophysiological recordings under well-defined shear stress. In this article, therefore, we provide a step by step detailed protocol routinely performed in our lab to achieve this goal6,7,10,11.
The overall goal of this method is to allow the real-time investigation of ion channel mechanoactivation under well-defined shear stress in the physiological range. This is achieved by modifying a standard parallel plate flow chamber to allow an electrophysiological pipette to be lowered into the chamber and access cells grown on the bottom plate during the real time exposure to flow, providing a unique approach to achieve this goal6,7,11. In contrast, standard parallel plate flow chambers, described in prior publications can be used for the real time imaging analysis of cells exposed to shear forces12 or other non-invasive approaches13,14 but not for electrophysiology. Similarly, the cone and plate apparatus, another powerful approach to expose cells to shear stress15,16 is also not suitable for electrophysiological recordings. Thus, these flow devices do not allow the investigation of shear stress sensitivity of ion channels. The difficulty in performing electrophysiological recordings under flow is the main reason for the paucity of information about the mechanisms responsible for these crucial effects.
In terms of the alternative approaches to achieve the same goal, there are none that are as accurate or controlled. Some earlier studies attempted to record ion channel activity under flow by exposing cells to a stream of liquid coming from another pipette brought to the vicinity of a cell from above17,18. This is highly non-physiological, as the mechanical forces generated under these conditions have little in common with the physiological profiles of shear stress in the blood vessels. Similar concerns apply to the attempts to simulate physiological shear stress by perfusion of open chambers. As discussed in detail in our earlier study10, an open liquid-air interface creates multiple disturbances and recirculation, which are non-physiological. To address all these concerns, we have designed a modified parallel plate (MPP) flow chamber, also referred to as the &#8220;minimally invasive flow device&#8221; in our earlier studies6,7,10,11, made from acrylic and extensively used in our lab. However, in spite of the fact that the original description of the design has been published almost 20 years ago and is the only flow device that allows performing electrophysiological recordings under well-defined shear stress, this methodology has not been adopted by other labs and there are only very few studies that attempt to record currents under flow. We believe, therefore, that providing a detailed description for using the MPP flow chamber will be of great help to researches who are interested in mechanosensitive ion channels and vascular biology.

<table><tbody><tr><td>0.2 &amp;micro;m sterile syringe filters</td><td>VWR</td><td>28145-501</td><td>Used for filtering electrophysiolgoical pipette solution</td></tr><tr><td>5 grade forceps</td><td>Fine Scientific Tools</td><td>1252-30</td><td>Used for transferring digested arteries to fresh solution</td></tr><tr><td>9&quot; Pasteur Pipet</td><td>Fisher Scientifc</td><td>13-678-20D</td><td>Used for mechanically disrupting digested arteries and transferring freshly isolated endohtelial cells&amp;nbsp;</td></tr><tr><td>12 mm diameter Cover glass circles</td><td>Fisher Scientifc</td><td>12-545-80</td><td>For use with studies involving cultured cells and multiple treatments. Cells adhered to the cover glass are used for patch clamp analyses</td></tr><tr><td>24 mm x 40 mm Rectangluar Cover glass</td><td>Sigma-Aldrich</td><td>CLS2975224</td><td>Cover glass to be added to MPP flow chamber pieces C (Figure 1)</td></tr><tr><td>24 mm x 50 mm Rectangular Cover glass</td><td>Sigma-Aldrich</td><td>CLS2975245</td><td>Cover glass to be added to MPP flow chamber E (Figure 1)</td></tr><tr><td>20 G syringe needles</td><td>Becton Dickinson and Co</td><td>305175</td><td>For use in mechanical disruption of digested mesenteric arteries</td></tr><tr><td>35 mm Petri dish</td><td>Genesee Scientific</td><td>32-103</td><td>For use in mechanical disruption of digested mesenteric arteries</td></tr><tr><td>Amphotericin B solubilized</td><td>Sigma-Aldrich</td><td>A9528-50MG</td><td>Used for generating the perforated whole-cell patch configuration.</td></tr><tr><td>Collagenase, type I</td><td>Worthington Biochemical</td><td>100 mg - LS004194</td><td>Enzyme used in our laboratory as a brief digestion following the initial cocktail of neutral protease and elastase</td></tr><tr><td>Dimethyl Sulfoxide (DMSO)</td><td>Fisher Scientifc</td><td>67-68-5</td><td>Solvent for Amphotericin B used in perforated whole-cell patch clamp</td></tr><tr><td>Elastase, lyophilized</td><td>Worthington Biochemical</td><td>25 mg - LS002290&amp;nbsp;</td><td>Enzyme used in our laboratory in a cocktail with neutral protease/dispase to begin digestion of arteries for endothelial cell isolation.</td></tr><tr><td>Falcon Tissue culture Plate, 6-well, Flat Bottom with Low Evaporation Lid&amp;nbsp;</td><td>Corning</td><td>353046</td><td>For use with studies involving cultured cells and multiple treatments</td></tr><tr><td>Neutral protease/dispase</td><td>Worthington Biochemical</td><td>10 mg- LS02100 50 mg - LS02104</td><td>Enzyme used in our laboratory in a cocktail with elastase to begin digestion of arteries for endothelial cell isolation</td></tr><tr><td>SylGard&amp;nbsp;</td><td>World Precision Instruments</td><td>SYLG184</td><td>Silicone elastomer for adhering the rectangular cover slip to the MPP flow chamber pieces C and E (Figure 1)</td></tr><tr><td>Tygon ND 10-80 tubing</td><td>Microbore Tubing</td><td>AAQ04133</td><td>ID: 0.05 in, OD: 0.09 in, inlet perfusion tubing for adminsitering flow to the chamber</td></tr></tbody></table>

electrophysiological recordings of single-cell ion currents under well-defined shear stress

Fluid shear stress is well known to play a major role in endothelial function. In most vascular beds, elevated shear stress from acute increases in blood flow triggers a signaling cascade resulting in vasodilation thereby alleviating mechanical stress on the vascular wall. The pattern of shear stress is also well known to be a critical factor in the development of atherosclerosis with laminar shear stress being atheroprotective and disturbed shear stress being pro-atherogenic. While we have a detailed understanding of the various intermediate cell signaling pathways, the receptors that first translate the mechanical stimulus into chemical mediators are not completely understood. Mechanosensitive ion channels are critical to the response to shear and regulate shear-induced cell signaling thereby controlling the production of vasoactive mediators. These channels are among the earliest activated signaling components to shear and have been linked to shear-induced vasodilation through promoting nitric oxide production (e.g., inwardly rectifying K+ [Kir] and transient receptor potential [TRP] channels) and endothelium hyperpolarizing factor (e.g., Kir and calcium-activated K+ [KCa] channels) and shear-induced vasoconstriction through an undetermined mechanism that involves piezo channels. Understanding the biophysical mechanism by which these channels are activated by shear forces (i.e., directly or through a primary mechano-receptor) could provide potential new targets to resolve the pathophysiology associated with endothelial dysfunction and atherogenesis. It is still a major challenge to record flow-induced activation of ion channels in real time using electrophysiology. The standard methods to expose cells to well-defined shear stress, such as the cone and plate rheometer and closed parallel plate flow chamber do not allow real time study of ion channel activation. The goal of this protocol is to describe a modified parallel plate flow chamber that allows real time electrophysiological recording of mechanosensitive ion channels under well-defined shear stress.

Multiple photographs showing different views of the MPP flow chamber on the microscope stage (upper panel) and a schematic representation of the MPP flow chamber (bottom panel) are shown in Figure 1. The schematic details the dimensions of the entire device and flow chamber. Figure 2 shows a photograph of the gravity perfusion system to the MPP flow chamber in our laboratory (upper panel). Also shown is a schematic representation...

Watch this Scientific Journal Video about Electrophysiological Recordings of Single-cell Ion Currents Under Well-defined Shear Stress at JoVE.com

Electrophysiological Recordings of Single-cell Ion Currents Under Well-defined Shear Stress

The goal of this protocol is to describe a modified parallel plate flow chamber for use in investigating real time activation of mechanosensitive ion channels by shear stress.

Fluid shear stress is well known to play a major role in endothelial function. In most vascular beds, elevated shear stress from acute increases in blood ...

electrophysiological-recordings-single-cell-ion-currents-under-well

Research

JoVE Journal

Immunology and Infection

6.1K Views.  University of Illinois at Chicago. Het gebruik van de gewijzigde parallelle plaat stroomkamer kan onderzoekers helpen bij het beantwoorden van belangrijke vragen met betrekking tot de functionele reactie van mechanosensitive ionkanalen op schuifstress. Het belangrijkste voordeel van deze techniek is dat het mogelijk maakt real-time onderzoek te stromen geactiveerde ionkanalen in een gemakkelijk te monteren, herbruikbare parallelle plaat flow kamer. Om te beginnen met de procedure, breng een dunne laag siliconen elastomeer oplo...

Video: Electrophysiological Recordings of Single-cell Ion Currents Under Well-defined Shear Stress

Whole-cell Recordings of Light Evoked Excitatory Synaptic Currents in the Retinal Slice

We use the whole-cell patch clamp technique to study the synaptic circuitry that underlies visual information processing in the retina. In this video, we will guide you through the process of performing whole-cell recordings of light evoked currents of individual cells in the retinal slice preparation. We use the aquatic tiger salamander as an animal model. We begin by describing the dissection of the eye and show how slices are mounted for electrophysiological recordings. Once the slice is placed in the recording chamber, we demonstrate how to perform whole-cell voltage clamp recordings. We then project visual stimuli onto the photoreceptors in the slice to elicit light-evoked current responses. During the recording we perfuse the slice with pharmacological agents, whereby an 8-channel perfusion system allows us to quickly switch between different agents.  The retinal slice preparation is widely used for patch clamp recordings in the retina, in particular to study amacrine or bipolar cells, which are not accessible in a whole-mount preparation.

This video shows the process of whole-cell voltage clamp recordings in the retinal slice of the aquatic tiger salamander. We demonstrate the preparation of the slice as well as how to perform patch clamp recordings during visual stimulation of the retina.

We use the whole-cell patch clamp technique to study the synaptic circuitry that underlies visual information processing in the retina. In this video, we ...

Biology

Enhancing Targeted Cancer Diagnosis by Quantifying Single-Cell Mechanics

Shear Assay Protocol for the Determination of Single-Cell Material Properties

Irregular biomechanics are a hallmark of cancer biology subject to extensive study. The mechanical properties of a cell are similar to those of a material. A cell's resistance to stress and strain, its relaxation time, and its elasticity are all properties that can be derived and compared to other types of cells. Quantifying the mechanical properties of cancerous (malignant) versus normal (non-malignant) cells allows researchers to further uncover the biophysical fundamentals of this disease. While the mechanical properties of cancer cells are known to consistently differ from the mechanical properties of normal cells, a standard experimental procedure to deduce these properties from cells in culture is lacking. 
This paper outlines a procedure to quantify the mechanical properties of single cells in vitro using a fluid shear assay. The principle behind this assay involves applying fluid shear stress onto a single cell and optically monitoring the resulting cellular deformation over time. Cell mechanical properties are subsequently characterized using digital image correlation (DIC) analysis and fitting an appropriate viscoelastic model to the experimental data generated from the DIC analysis. Overall, the protocol outlined here aims to provide a more effective and targeted method for the diagnosis of difficult-to-treat cancers.

Author Spotlight: Shear Assay Protocol for the Determination of Single-Cell Material Properties  

This protocol outlines the quantification of the mechanical properties of cancerous and non-cancerous cell lines in vitro. Conserved differences in the mechanics of cancerous and normal cells can act as a biomarker that may have implications in prognosis and diagnosis.

Irregular biomechanics are a hallmark of cancer biology subject to extensive study. The mechanical properties of a cell are similar to those of a ...

Cancer Research

Designing Microfluidic Devices for Studying Cellular Responses Under Single or Coexisting Chemical/Electrical/Shear Stress Stimuli

Microfluidic devices are capable of creating a precise and controllable cellular micro-environment of pH, temperature, salt concentration, and other physical or chemical stimuli. They have been commonly used for in vitro cell studies by providing in vivo like surroundings. Especially, how cells response to chemical gradients, electrical fields, and shear stresses has drawn many interests since these phenomena are important in understanding cellular properties and functions. These microfluidic chips can be made of glass substrates, silicon wafers, polydimethylsiloxane (PDMS) polymers, polymethylmethacrylate (PMMA) substrates, or polyethyleneterephthalate (PET) substrates. Out of these materials, PMMA substrates are cheap and can be easily processed using laser ablation and writing. Although a few microfluidic devices have been designed and fabricated for generating multiple, coexisting chemical and electrical stimuli, none of them was considered efficient enough in reducing experimental repeats, particular for screening purposes. In this report, we describe our design and fabrication of two PMMA-based microfluidic chips for investigating cellular responses, in the production of reactive oxygen species and the migration, under single or coexisting chemical/electrical/shear stress stimuli. The first chip generates five relative concentrations of 0, 1/8, 1/2, 7/8, and 1 in the culture regions, together with a shear stress gradient produced inside each of these areas. The second chip generates the same relative concentrations, but with five different electric field strengths created within each culture area. These devices not only provide cells with a precise, controllable micro-environment but also greatly increase the experimental throughput.

Micro-fabricated devices integrated with fluidic components provide an in vitro platform for cell studies mimicking the in vivo micro-environment. We developed polymethylmethacrylate-based microfluidic chips for studying cellular responses under single or coexisting chemical/electrical/shear stress stimuli.

Microfluidic devices are capable of creating a precise and controllable cellular micro-environment of pH, temperature, salt concentration, and other ...

Bioengineering

Whole-cell Patch-clamp Recordings in Brain Slices

Whole-cell patch-clamp recording is an electrophysiological technique that allows the study of the&#160;electrical properties of a substantial part of the neuron. In this configuration, the micropipette is in tight contact with the cell membrane, which prevents current leakage and thereby provides more accurate ionic current measurements than the previously used intracellular sharp electrode recording method. Classically, whole-cell recording can be performed on neurons in various types of preparations, including cell culture models, dissociated neurons, neurons in brain slices, and in intact anesthetized or awake animals. In summary, this technique has immensely contributed to the understanding of passive and active biophysical properties of excitable cells. A major advantage of this technique is that it provides information on how specific manipulations (e.g., pharmacological, experimenter-induced plasticity) may alter specific neuronal functions or channels in real-time. Additionally, significant opening of the plasma membrane allows the internal pipette solution to freely diffuse into the cytoplasm, providing means for introducing drugs, e.g., agonists or antagonists of specific intracellular proteins, and manipulating these targets without altering their functions in neighboring cells. This article will focus on whole-cell recording performed on neurons in brain slices,&#160;a preparation that has the advantage of recording neurons in relatively well preserved brain circuits, i.e., in a physiologically relevant context. In particular,&#160;when combined with appropriate pharmacology, this technique is a powerful tool allowing identification of specific neuroadaptations that occurred following any type of experiences, such as learning, exposure to drugs of abuse, and stress. In summary, whole-cell patch-clamp recordings in brain slices provide means to measure in ex vivo preparation long-lasting changes in neuronal functions that have developed in intact awake animals.

This protocol describes basic procedural steps for performing whole-cell patch-clamp recordings. This technique allows the study of&#160;the electrical behavior of neurons, and when performed in brain slices, allows the assessment of various neuronal functions from neurons that are still integrated in relatively well preserved brain circuits.

Whole-cell patch-clamp recording is an electrophysiological technique that allows the study of the&#160;electrical properties of a substantial part of the ...

Neuroscience

In Utero Electroporation Approaches to Study the Excitability of Neuronal Subpopulations and Single-cell Connectivity

The nervous system is composed of an enormous range of distinct neuronal types. These neuronal subpopulations are characterized by, among other features, their distinct dendritic morphologies, their specific patterns of axonal connectivity, and their selective firing responses. The molecular and cellular mechanisms responsible for these aspects of differentiation during development are still poorly understood.
Here, we describe combined protocols for labeling and characterizing the structural connectivity and excitability of cortical neurons. Modification of the in utero electroporation (IUE) protocol allows the labeling of a sparse population of neurons. This, in turn, enables the identification and tracking of the dendrites and axons of individual neurons, the precise characterization of the laminar location of axonal projections, and morphometric analysis. IUE can also be used to investigate changes in the excitability of wild-type (WT) or genetically modified neurons by combining it with whole-cell recording from acute slices of electroporated brains. These two techniques contribute to a better understanding of the coupling of structural and functional connectivity and of the molecular mechanisms controlling neuronal diversity during development. These developmental processes have important implications on axonal wiring, the functional diversity of neurons, and the biology of cognitive disorders.

In Utero Electroporation Approaches to Study the Excitability of Neuronal Subpopulations and Single-cell Connectivity

This manuscript provides protocols that use in utero electroporation (IUE) to describe the structural connectivity of neurons at the single-cell level and the excitability of fluorescently labeled neurons. Histology is used to characterize dendritic and axonal projections. Whole-cell recording in acute slices is used to investigate excitability.

The nervous system is composed of an enormous range of distinct neuronal types. These neuronal subpopulations are characterized by, among other features, ...

Isolation of Human Atrial Myocytes for Simultaneous Measurements of Ca2+ Transients and Membrane Currents

The study of electrophysiological properties of cardiac ion channels with the patch-clamp technique and the exploration of cardiac cellular Ca2+ handling abnormalities requires isolated cardiomyocytes. In addition, the possibility to investigate myocytes from patients using these techniques is an invaluable requirement to elucidate the molecular basis of cardiac diseases such as atrial fibrillation (AF).1 Here we describe a method for isolation of human atrial myocytes which are suitable for both patch-clamp studies and simultaneous measurements of intracellular Ca2+ concentrations. First, right atrial appendages obtained from patients undergoing open heart surgery are chopped into small tissue chunks ("chunk method") and washed in Ca2+-free solution. Then the tissue chunks are digested in collagenase and protease containing solutions with 20 &mu;M Ca2+. Thereafter, the isolated myocytes are harvested by filtration and centrifugation of the tissue suspension. Finally, the Ca2+ concentration in the cell storage solution is adjusted stepwise to 0.2 mM. We briefly discuss the meaning of Ca2+ and Ca2+ buffering during the isolation process and also provide representative recordings of action potentials and membrane currents, both together with simultaneous Ca2+ transient measurements, performed in these isolated myocytes.

Isolation of Human Atrial Myocytes for Simultaneous Measurements of Ca2+ Transients and Membrane Currents

We describe the isolation of human atrial myocytes which can be used for intracellular Ca2+ measurements in combination with electrophysiological patch-clamp studies.

The study of  electrophysiological properties of cardiac ion channels with the patch-clamp  technique and the exploration of cardiac cellular Ca2+ ...

Mechano-Node-Pore Sensing: A Rapid, Label-Free Platform for Multi-Parameter Single-Cell Viscoelastic Measurements

Cellular mechanical properties are involved in a wide variety of biological processes and diseases, ranging from stem cell differentiation to cancer metastasis. Conventional methods for measuring these properties, such as atomic force microscopy (AFM) and micropipette aspiration (MA), capture rich information, reflecting a cell's full viscoelastic response; however, these methods are limited by very low throughput. High-throughput approaches, such as real-time deformability cytometry (RT-DC), can only measure limited mechanical information, as they are often restricted to single-parameter readouts that only reflect a cell's elastic properties. In contrast to these methods, mechano-node-pore sensing (mechano-NPS) is a flexible, label-free microfluidic platform that bridges the gap in achieving multi-parameter viscoelastic measurements of a cell with moderate throughput. A direct current (DC) measurement is used to monitor cells as they transit a microfluidic channel, tracking their size and velocity before, during, and after they are forced through a narrow constriction. This information (i.e., size and velocity) is used to quantify each cell's transverse deformation, resistance to deformation, and recovery from deformation. In general, this electronics-based microfluidic platform provides multiple viscoelastic cell properties, and thus a more complete picture of a cell's mechanical state. Because it requires minimal sample preparation, utilizes a straightforward electronic measurement (in contrast to a high-speed camera), and takes advantage of standard soft lithography fabrication, the implementation of this platform is simple, accessible, and adaptable to downstream analysis. This platform's flexibility, utility, and sensitivity have provided unique mechanical information on a diverse range of cells, with the potential for many more applications in basic science and clinical diagnostics.

Presented here is a method to mechanically phenotype single cells using an electronics-based microfluidic platform called mechano-node-pore sensing (mechano-NPS). This platform maintains moderate throughput of 1-10 cells/s while measuring both the elastic and viscous biophysical properties of cells.

Cellular mechanical properties are involved in a wide variety of biological processes and diseases, ranging from stem cell differentiation to cancer ...

Measurement of Extracellular Ion Fluxes Using the Ion-selective Self-referencing Microelectrode Technique

Cells from animals, plants and single cells are enclosed by a barrier called the cell membrane that separates the cytoplasm from the outside. Cell layers such as epithelia also form a barrier that separates the inside from the outside or different compartments of multicellular organisms. A key feature of these barriers is the differential distribution of ions across cell membranes or cell layers. Two properties allow this distribution: 1) membranes and epithelia display selective permeability to specific ions; 2) ions are transported through pumps across cell membranes and cell layers. These properties play crucial roles in maintaining tissue physiology and act as signaling cues after damage, during repair, or under pathological condition. The ion-selective self-referencing microelectrode allows measurements of specific fluxes of ions such as calcium, potassium or sodium at single cell and tissue levels. The microelectrode contains an ionophore cocktail which is selectively permeable to a specific ion. The internal filling solution contains a set concentration of the ion of interest. The electric potential of the microelectrode is determined by the outside concentration of the ion. As the ion concentration varies, the potential of the microelectrode changes as a function of the log of the ion activity. When moved back and forth near a source or sink of the ion (i.e. in a concentration gradient due to ion flux) the microelectrode potential fluctuates at an amplitude proportional to the ion flux/gradient. The amplifier amplifies the microelectrode signal and the output is recorded on computer. The ion flux can then be calculated by Fick&#8217;s law of diffusion using the electrode potential fluctuation, the excursion of microelectrode, and other parameters such as the specific ion mobility. In this paper, we describe in detail the methodology to measure extracellular ion fluxes using the ion-selective self-referencing microelectrode and present some representative results.

Transporters in cell membranes allow differential segregation of ions across cell membranes or cell layers and play crucial roles during tissue physiology, repair and pathology. We describe the ion-selective self-referencing microelectrode that allows the measurement of specific ion fluxes at single cells and tissues in vivo.

Cells from animals, plants and single cells are enclosed by a barrier called the cell membrane that separates the cytoplasm from the outside. Cell layers ...

Application of Electrophysiology Measurement to Study the Activity of Electro-Neutral Transporters

The transport of ions through cell membranes ensures the fine control of ion content within and outside the cell that is indispensable for cell survival. These transport mechanisms are mediated by the activities of specialized transporter proteins. Specifically,pH dynamics are finely controlled by plasma membrane proton (H+) extrusion systems, such as the Na+/H+ exchanger (NHE) protein family. Despite extensive efforts to study the mechanisms underlying NHE regulation, our current understanding of the biophysical and molecular properties of the NHE family is inadequate because of the limited availability of methods to effectively measure NHE activity. In this manuscript, we used H+-selective electrodes during whole-cell patch clamping recording to measure NHE-induced H+ flux. We proposed this approach to overcome some limitations of typically used methods to measure NHE activity, such as radioactive uptake and fluorescent membrane permeants. Measurement of NHE activity using the described method enables high sensitivity and time resolution and more efficient control of intracellular H+ concentrations. H+-selective electrodes are based on the fact that transporter activity creates an ion gradient in close proximity to the cell membrane. An H+-selective electrode moving up to and away from the cell membrane in a repetitive, oscillatory fashion records a voltage difference that is dependent on H+ flux. While H+-selective electrodes are used to detect H+ flux moving out of the cell, the patch clamp method in the whole-cell configuration is used to control the intracellular ion composition. Moreover, application of the giant patch clamp technique allows modification of the intracellular composition of not only ions but also lipids. The transporter activity of NHE isoform 3 (NHE3) was measured using this technical approach to study the molecular basis of NHE3 regulation by phosphoinositides.

This manuscript describes the applications of proton-selective electrodes and patch clamping methods to measure the activity of proton transport systems. These methods overcome some limitations of techniques commonly used to study proton transport activity, such as moderate sensitivity, time resolution and insufficient intracellular milieu control.

The transport of ions through cell membranes ensures the fine control of ion content within and outside the cell that is indispensable for cell survival. ...

Recording of Inward Rectifying K+ Currents in Freshly Isolated Basilar Artery Smooth Muscle Cells by Patch Clamp Technique

Cerebrovascular disease is a prevalent condition among the elderly, with its incidence steadily rising. The basilar artery is a critical cerebral vessel that supplies the pons, cerebellum, posterior brain regions, and inner ear. Potassium (K+) channel activity plays a significant role in determining vascular tone by regulating the cell membrane potential. Activation of inward rectifying K+ (Kir) channels, like other K+ channels, leads to cell membrane hyperpolarization and vasodilation. In this study, freshly isolated smooth muscle cells from the basilar artery were used to record Kir currents via the whole-cell patch clamp technique. The effects of 100 &#181;mol/L BaCl2, a Kir channel inhibitor, and 10 &#181;mol/L sodium nitroprusside (SNP), a nitro vasodilator, on Kir channel currents were investigated. The results demonstrated that BaCl2 inhibited Kir channel currents in basilar artery smooth muscle cells, whereas SNP enhanced these currents. This protocol provides a comprehensive guide for preparing freshly isolated arterial smooth muscle cells and recording Kir channel currents using the patch clamp technique, offering a valuable resource for researchers seeking to master this method.

This protocol describes a rapid and efficient method for isolating smooth muscle cells from the rat basilar artery and recording inward rectifying potassium channel currents in these cells using the whole-cell patch clamp technique. It offers a novel approach for researchers studying the basilar artery and ion channels.

Cerebrovascular disease is a prevalent condition among the elderly, with its incidence steadily rising. The basilar artery is a critical cerebral vessel ...

Electrophysiological Recordings of Single-cell Ion Currents Under Well-defined Shear Stress

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