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Tutorials for Electrophysiological Recordings in Tissue Engineering Chuang Du, Will Collins, Will Cantley, Disha Sood, and David L Kaplan ACS Biomater. Sci. Eng., Just Accepted Manuscript • DOI: 10.1021/acsbiomaterials.7b00318 • Publication Date (Web): 11 Aug 2017 Downloaded from http://pubs.acs.org on August 21, 2017

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ACS Biomaterials Science & Engineering

Tutorials for Electrophysiological Recordings in Neuronal Tissue Engineering Chuang Du, Will Collins,+ Will Cantley,# Disha Sood, and David L Kaplan* Department of Biomedical Engineering, Tufts University, 4 Colby Street, Medford MA 02155, USA

AUTHOR INFORMATION Corresponding Author: *E-mail: [email protected] Tufts University Science and Technology Center 4 Colby Street, Medford, MA 02155 Additional Affiliations: +

Program in Pharmacology and Experimental Therapeutics, Sackler School of Graduate Biomedical

Sciences, Tufts University, Boston, MA 02111, USA #

Program in Cellular, Molecular, and Developmental Biology, Sackler School of Graduate Biomedical

Sciences, Tufts University, Boston, MA 02111, USA

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ABSTRACT

Electrophysiology is a powerful tool to examine cellular functions, but the use of the techniques remains challenging outside of physiology and neuroscience fields. We aim to provide a practical methods guide for electrophysiological recordings to nonexperts in the field to help with the utility of these important research tools. We focus on two techniques that are critical in the context of tissue engineering, wholecell patch clamp recording for assessing cellular functions and extracellular field potential recording for evaluating network activities. Specific examples are presented to demonstrate how the techniques were applied to tissue engineering studies.

Keywords: 2D, 3D, Patch Clamp, Whole-Cell Recording, Extracellular Field Potential

INTRODUCTION

Bio-electricity is one of the fundamental properties of the cells. In excitable cells, membrane potential and its changes are used to modulate cell functions, and to encode, propagate, transmit, and integrate cell information. The study of the bio-electrical activity of cells thus provides a powerful tool to examine cellular functions, and gave rise to the discipline of electrophysiology. Employing electrical means, electrophysiology has evolved from simply measuring and describing electrical activity of cells and tissues to exploring the molecular structures of ion channel proteins.1-6 Many in depth, theoretical, and historical reviews of electrophysiological techniques exist.7-12 Here our goal is to provide a more useful methods guide for the nonexpert in the field, to facilitate broader utility and more accurate assessments of biological systems using electrophysiological techniques. 2


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The guiding principle of electrophysiology is simple and is based on Ohm’s law, which states that the current through a conductor between two points is directly proportional to the potential difference across the two points (I = V / R). However, the use of these techniques remains challenging outside of physiology and neuroscience fields. We aim to provide a practical methods guide for electrophysiological recordings in the context of tissue engineering. We focus on two main aspects that are critical to tissue engineering, cellular functions and network activities. We have used whole-cell patch clamp techniques to study cellular functions, and network activities were examined with extracellular field potential recording. Both techniques are mainly used in our neuronal tissue engineering studies, but the whole-cell patch clamp recording can also be applied to other cell types as diverse as cardiac and skeletal myocytes, and cancer cell-line cells. Table 1 summarizes the utilities and limitations of these two techniques. This methods guide will cover mainly the practical aspects. In each section, a general principle description of the method is given first, followed by a practical guide with more details. Specific examples are presented to demonstrate how the techniques were applied to our tissue engineering studies.

Aside from the two main methods of focus in this paper, there are other options that can be considered. For example, voltage-sensitive dyes or genetically expressed voltage-sensitive fluorescent proteins have been widely used for assessments of bioelectric features in biological systems.13-17 Many challenges are present with any electrophysiological assessments of biological systems. One of the limitations with classical electrophysiology is that it is inherently dependent on the point of observation within a volume of cells and tissues. Recordings with multiple electrode arrays (MEA) can alleviate this problem by increasing the number of points of observation. Optical imaging with voltage-sensitive dyes or 3



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fluorescent proteins overcomes this limitation by scanning the whole field. But the optical techniques have their own challenges as well. These include difficulties of extracting exact physiological bioelectrical values from optical ratio-metric measurements, interferences with the physiological functions of cell membranes, phototoxicity, and low signal-to-noise ratio. We do not plan to cover these techniques, but there are suitable recent reviews to help.15,18-20

Table 1. Utilities and limitations of the whole-cell patch clamp recording and the extracellular field potential recording methods.

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Method

Utility, type of information

Limitations

Whole-Cell Intracellular Recording

Cellular information from specific cell, including resting membrane potential, spontaneous and evoked action potentials, voltage- and ligandactivated membrane currents.

High technical difficulty;

2D Extracellular Recording

Low technical difficulty;

Limited to detecting spiking information;

Cells can aggregate in 2D cultures and clusters can be visually identified; Information includes spontaneous and evoked spikes from cell clusters in 2D cultures; Synaptic interactions between cells can be studied by stimulation-evoked responses and by pharmacological tools. 3D Extracellular Recording

Low technical difficulty; Information includes spontaneous and evoked spikes from cells present within 3D cultures; Synaptic interactions between cells can be studied by stimulation-evoked responses and by pharmacological tools.

Whole-Cell Patch Clamp Recording

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Limited to 2D cultures thus lacking information on 3D interactions.

Spike amplitude information is not quantifiable; Source of the detected activity is difficult to determine, but the problem can be overcome by employing optogenetic tools.

Limited to detecting spiking information; Spike amplitude information is not quantifiable; Source of the detected activity is difficult to determine, but the problem can be overcome by employing optogenetic tools; Cells or cellular clusters can not be identified visually, making the recording very random. Increasing coverage of sampling areas might alleviate the randomness problem.


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Generally, the method follows what was originally described by Hamill et al.21 In principle, a direct, low resistant path is established between the cell and the electrode after the tip of the electrode forms an extremely high resistant gigaohms seal with the cell. This direct, low resistant path allows the electrode to detect all electrical current and potential changes coming from the cell as a whole across its entire cellular membrane, giving rise to its name of whole-cell recording. Briefly, acutely dispersed neurons or other cells are plated on poly-D-lysine (or other substrate)-coated 35 mm culture dishes, or on substrate-coated 10 mm glass cover slips, and are maintained at 37oC in 5% CO2 incubators. At the time of recording, the dishes or cover slips are placed inside a recording chamber that contains an extracellular solution. Cells are visualized with a phase-contrast inverted microscope. A patch electrode filled with an intracellular solution is secured into an electrode holder with a suction port and placed on top of the cells of interest (Fig. 1B1). The electrode tip is gently dipped into the cell membrane (Fig. 1B2), then forms a gigaohms seal with the membrane when a slight suction is applied (Fig. 1B3). A stronger suction after that breaks the membrane patch within the giga seal and establishes a direct connection between the inside of the cells (cytosol) and the patch electrode via the intracellular solution (Fig. 1B4). Membrane potentials and whole-cell membrane currents can then be measured either at room temperature or at a desired temperature by whole-cell patch-clamp using a patch amplifier. Recorded signals can be processed, digitized, displayed, recorded, and analyzed in a computer. Besides whole-cell recording, other patch clamp recording configurations also can be used. These include cell-attached, inside-out, outside-out, and perforated patch (a whole-cell patch variation), which are outside of the focus of this methods paper.

Practical Guides for Whole-Cell Patch Clamp Recording 1. Basic Recording Set-Up 6


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The basic recording set-up consists of three major components: mechanical-positioning, visualoptical, and electrical. The mechanical control starts with a vibration-free table, such as a TMC (Technical Manufacturing Corporation) or Newport air table. The micron precision position control is achieved by electric and hydraulic micromanipulators, such as Sutter and Narashige. When setting up the micromanipulator, a 45o approach is a good setting for an inverted microscope, and can be as shallow as 20o for upright microscope. The visual-optical component consists of a high magnification, inverted microscope. An inverted microscope is preferred over an upright microscope because it provides ample working space and easy access to the cell preparations. Finally, the electrical component consists of an electrical noise shielding mechanism such as a Faraday cage, a patch clamp amplifier such as an Axopatch amplifier, and a data acquisition system that includes a digitizer such as the Digidata, acquisition software such as the pClamp software suite, and a computer. The set-up is usually built like a tower on top of an air table, with the microscope as the center piece (Fig. 1A). One or multiple micromanipulators and amplifier headstages are positioned around the optical center of the microscope. The headstage has connections for the electrode holder and the signal ground. The ground can be simply made with chlorided, coiled silver wire, by coiling bare silver wire, then dipping it into bleach for a few hours. Alternatively, one can use prefabricated Ag/AgCl pellets available from commercial electrophysiology suppliers such as WPI and Warner Instruments. A well chlorinated silver wire is as good as Ag/AgCl pellets. It is easy to make, but may not last as long as Ag/AgCl pellets. For this reason, Ag/AgCl pellets should be the first choice. Usually, there is no concern for having the AgCl in direct contact with the bathing solution as the solution is Cl- based. The amplifier and the data acquisition system are positioned outside of the Faraday cage, often housed in a rig to further shield any interference these instruments may produce from affecting the recording in the cage. 2. Solutions 7



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In the whole-cell recording configuration, the recording solutions usually mimic the extracellular and the intracellular ionic conditions, meaning high Na+ for extracellular solution and high K+ and low chloride for intracellular solution. A common set of recording solutions that we use in our recording consists of the following components in mM: NaCl 140, KCl 2.8, CaCl2 2, MgCl2 2, HEPES 10, and Dglucose 10, with pH adjusted to 7.4 with NaOH (for extracellular solution), and K gluconate 140, NaCl 10, MgCl2 2, HEPES 10, EGTA 1, MgATP 4, and NaGTP 0.3, with pH adjusted to 7.3 with KOH (for intracellular solution). Depending on the experimental needs, the compositions of the solutions can be changed. For example, external Na+ and Ca2+ can be substituted by K+ to identify the carrier of inward currents, external K+ can be elevated in place of Na+ to produce membrane depolarization, and cesium can replace internal K+ to block voltage-dependent K channels. All the solution changes are done under equal molar fashion by replacing one ion with another with no change in osmolarity. 3. Environment Control Environment control consists mainly of solution exchange control and temperature control. For solution exchange, several arrangements can be made: bath perfusion, focal perfusion, focal pressure ejection, and fast solution switch with piezoelectric drive. The bath perfusion method is global and slow, but is easy to set up. The focal perfusion is localized and fast, and requires more effort to set it up. The piezo-driven solution switch is the fastest solution exchange method and requires the use of specialized instruments. The flow of the solutions can be achieved by gravity, by air pressure or by using pumps. To control the temperature of all or just the recorded cells, solution can be heated around and in the chamber, or be preheated just before reaching to the cells. The former is efficient relative to the latter and heats up a large area. The latter is more specific in area but not efficient due to a larger temperature gradient through a solution line instead of directly heating up the whole volume.

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4. Step-By-Step Procedures, using our setup as a guide - Turn on the computer, but not the acquisition program yet. - Turn on the instruments, first the patch amplifier (Axopatch-1D), then the data acquisition digitizer (Digidata 1440A). Set the amplifier in V-CLAMP mode. - Now turn on the acquisition program (Clampex). By default, the program should connect to the digitizer automatically when it was already set up before. If not, go to Configuration/Digitizer. Click Change, select the digitizer type, then click Scan. Click OK after the program finds the digitizer. The digitizer window now will show the proper digitizer, and the yellow READY light in the digitizer (Digidata 1440A) turns on. Click OK to exit the Digitizer window. - Pull electrodes from 1.5 mm diameter borosilicate glass capillaries (Sutter, BF150-86-10) with a pipette puller (Sutter P-87). We used a three-loops pull with the following settings: heat 810, pull 0, velocity 30, and time 200. The resulting pipettes had resistances of 3-5 MΩ when filled with our typical intracellular solution. Usually, increasing the heat setting will increase the pipette resistance, and vice versa. For more details, one can consult “The Pipette Cookbook” provided by the puller manufacturer (www.sutter.com). For whole-cell recording, fire-polishing of the pipette tips provides little additional benefit, therefore is not necessary. - Fill the recording chamber with extracellular solution. Put cells (coverslip) in the recording chamber, connect the ground electrode. Find the cell to be recorded. - Fill electrode with intracellular solution. Remove air bubbles inside with tapping and shaking to the electrode. - Insert the electrode into the holder and tighten. Apply a little positive air pressure to the inside of the electrode through the air pressure tubing line. Close off the air pressure tubing line to maintain 9



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the small positive pressure. The positive air pressure can be applied by mouth, or with a 3cc syringe. - Swing the manipulator into the vertical position above the cell, lower the electrode into solution. - Find the electrode tip, move it right on top of the cell, as close as possible but not touching the cell. - Open the Membrane Test program. Select Bath, then Start. The program applies a testing voltage (1 mV) to the electrode. The testing pulses produce current responses, the amplitude of which is a function of the electrode resistance (Fig. 1B1). The program also computes the responses and shows you the calculated electrode resistance. - Monitoring the Membrane Test screen, null the junction potential from the amplifier first, then lower the electrode tip into the cell membrane. The current response of the electrode will not change until it touches the membrane. - When the tip touches the cell, the current response decreases and the resistance increases (Fig. 1B2). Lower the tip further until the current response decreases to about half of the initial level, or when the tip lowering produces no further change in current response, then release electrode air pressure. A giga seal will form normally as the current response decreases to zero (Fig. 1B3). The rate of success of this step depends mostly on the health of the cells. In good, healthy rat cortical cells, the success rate can be up to 100%. In the other extreme (less healthy cells), it can be 0%. In this case, apply a small suction to form a seal. - Switch to Cell in the Membrane Test program. A -70 mV holding potential is now applied to the patch. The zero current response line remains with small capacitive transients at the onset and the end of the testing pulses. Next, break into the whole-cell by a hard, controlled suction. The capacitive component of the current response has a sudden increase (both in amplitude and in the decay phase) when this happens, reflecting a sudden increase in membrane area from a small patch 10


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to a whole-cell (Fig. 1B4). The holding current will remain relatively the same as before when the seal is maintained, while the current response shows a small increase, reflecting the leak conductance already present on the cell membrane. If the membrane patch is hard to break by suction, use the Zap function of the amplifier. The most effective way is to use a combination of light zap followed by a controlled suction. - At this stage, the Membrane Test program will compute several important values: access resistance, and membrane resistance and capacitance. With the lower resistance pipettes (3-5 MΩ) we used in our experiments, the access resistance was not a problem. It was mostly around 10-15 MΩ. For whole-cell recording, higher resistant pipettes (~10 MΩ) can be used, but make the wholecell conformation difficult to achieve. On the other hand, whole cell capacitance and series resistance might need to be compensated, but the issues are complicated. In simple terms, cell capacitance slows down the membrane potential charging step, and series resistance compromises the accuracy of membrane potential charging and reading. Therefore, it is best to keep them under control by compensating them electronically. In old amplifiers, these are done by adjusting fast and slow capacitance, and series resistance compensations. In new amplifiers, the task can be accomplished automatically by a click of button. In our experiments, we were not particularly concerned with these issues because we used low resistance electrodes to keep the series resistance error in check, and we were mainly interested in slow, steady state responses that were less sensitive to capacitance slowing. Therefore, whole-cell capacitance and series resistance compensations were not done in our recordings. - Once the whole-cell configuration is formed, several types of recording can proceed according to the experimental needs by selecting an amplifier mode. Remaining in the V-CLAMP mode, one can record the whole-cell membrane currents, either with or without changes in holding potential.

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Switching to I = 0 mode, the amplifier meter now shows the membrane potential, and one can record the resting membrane potential. Switching to I-CLAMP mode, one can record membrane potential changes in response to injection of hyperpolarizing or depolarizing currents. 5. Data Analysis We use the analysis program of the pCLAMP suite (Clampfit) for our primary data detection, data extraction, and figure export. Further analysis can be done with Microsoft Excel and other statistic and chart making programs. For membrane potential values, all-point averages of a fixed period of time (10 to 30 sec) can be used. For action potential measurements, averages of the individual measurement are used. For voltage-evoked membrane current changes, individual response or peak amplitudes of the individual responses can be averaged.

Specific Example 1: Recording Membrane Potentials of Rat Cortical Neurons Cultured rat embryonic day 18 cortical neurons were plated on poly-D-lysine-coated 10 mm glass cover slips for 4-6 days and placed inside a recording chamber (P1, Warner Instruments) that contains extracellular solution (140 mM NaCl, 2.8 mM KCl, 2 mM CaCl2, 2 mM MgCl2, 10 mM HEPES, and 10 mM D-glucose, with pH adjusted to 7.4 with NaOH). Cells were visualized with a Zeiss Axiovert 100 inverted microscope. Membrane potentials were measured at room temperature by whole-cell patch-clamp using an Axopatch 1D amplifier (Axon Instruments) in current-clamp mode. Patch electrodes were pulled from 1.5 mm diameter borosilicate glass capillaries (Sutter, BF150-86-10) with a Sutter P-87 microelectrode puller, and had 4-6 MΩ resistances when filled with an intracellular solution that contained 140 mM potassium gluconate, 10 mM NaCl, 2 mM MgCl2, 10 mM HEPES, 1 mM EGTA, 4 mM MgATP, and 0.3 mM NaGTP, with pH adjusted to 7.3 with KOH. Membrane potential data were filtered by the amplifier-incorporated 4-pole Bessel filter at 2 KHz, and digitized at 5 KHz by a Molecular Devices 12


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digitizer (Digidata 1440A) using a Dell Precision 340 computer, with pClamp 10 software (Molecular Devices). To test the functionality of the recorded neuron, depolarizing currents (5 step protocol from 100 to 500 pA, at 100 pA step) were routinely injected into the cells to elicit action potentials. We recorded resting membrane potentials from control neurons and from neurons that were treated for 24 hours with ivermectin, a chloride channel opener. The resting membrane potentials of control vs. ivermectin-treated neurons were -73.5 + 5.9 mV and -43.1 + 3.4 mV (mean + SEM, n = 12 for both groups) (Fig. 2A-C). We also measured the membrane potentials of neurons under same culture and treatment conditions with a ratiometric di-8-ANEPPS dye (Invitrogen), and found that 1% change in ANEPPS ratio corresponded to 1 mV change in resting membrane potential (Fig. 2D).22

Specific Example 2: Recording Membrane Currents of Rat Cortical and Lunds Human Mesencephalic (LUHMES) Cells Cultured rat embryonic day 18 cortical neurons and LUHMES cells23 were plated on poly-D-lysine-coated 10 mm glass cover slips 4-6 days ago and placed inside a recording chamber (P1, Warner Instruments) that contained extracellular solution (140 mM NaCl, 2.8 mM KCl, 2 mM CaCl2, 2 mM MgCl2, 10 mM HEPES, and 10 mM D-glucose, with pH adjusted to 7.4 with NaOH). Cells were visualized with a Zeiss Axiovert 100 inverted microscope. Voltage-activated membrane currents were activated by 400 ms, increasing depolarization steps (delta = 10 mV) from a holding potential of -70 mV to +60 mV, and measured at room temperature by whole-cell patch-clamp using an Axopatch 1D amplifier (Axon Instruments) in voltage-clamp mode. Patch electrodes were pulled from 1.5 mm diameter borosilicate glass capillaries (Sutter) with a Sutter P-87 microelectrode puller, and had 4-6 MΩ resistances when filled with an intracellular solution that contained 140 mM potassium gluconate, 10 mM NaCl, 2 mM MgCl2, 10 mM HEPES, 1 mM EGTA, 4 mM MgATP, and 0.3 mM NaGTP, with pH adjusted to 7.3 with KOH. 13



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Membrane currents were filtered by the amplifier-incorporated 4-pole Bessel filter at 2 KHz, and digitized at 5 KHz by a Molecular Devices digitizer (Digidata 1440A) using a Dell Precision 340 computer, with pClamp 10 software (Molecular Devices). Both inward and outward whole-cell membrane currents were recorded. In cortical neurons, there were robust inward and outward currents that reached nA peak amplitude (Fig. 3A). The inward currents were a mixture of Na and Ca current, with the bulk of peak current responses representing mostly the Na current. Similarly, the outward K currents likely represent a mixture of multiple subtypes. The total peak inward and outward currents were measured, and we did not attempt to further examine them. In contrast, the LUHMES cells had different inward and outward current profiles. In undifferentiated LUHMES cells, the inward currents were close to zero and the outward currents were very small and appeared to be of the delayed activated type only (Fig. 3B). In differentiated LUHMES cells (40 ng/mL basic Fibroblast Growth Factor, 1 ug/mL tetracycline, 1 mM dibutyryl cAMP & 2ng/mL Glial Derived Neurotrophic Factor)23, the inward currents started to develop, and the outward currents became larger while maintaining the delayed activation characteristic (Fig. 3C).

Extracellular Field Potential (EFP) Recording In contrast to whole-cell patch clamp recording, extracellular field potential recording is simpler, and has been widely used in many in vivo and in vitro scientific experimentations, as well as in many clinical settings.11,24-27 Well-known examples include Single and Multiple Unit Recording (SUR and MUR), Local Field Potential (LFP) recording, Electroencephalogram (EEG), Electrocorticogram (ECoG), Electrocardiogram (EKG), and Electromyogram (EMG). In principle, an electrode is placed in an extracellular space and detects potential differences at its tip vs. a reference electrode. When the reference electrode is sufficiently far away and isolated from the local electrical activities, the recording electrode detects the potential changes at its location, referred to as a monopolar or direct recording. 14


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When the reference electrode is close to the recording electrode, the recording picks up only potential differences between them at the recording site, since both of them have more or less similar potential changes arising from activities of the surrounding areas, which cancel each other out between them. This is referred as bipolar or differential recording. The potential at the tip of the electrode is the summation of all electrical activities, near and far, fast and slow, including action potentials, synaptic and non-synaptic potentials due to receptor activations, potentials from voltage- and messenger-gated ion channel openings and closings, transporter/pump-generated potentials, and pace-making potentials. The close and nearby activities dominate the recorded potential changes while the contribution of farther away activities decreases with distance. The filtering effect of the intra- and extra-cellular spaces makes the fast spiking event recording local, giving rise to single- and multi-unit recordings. The slow potential changes on the other hand can be recorded from farther away because they are affected much less by the space filtering. But the slow potential changes often cancel each other out when they occur randomly. When they happen in a coordinated fashion such as in synaptic activations within a densely packed dendritic field, the synchronized synaptic responses can be readily recorded, referred to customarily as Local Field Potential (LFP). Although extracellular electrodes record field potential changes that are local, we will call it extracellular field potentials (EFP), so as not to be mixed up with the LFP that has already been in use by neuroscientists for decades to describe specifically the postsynaptic potentials. A flexible or rigid microelectrode, made with metal or pulled from glass micropipette, is placed close to or inside the tissue to be recorded (Fig. 4A). The number of electrodes is limited only by the availability of the input channels of the amplifiers. One can record from any tissue that is capable of generating electrical activity, such as whole organisms, organs, tissue slices, and tissue cultures. Electrical potential signals around the electrode tip are picked up either directly against a common ground placed at a distance from the site, or differentially against a nearby electrode, and amplified by low noise voltage 15



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amplifiers. For high impedance glass microelectrodes, a matching high impedance preamplifier is required. The inflexibility and requirement for the additional instrument for the glass microelectrode nonetheless make the system more sensitive to the electrical potential changes around the electrode tips, enabling better detection of local electrical activity. The amplified signals can be conditioned through filtering depending on experimental needs. For example, to detect slow field potential changes, a low low-pass filter combined with a low high-pass filter is used to filter out the high frequency components of the signals. On the other hand, a high low-pass filter in combination with a high highpass filter can be used to smooth out the slow potential changes and to detect high frequency events such as action potentials. In the context of biomedical and tissue engineering, the slower synaptic potentials are usually not well organized and coordinated due to the lack of the structural development and organization, we thus focus on detecting the fast spiking activities in the tissue (Fig. 4B&C).

Practical Guides for EFP Recording 1. Basic Recording Set-Up Compared to whole-cell patch clamp recordings, the basic recording set-up for extracellular field potential recording is much simpler. A vibration-free table such as a TMC or Newport air table is good to have, but can be substituted with any stable table top. A fine manipulator on top of coarse movement control is normally sufficient for positioning the recording electrode on or inside the cell preparations. The visual-optical component usually consists of a stereo dissecting microscope with zooming or stepping amplification options. The electrical component is similar to the patch clamp set-up, and consists of an electrical noise shielding mechanism such as a Faraday cage, an extracellular potential amplifier such as a WPI or a NPI amplifier, and a data acquisition system that includes a digitizer such as the Digidata, acquisition software such as the pClamp software suite, and 16


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a computer. The set-up is usually built like a tower on top of an air table, with the dissecting microscope as the center piece. One or multiple manipulators and amplifier headstages are positioned around the optical center of the microscope. The headstage has connections for the electrode holder and the signal ground. The ground can be any chlorided silver wire (see set-up for whole-cell recording for details). The amplifier, the data acquisition system, and other instrument such as stimulator are positioned outside of the Faraday cage, often housed in a rig to further shield any interference these instruments may produce from affecting the recording in the cage. 2. Solutions In extracellular field recording configuration, the recording solutions usually mimic the extracellular ionic conditions, meaning high Na+ and low K+ in the solution. A common extracellular solution that we use in our recording consists of the following components in mM: NaCl 140, KCl 2.8, CaCl2 2, MgCl2 2, HEPES 10, and D-glucose 10, with pH adjusted to 7.4 with NaOH (for extracellular solution). The solution is used to bathe the preparations, as well as to fill the recording electrode. 3. Environment Control-Temperature Control Similar to the whole-cell patch clamp recording, environment control in extracellular field recording consists mainly of solution exchange control and temperature control, but in a simpler way due to the nonspecific and slow nature of the technique. Solution exchange can be done manually or by a gravity perfusion setup. Temperature control can be achieved by heating around and within the recording chamber, or by heating the inflow solution. 4. Step-By-Step Procedures, using our setup as a guide - Turn on the computer, but not the acquisition program yet. - Turn on the instruments, first the data acquisition digitizer (Digidata 1550), then the A-M Systems 17



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Differential AC amplifier (Model 1700). The bridge amplifier (BA-03X) can be turned on now, or after the electrode is inserted into the solution. The Model 1700 amplifier was connected to the BA-03X in series (BA-03X output to 1700 input) to provide additional amplification and to convert the BA03X’s DC signal into a differential one. With a high impedance headstage, model 1700 can be used by itself as well. - Now turn on the acquisition program (Clampex). By default, the program should connect to the digitizer automatically when it was already set up before. If not, go to Configuration/Digitizer. Click Change, select the digitizer type, then click Scan. Click OK after the program finds the digitizer. The digitizer window now will show the digitizer and the digitizer yellow READY light turns on. Click OK to exit the Digitizer window. - Fill the recording chamber with extracellular solution. Place the preparation in the chamber. Locate the recording site. - Pull electrodes from 1.5 mm diameter borosilicate glass capillaries (Sutter, BF150-86-10) with a pipette puller (Sutter P-87). We used a single pull with the following settings: heat 815, pull 60, velocity 85, time 150. When filled with our standard extracellular solution, the electrodes had a resistance of 80-100 MΩ. Increasing heat setting will increase the electrode resistance, and vice versa. - Fill the electrode with extracellular solution. Remove air bubbles inside with tapping and shaking of the electrode. Insert electrode into the holder and tighten. - Swing out the manipulator, insert the electrode holder into the amplifier headstage. Return the manipulator into the center position. Lock the manipulator, lower the electrode into solution. - Turn on the bridge amplifier if it is not on yet. Find electrode tip, move it right on top of the 18


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recording site. Test the electrode resistance. Move the electrode into the preparation as close as possible, but not touching. The extracellular field recording is now ready to proceed. 5. Data Analysis We use the analysis program of the pCLAMP suite (Clampfit) for our primary data detection, data extraction, and figure export. To detect spikes, we use Clampfit’s Event Detection function. One time peak-to-peak baseline noise amplitude is used as the threshold for spike detection. Analysis of detected spikes are done in Clampfit. Further analysis can be done with Microsoft Excel and other statistic and chart making programs. Spike frequency data are usually expressed as Hz or number of spikes per 20 seconds.

Specific Example 3: Recording Spiking Activity of Rat Cortical Neurons in Cultures and in 3D Silk Sponges Dissociated day 18 rat embryonic cortical neurons were seeded on poly-D-lysine-coated 35-mm Corning Culture Dishes or in 3D silk protein-based scaffolds. After a period of incubation (10 days to 3 months), the electrical spiking activities of the neurons in cultures or in 3D scaffolds were recorded extracellularly at room temperature by sharp glass microelectrodes using an NPI Bridge Amplifier (BA-03X, Tamm, Germany). Cells in cultures or in 3D scaffolds were immersed in extracellular solution (140 mM NaCl, 2.8 mM KCl, 2 mM CaCl2, 2 mM MgCl2, 10 mM HEPES, and 10 mM D-glucose, with pH adjusted to 7.4 with NaOH). Sharp glass microelectrodes were pulled using Sutter Borosilicate Glass (BF150-86-10) with a Sutter Micropipette Puller (P-87). The microelectrodes were filled with extracellular solution and had a resistance of 60-80 MΩ. The cell cultures and 3D scaffolds were visualized with a Wild M3C Dissecting Microscope (Switzerland). The recording electrodes were positioned close to the cultured tissues or the silk sponges. Fast field potential changes (spikes) were recorded with the NPI amplifier at a bandwidth of 19



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0.3-10 kHz and were further amplified with an A-M Systems Differential AC amplifier (Model 1700) to a combined total gain of 10,000x. The signals were digitized at 10 KHz by a Molecular Devices digitizer (Digidata 1550) using a Dell Optiplex GX620 computer with pClamp 10 software (Molecular Devices). In 2 weeks long 3D cultures, frequent spikes were recorded (Fig. 5A1). Spikes were defined as sharp changes in field potential in sub millisecond ranges, and were counted as events when they crossed the detection threshold set at one time of the average peak-to-peak amplitude of baseline noise. The peakto-peak baseline noises of the recordings were between 0.25 to 0.3 mV, and the detection thresholds were set accordingly between 0.25 to 0.3 mV. The directions of the spikes could be up or down, or both. The amplitudes of the events were in a continuing scale from the detection threshold (0.25-0.3 mV) to the data acquisition's max amplitude of 1 mV, due to our amplification factor of x10,000. The spikes were blocked by neuronal sodium channel blocker tetrodotoxin (TTX, 10 µM) (Fig. 5A2), demonstrating that the spikes resulted from neuronal sodium channel firings. In another set of experiments, we tested the effects of extracellular matrix (ECM) and astrocyte-secreted matricellular proteins such as SPARC (secreted protein, acidic and rich in cysteine), Hevin, TSP2 (thrombospondin 2) on neuronal activity in 2 week long 3D collagen-based cultures. Matricellular proteins appeared to enhance the neuronal spikes in 25% of preparations (Fig. 5B).28

Specific Example 4: Recording Spontaneous Spiking Activity and Evoked Synaptic Potentials of iPSCs in 3D Silk Sponges Different stages of induced pluripotent stem cells (iPSCs) were seeded in 3D silk protein-based scaffolds. After a period of incubation (10 days to 3 months), the electrical spiking activities of the neurons in 3D scaffolds were recorded extracellularly at room temperature by sharp glass microelectrodes using an NPI Bridge Amplifier (BA-03X, Tamm, Germany). Cells in 3D scaffolds were immersed in extracellular

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solution (140 mM NaCl, 2.8 mM KCl, 2 mM CaCl2, 2 mM MgCl2, 10 mM HEPES, and 10 mM D-glucose, with pH adjusted to 7.4 with NaOH). Sharp glass microelectrodes were pulled using Sutter Borosilicate Glass (BF150-86-10) with a Sutter Micropipette Puller (P-87). The microelectrodes were filled with extracellular solution and had a resistance of 60-80 MΩ. The 3D scaffolds were visualized with a Wild M3C Dissecting Microscope (Switzerland). The recording electrodes were positioned close to the silk sponges. Fast field potential changes (spikes) were recorded with the NPI amplifier at a bandwidth of 0.3-10 kHz and were further amplified with an A-M Systems Differential AC amplifier (Model 1700) to a combined total gain of 10,000x. The signals were digitized at 10 KHz by a Molecular Devices digitizer (Digidata 1550) using a Dell Optiplex GX620 computer with pClamp 10 software (Molecular Devices). Spikes were defined and detected as described above in Specific Example 3. Electrical stimulations (40 V, 1 ms pulses) of the cultured tissues were generated via a Grass S44 Stimulator, passed through a Grass Stimulus Isolation Unit (SIU5), and delivered through a platinum parallel bipolar electrode (FHC PBSB0875) with a distance of 800 μm between two tips positioned near the recording electrode. In 12 weeks old preparations, many spikes were present (Fig. 6A&B, upper panels). Application of a cocktail of glutamate receptor blockers (6-cyano-7-nitroquinoxaline-2,3-dione (CNQX) 50 μM, (2R)-amino-5phosphonopentanoate (AP5) 250 μM) reduced the presence of the neuronal spikes (Fig. 6A, lower panel), indicating that many neuronal spikes resulted from activation of glutamate receptors. When these preparations were subjected to controlled cortical impact (CCI) like injury (adapted from Yang et al.29) (in brief a 3mm flat tip impounder impacted the scaffold at 6m/s for 100ms duration to a depth of 0.6mm), the neuronal spiking activity was no longer detected 24 hours after (Fig. 6B, lower panel). In 812 week old preparations, we also examined the neuronal responses evoked by electrical stimulation (Fig. 7). Here, 30 extracellular field potential responses to the stimulation pulses (40V, 1 ms, at 0.1 Hz) were first recorded (Fig. 7A). These responses comprised both the electrical field changes resulting from the stimulations themselves, and the evoked neuronal responses. To dissect out the neuronal 21



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responses, we used the neuronal sodium channel blocker TTX (5 μM). Five min after TTX was applied, another 30 extracellular field potential responses to the same stimulation pulses were recorded (Fig. 7B), and averaged digitally (Fig. 7C). This average response (TTX average) represented the direct field changes resulting from the stimulations. We then subtracted the TTX average response from the first 30 stimulation responses digitally. This resulted in the extraction of the evoked neuronal responses that appeared to comprise a direct neuronal spike firing and a delayed, slower potential response that likely represented the evoked synaptic potentials (Fig. 7D). It is unlikely that the spikes are an artifact from the TTX average subtractions, mainly because the spikes can be absent, or can go up, or down, or up and down, or down and up, or can be small or large. This variability looks to be a biological fact rather than an artifact.

Power Spectrum Considerations (Analysis) Signals from extracellular field potential recordings are very sensitive to filter settings. When not filtered, the signals are dominated by high frequency baseline noises. When the high-pass and the low-pass filters are set at a low frequency range, slow potential fluctuations like those of LFPs can be detected. When the high-pass and the low-pass filters are set at a high frequency range, only the high frequency components of the signals will be detected. In our experimental conditions, the well-organized synaptic structures that produces LFPs are likely not present. We therefore set our filter setting on the high frequency side (a bandwidth of 0.3-10 kHz) to record neuronal spikes. A relevant question pertaining to experimental needs is that besides the obvious spikes as detected by our criteria, whether the baseline noises contain additional information about the underlying neuronal activity within the 3D preparations. To this end, we employed power spectrum analysis to determine the frequency properties of our recorded signals under various conditions. We first tested whether neuronal activity blocker TTX 22


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affected the power spectra of the baseline with or without the presence of neuronal spikes. The presence of TTX did not affect either the baseline or the power spectra (Fig. 8A). When neuronal spiking activity was present, the power spectrum was not different from when the baseline was without spikes (Fig. 8B). TTX blocked the neuronal spiking activity, but did not affect the power spectra (Fig. 8C). We next tested whether the power spectra in the presence of neuronal spikes would be affected by a cocktail of glutamate receptor blockers that block the neuronal spiking activity. Here again, glutamate blockers blocked the neuronal spiking activities, demonstrating that the neuronal spikes depend on glutamatergic receptor activation. But the power spectra were not changed by the actions of the receptor blockers (Fig. 8D). Taken together, these results show that the baseline under our experimental conditions does not contain neural activity information. They also show that, when neuronal spiking activity is not at a high frequency, sustained firing level as in the whole brain, its contribution to the power spectra is negligible, and that the power spectra alone would not be a useful index for neuronal activities in experiments under the same or similar conditions.

CONCLUSIONS Electrophysiology is a powerful and valuable tool for functional assessments in tissue engineering studies. Whole-cell patch clamp recording of excitable and nonexcitable cells can provide functional determination of the cell properties such as resting membrane potential, action potentials, and membrane ion channel expression. Extracellular field potential recordings can be used to evaluate network neuronal activities within artificial 3D constructs. When combined with field stimulation, extracellular field potential recording can yield information on network synaptic interactions.

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ASSOCIATED CONTENT The following supporting material is available free of charge: EPhysVideo.mp4 (a brief video demonstration of the electrophysiological techniques presented in this tutorial)

ACKNOWLEDGEMENTS The authors thank Prof. Barry Trimmer for providing laboratory facility and some of the electrophysiological recording equipment, Amy Thurber for helping with early initial setting up of the equipment, and Dr. Nurdan Oezkucur for preparing cell cultures used in Specific Example 1. Will Collins, Will Cantley, and Disha Sood contributed equally to the study. The project was supported by the NIH (AR005593, AR061988, R01NS092847), the Keck Foundation, and the Allen Center.

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REFERENCES (1) Hodgkin, A.L.; Huxley, A.F. Action potentials recorded from inside a nerve fibre. Nature 1939, 144, 710-711. (2) Brazier, M.A. The electrical activity of the nervous system: Electrical signals are the neurophysiologist’s clue to coding in the nervous system. Science 1964, 146, 1423-1428. (3) Eccle, J.C. The ionic mechanisms of excitatory and inhibitory synaptic action. Ann N Y Acad. Sci. 1966, 137, 473-94. (4) Coraboeuf, E. Ionic basis of electrical activity in cardiac tissues. Am. J. Physiol. 1978, 234, H101-116. (5) Neher, E.; Sakmann, B. Single-channel currents recorded from membrane of denervated frog muscle fibres. Nature 1976, 260, 799-802. (6) Mackinnon, R. Using mutagenesis to study potassium channel mechanisms. J. Bioenerg. Biomembr. 1991, 23, 647-663. (7) Brown, K.T.; Flaming, D.G. Advanced Micropipette Techniques for Cell Physiology. Willey, New York, 1986. (8) Rudy, B.; Iverson, L.E. Methods in Enzymology. Vol. 207: “Ion Channels.” Academic Press, San Diego, 1992. (9) Sakmann, B.; Neher, E. Single Channel Recording. 2nd ed. Plenum, New York, 1995. (10) Hille, B. Ion Channels of Excitable Membranes. 3rd ed. Sinauer Associates, Sunderland, MA, 2001. (11) Vertes, R.P.; Stackman, R.W. Jr. Electrophysiological Recording Techniques. Springer Protocols, Neuromethods Series Nr. 54, Humana Press, New York, 2011. (12) Verkhratsky, A.; Parpura, V. History of electrophysiology and the patch clamp. Methods Mol. Biol. 2014, 1183, 1-19. (13) Ebner, T.J.; Chen, G. Use of voltage-sensitive dyes and optical recordings in the central nervous system. Prog. Neurobiol. 1995, 46, 463-506.

(14) Baker, B.J.; Kosmidis, E.K.; Vucinic, D.; Falk, C.X.; Cohen, L.B.; Djurisic M, Zecevic D. Imaging brain activity with voltage- and calcium-sensitive dyes. Cell. Mol. Neurobiol. 2005, 25, 245-282. (15) Kralj, J.M.; Douglass, A.D.; Hochbaum, D.R.; Maclaurin, D.; Cohen, A.E. Optical recording of action potentials in mammalian neurons using a microbial rhodopsin. Nat. Methods 2011, 9, 90-95. (16) Adams, D.S.; Levin, M. General principles for measuring resting membrane potential and ion concentration using fluorescent bioelectricity reporters. Cold Spring Harbor Protoc. 2012, 2012, 385-397. (17) Jin, L.; Han, Z.; Platisa, J.; Wooltorton, J.R.; Cohen, LB.; Pieribone, V.A. Single action potentials and subthreshold electrical events imaged in neurons with a fluorescent protein voltage probe. Neuron 2012, 75, 779-785. 25



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(18) Lippert, M.T.; Takagaki, K.; Xu, W.; Huang, X.; Wu, J.Y. Methods for voltage-sensitive dye imaging of rat cortical activity with high signal-to-noise ratio. J Neurophysiol. 2007, 98, 502-512. (19) Carlson, G.C.; Coulter, D.A. In vitro functional imaging in brain slices using fast voltage-sensitive dye imaging combined with whole-cell patch recording. Nat. Protoc. 2008, 3, 249-255. (20) Adams, D.S.; Levin, M. Measuring resting membrane potential using the fluorescent voltage reporters DiBAC4(3) and CC2-DMPE. Cold Spring Harbor Protoc. 2012, 2012, 459-464. (21) Hamill, O.P.; Marty, A.; Neher, E.; Sakmann, B.; Sigworth, F.J. Improved patch-clamp techniques for high-resolution current recording from cells and cell-free membrane patches. Pflugers Arch. 1981, 391, 85-100. (22) Ozkucur, N.; Quinn, K.P.; Pang, J.C.; Du, C.; Georgakoudi, I.; Miller, E.; Levin, M.; Kaplan, D.L. Membrane potential depolarization causes alterations in neuron arrangement and connectivity in cocultures. Brain Behav. 2015, 5, 24-38. (23) Lotharius, J.; Barg, S.; Wiekop, P.; Lundberg, C.; Raymon, H.K.; Brundin, P. Effect of mutant αsynuclein on dopamine homeostasis in a new human mesencephalic cell line. J. Biol. Chem. 2002, 277, 38884-38894. (24) Bukalo, O.; Dityatev, A. Analysis of neural cell functions in gene knockout mice: electrophysiological investigation of synaptic plasticity in acute hippocampal slices. Methods Enzymol. 2006, 417, 52-66. (25) Chorev, E.; Epsztein, J.; Houweling, A.R.; Lee, A.K.; Brecht, M. Electrophysiological recordings from behaving animals--going beyond spikes. Curr. Opin. Neurobiol. 2009, 19, 513-519. (26) Buzsaki, G.; Anastassiou, C.A.; Koch, C. The origin of extracellular fields and currents--EEG, ECoG, LFP and spikes. Nat. Rev. Neurosci. 2012, 13, 407-420. (27) Saffitz, J.E.; Corradi, D. The electrical heart: 25 years of discovery in cardiac electrophysiology, arrhythmias and sudden death. Cardiovasc. Pathol. 2016, 25, 149-157. (28) Sood, D.; Chwalek, K.; Stuntz, E.; Pouli, D.; Du, C.; Tang-Schomer, M.; Georgakoudi, I.; Black, L.; Kaplan, D. L. (2016). Fetal brain extracellular matrix boosts neuronal network formation in 3D bioengineered model of cortical brain tissue. ACS Biomater. Sci. Eng. 2016, 2, 131-140. (29) Yang, J.; You, Z.; Kim, H.H.; Hwang, S.K.; Khuman, J.; Guo, S.; Lo, E.H.; Whalen, M. J. Genetic analysis of the role of tumor necrosis factor receptors in functional outcome after traumatic brain injury in mice. J. Neurotrauma 2010, 27, 1037-1046.

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FIGURE LEGENDS Fig. 1: Basics of whole-cell patch clamp recording. A: Photogragh of a patch clamp recording setup, showing some of the main components. B: Schematic process of obtaining a whole-cell patch clamp recording. B1: An electrode is immersed into the solution. A voltage testing pulse is applied to the electrode, producing a current response inversely proportional to the resistance of the electrode. B2: The electrode touches and is pressed onto the cell. The test pulse responses become smaller as the electrode dipped into the cell membrane. B3: A slight suction reduces the testing pulse responses to a near straight line, indicating the formation of a gigaohms seal between the electrode and the membrane patch. The small transients at the beginning and at the end of the test pulses are capacitant transients of the membrane patch. B4: A further suction breaks the membrane patch, resulting in a whole-cell patch clamp. The transients at the beginning and at the end of the test pulses increase greatly due to a sudden increase in membrane area from a tinny patch to a whole-cell.

Fig. 2: Recording of resting membrane potential from rat cortical neurons. Reproduced with permission from ref. 22 Copyright 2014 Ozkucur, N.; Quinn, K.P.; Pang, J.C.; Du, C.; Georgakoudi, I.; Miller, E.; Levin, M.; Kaplan, D.L. Brain and Behavior published by Wiley Periodicals, Inc. A&B: Resting membrane potential of cortical neurons in control and 24 hrs after ivermectin treatment. C: Effect of ivermectin treatment on resting membrane potential. ** indicates statically significant different (n = 12). D: Effect of ivermectin treatment on the ratiometric measurement of a voltage sensitive dye (di-8-ANEPPS) in rat cortical neurons.

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Fig. 3: Recording of voltage-activated membrane currents from rat cortical neurons and LUHMES cells. Membrane currents were activated by increasing depolarizing voltage steps (delta = 10 mV, from -70 to +60 mV, on top of the traces). Current responses were leak-subtracted. On the left were full current responses to the voltage protocol. On the right were expansions in time scale (25x) showing the initial inward and outward current responses. Rat cortical neurons expressed robust inward and complex outward currents (A). Undifferentiated LUHMES cells had no inward and very small simple outward currents (B), while differentiated LUHMES cells started to express more inward and outward currents (C).

Fig. 4: Recording of extracellular field potential from 3D neuronal preparations. A: Schematic diagram and photograph showing the position of electrode in the extracellular space within the 3D preparations. The photograph also showed the position of bipolar stimulating electrodes. B: Example of recorded spiking activity. C: Expanded time scale (1000x) showing details of one recorded spike denoted by * in B.

Fig. 5: Neuronal spikes in 3D preparations of rat cortical neurons. A: Spiking activities before (A1) and in the presence of TTX (A2). Spiking activities were all blocked, demonstrating their neuronal origin. On the right are corresponding frequency plots. B: Effects of fetal extracellular matrix and astrocyte-secreted matricellular proteins (SPARC, Hevin, & TSP2, see text) on neuronal spiking activity. Matricellular proteins produced a robust enhancement of the neuronal spikes in 25% of the preparations.

Fig. 6: Neuronal spikes in 3D preparations of induced pluripotent stem cells. A: Example traces before (A1) and in the presence of glutamate receptor antagonists (CNQX&AP5, A2). Most spikes were blocked by the glutamate receptor antagonists, demonstrating that they were dependent on glutamate receptor 28


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activation. B: Spikes present in control preparations (B1) were absent in preparations 24 hrs after being subject to concoction injury (B2).

Fig. 7: Evoked neuronal responses in 3D preparations of iPSCs. A: Potential responses were recorded and aligned in response to 1 ms stimulation (40 V). The horizontal bars in each panel indicate when the stimuli were applied. B: After recording A, the same responses were recorded in the presence of TTX, eliminating the neural component from A. C: The B responses were digitally averaged, resulting in a “TTX Average.” D: Evoked responses in A were subtracted digitally by TTX Average. The resulting traces represent the TTX-sensitive component of the evoked responses, likely comprising of fast evoked spikes and slower synaptic potentials. The vertical bar in C represents 0.5 mV and applies to all panels.

Fig. 8: Power spectral analysis of baseline and spiking activity in 3D neural preparations. In each panel, 60 sec long recorded traces under two conditions are presented on the left, and the corresponding power spectra are on the right. The bandwidth of the recordings was set at 0.3-10 kHz. The vertical scale bar in A represents 0.5 mV and applies to all traces. A: Baseline noise with or without the presence of TTX. TTX did not visibly change the baseline nor changed the power spectrum profile. B: The presence of spikes did not alter the power spectrum vs. that of the baseline noise. C: When spikes were blocked by TTX, the power spectra were not different before vs. after TTX. D: When spikes were blocked by glutamate receptor antagonists CNQX&AP5, the power spectra were not different before vs. after the drugs.

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1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42

Fig. 7 A. Evoked Potentials

B. Evoked Potentials + TTX

Stimuli

Stimuli

D. Evoked Potentials - TTX Average

C. TTX Average

Evoked Spikes

Synaptic Potentials

Stimuli

Stimuli


Page 36 of 39

Page 37 of 39

1e-5

Baseline

Baseline

1e-6

1e-6

+ TTX

+ Spikes

1e-7

+ TTX

1e-8 1

C Control

+ TTX

10

100

1

D Control Control + TTX

1

10

100

1000

Frequency (Hz)


10

100

1000

Frequency (Hz)

1e-5 1e-6

+ CNQX/AP5

1e-8

+ Spikes

1e-8

1000

1e-7

Baseline

1e-7

Frequency (Hz)

1e-5 1e-6

1e-5

Amplitude (mV² / Hz)

Baseline

B


A


Fig. 8


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42


Control

1e-7

+ CNQX/ AP5

1e-8 1

10

100

1000

Frequency (Hz)


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48

For Table of Contents Use Only


Page 38 of 39

Page 39 of 39

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For Table of Contents Use Only

Tutorials for Electrophysiological Recordings in Neuronal Tissue Engineering Du, Chuang; Collins, Will; Cantley, Will; Sood, Disha; Kaplan, David


Tutorials for Electrophysiological Recordings in ... - ACS Publications

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