This work was supported by grants from the German Research Foundation (Si 1747/1-1), the Else Kr&#246;ner-Fresenius-Stiftung, and the Deutsche Stiftung f&#252;r Herzforschung.

1. Preparation of iPSC-derived Cardiomyocytes for Imaging
NOTE: Methods for iPSC culture and cardiac differentiation have been published before12,13,14 and are not discussed here in detail. The purification of iPSC-CMs by manual microdissection, magnetic cell separation, or lactate selection is recommended, depending on the differentiation protocol used. For the following protocol, microdissected explants from beating areas, generated using a monolayer differentiation protocol13, were taken on day 15 of a cardiac differentiation and cultured until day 30 on fibronectin-coated plates as described before10.
<ol>
	<li>Take the cell culture plates out of the incubator and manually collect the cardiac explants into microcentrifuge tubes using a 200 &#181;L pipette. After sedimentation, aspirate the cell culture supernatant and gently wash the explants 2x with Hank&#8217;s balanced salt solution (HBSS).</li>
	<li>Dissociate iPSC-CMs by adding collagenase type II (430 U/mL in HBSS) and incubate them at 37 &#176;C for 30 min.</li>
	<li>After the sedimentation of the remaining large cell clumps, collect the supernatant that contains the dissociated single iPSC-CMs and add this to fresh CM maintenance medium containing a high fetal bovine serum (FBS) concentration (DMEM-F12, 20% FBS, 2 mM L-glutamine, 1:100 MEM non-essential amino acids, 100 U/mL penicillin-streptomycin, and 0.1 mM &#946;-mercaptoethanol).</li>
	<li>Repeat the dissociation if large cell clumps remain. Collect single iPSC-CMs by centrifugation and resuspend them in CM medium containing a low FBS concentration (2% FBS).</li>
	<li>Re-seed single iPSC-CMs in a density allowing the subsequent imaging of single cells on a 3.5 cm glass-bottom cell culture microdish that has been coated with fibronectin (2 &#181;g/cm2). Optimize the seeding density before engaging in high-throughput experiments. Usually, 5&#8211;10 cardiac explants per 3.5 cm dish are sufficient; however, this heavily depends on the explant size and purity.</li>
	<li>Culture the single iPSC-CMs after their dissociation in CM maintenance medium containing 2% FBS for at least 48 h to ensure a sufficient recovery. 
	Note: In the next steps, the iPSC-CMs are transduced with a lentivirus encoding the GEVI VSFP-CR under the control of an MLC2v enhancer10 to achieve a ventricular-specific GEVI expression. Alternatively, lentiviral constructs allowing to express the GEVI specifically in nodal- or atrial-like cells, or an unspecific construct harboring the ubiquitously-expressed PGK promoter, can be used10. Lentiviral plasmids are available upon request. 
	CAUTION: When working with lentiviral particles, ensure that the working environment has the adequate biosafety level, adhere to the safety instructions for working with potentially infectious agents, and take the necessary safety precautions.</li>
	<li>Prepare the infection medium by mixing the lentivirus10-containing cell culture supernatant, which has been produced in HEK293 cells as described before15, with CMs maintenance medium (see step 1.3) in a 1:1 ratio.</li>
	<li>Add hexadimethrine bromide at a final concentration of 8 &#181;g/mL to enhance the infection efficiency. 
	NOTE: Due to the high infection efficiency of iPSC-CMs prior concentration of lentivirus through ultracentrifugation is usually not necessary.</li>
	<li>Aspirate the CM maintenance medium from the single iPSC-CMs and replace it with the infection medium prepared during steps 1.7 and 1.8. Culture the infected single iPSC-CMs in the infection medium for 12 h at 37 &#176;C. Then, aspirate the medium and replace it again with CM maintenance medium. 
	NOTE: A fluorescence signal from the GEVI appears 48 h after the infection. An imaging of the membrane potential recordings is recommended, 72 h after infection at the earliest, to ensure a proper signal strength. Single infected iPSC-CMs can be cultured for at least 3 weeks and imaged sequentially. If extensive photobleaching occurs during an imaging session, the fluorescence signal recovers over time.</li>
	<li>Before imaging, exchange the cell culture medium by Tyrode&#8217;s solution supplemented with Ca2+ (135 mM NaCl, 5.4 mM KCl, 1 mM MgCl2, 10 mM glucose, 1.8 mM CaCl2, and 10 mM HEPES; pH 7.35).</li>
</ol>
2. Optical Membrane Potential Recordings
<ol>
	<li>Place the imaging chamber containing the iPSC-CMs on the stage of an inverted epifluorescence microscope equipped with an image splitter, the appropriate filter sets, and a camera (e.g., an sCMOS camera) capable of high-speed imaging at a high sensitivity. 
	NOTE: For example, use a 480/40 nm band-pass excitation filter combined with a 500 nm long-pass dichroic mirror in the microscope, and a 568 nm long-pass dichroic mirror combined with 520/28 nm and 630/75 nm band-pass emission filters in the image splitter. For single-cell imaging, a high-magnification, high-numerical-aperture oil immersion objective provides the best signal-to-noise ratio.</li>
	<li>If electrical pacing is required, install pacing electrodes in the imaging chamber and connect them to a stimulus generator. 
	NOTE: We used a pacing inset that contained two platinum electrodes and fitted into the 3.5 cm glass-bottom cell culture dishes. The cells located between the electrodes were imaged. Typical pacing parameters are rectangular pulses of 5 ms in duration, with an amplitude of 10 V/cm electrode distance.</li>
	<li>Optionally, use a heated microscope stage to ensure a stable temperature of the iPSC-CMs during imaging, or even a microscope incubation chamber set-up if long-term imaging is intended.</li>
	<li>Bring the cells to focus and place a cell expressing the GEVI into the center of the viewing field. 
	NOTE: This is most conveniently done by using the eyepieces of the microscope using a green fluorescent protein (GFP) or a red fluorescent protein (RFP) filter set to identify cells expressing the GEVI.</li>
	<li>Set the optical path so that emitted light is routed to the image splitter. Switch the filter cube in the microscope to the one to be combined with the filters in the image splitter (see Table of Materials).</li>
	<li>In the software controlling the camera, set the acquisition settings so that high-speed imaging (e.g., 100 frames per second, with an exposure time of 10 ms per frame) is possible. 
	Note: Usually, this involves the binning of camera pixels (e.g., 8 x 8 pixels of the camera chip are binned to generate one pixel in the time-lapse movie).</li>
	<li>Set up the image splitter according to the manufacturer&#8217;s instruction. The two images representing the two different emission wavelength bands should be adjacent to each other, each occupying one half of the image. 
	NOTE: This step has to be performed only 1x at the beginning of each imaging session.</li>
	<li>Check and, if necessary, adjust the focus of the image produced by the camera.</li>
	<li>Adjust the brightness of the illumination light so that any saturation of the pixels is avoided; this can be most easily done by using a color palette in which saturated pixels are represented by a unique color. 
	NOTE: During all these preparations prior to imaging, limit the exposure of the sample to the excitation light to the necessary amount (i.e., do not leave the light on for extended periods of time) to avoid photobleaching of the GEVI.</li>
	<li>Set up the acquisition of the time series (e.g., 5,000 images at 100 frames per second for a series duration of 50 s). Dim the light in the room (including computer monitors) to avoid stray light influencing the measurement. If the excitation light shutter is not controlled by the imaging software, open it just before starting the acquisition.</li>
	<li>Start the acquisition of the image series. If electrical pacing is desired, start the pacing sequence at the appropriate time point, unless it is automatically initiated by the imaging software. If the application of a pharmacological agent is wanted, do this either manually (e.g., by pipetting 100 &#181;L of the agent in a tenfold end concentration to the recording chamber containing 900 &#181;L of buffer, and gently mixing it with the pipette) or by using a constant-flow perfusion system.</li>
	<li>When the acquisition is finished, close the excitation light shutter if necessary. Save the recording to the hard drive.</li>
	<li>Leaving the recording settings (i.e., exposure time, frame rate, and light intensity) unchanged, perform a recording of the background fluorescence (as in step 2.11) over a region of the coverslip not containing cells or over another coverslip on which no cells have been seeded. 
	NOTE: If sequential measurements are performed without changing the acquisition settings, only one background fluorescence recording is necessary for all these measurements.</li>
	<li>If desired, perform additional experiments (steps 2.4&#8211;2.13) on different iPSC-CMs located on the same coverslip. Be aware that all CMs on the coverslip will be affected in case a drug was applied.</li>
</ol>
3. Analysis
&#62;NOTE: Depending on the imaging software used (which is typically a proprietary software package provided by the manufacturer of the camera or the whole fluorescence imaging system), it may be possible to perform the analysis of the acquired images partly or even entirely within this software package. However, here a workflow of image analysis that can be performed with open source software (i.e., the image analysis platform ImageJ)16, which can be conveniently installed using a distribution such as Fiji17, and the R package for statistical computing18 are described. Briefly, regions of interest (ROIs) representing cells or background are drawn in ImageJ, and the mean fluorescence in these ROIs over time is exported to a file to be, then, further analyzed in R or, alternatively, with spreadsheet software.
<ol>
	<li>In the imaging software, save or export the acquired image time series (both the recording containing the cells and the corresponding background recording) to a format (e.g., TIF) that can be read by ImageJ. Alternatively, use the BioFormats plugin (https://imagej.net/Bio-Formats), which provides routines enabling ImageJ to read proprietary file formats from different brands of imaging system manufacturers. 
	NOTE: The following steps assume that the imaging intervals are the same throughout the recording. If this is not the case, it may be necessary to export the timing information from the imaging software and include it in the further analysis.</li>
	<li>Open the TIF stack containing the cells in ImageJ.</li>
	<li>Use the freehand selection tool to draw an ROI over a fluorescent cardiomyocyte in the red channel. Ensure that the ROI is large enough so that the cell does not move out of the ROI while contracting.</li>
	<li>Open the ROI manager plugin (&#39;Analyze | Tools | ROI Manager&#39;) and press the &#39;Add [t]&#39; button to add this ROI as ROI1.</li>
	<li>Drag the ROI over the same cell in the green channel and add this ROI as ROI2.</li>
	<li>In the &#39;Analyze | Set Measurements&#39; menu, unselect all options except for &#39;Mean gray value&#39;. 
	NOTE: This has to be done only 1x in an ImageJ session.</li>
	<li>In the ROI manager, press the &#39;More | Multi Measure&#39; button. Select the &#39;Measure all slices&#39; and &#39;One row per slice&#39; options and press &#39;OK&#39;. A window opens containing a spreadsheet with three rows (the slice number and the mean fluorescence in the two ROIs representing the cell in the red and in the green channel, respectively).</li>
	<li>Use &#39;File | Save As&#39; to save the spreadsheet into a comma-separated value (CSV) file.</li>
	<li>Close the window with the image stack and the spreadsheet window without closing the ROI manager window. Open the TIF stack containing the background measurement.</li>
	<li>In the ROI manager, press the &#39;More | Multi Measure&#39; button. Select the &#39;Measure all slices&#39; and &#39;One row per slice&#39; options and press &#39;OK&#39;.</li>
	<li>Use &#39;File | Save As&#39; to save the spreadsheet with the background data into another CSV file. 
	NOTE: The following calculations can be done with spreadsheet software. We used the R software18. A simple example script that reads the cell and background data from the files &#8220;cell.csv&#8221; and &#8220;background.csv&#8221; performs the calculations, and plots the time course of the membrane potential signal is provided as Supplementary Code File 1.</li>
	<li>From the background recording, calculate the average intensity in the red and in the green channel. Subtract this background signal from the signals representing the cell in the red and green channel to calculate background-corrected RFP and GFP signals, respectively.</li>
	<li>As a surrogate for the membrane potential, calculate the ratio RFP/GFP. 
	NOTE: This ratio is dimensionless. Alternatively, it can be normalized to its initial value (i.e., &#916;R/R0). Importantly, useful information is contained in the temporal changes rather than in the absolute values of this ratio. Due to uneven photobleaching of the GFP and the RFP, a baseline drift in this ratio may occur. If necessary, such a baseline drift may be corrected by constructing a baseline curve and subtracting it from the RFP/GFP ratio10.</li>
</ol>

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The authors have nothing to disclose.

The method described here enables an optical recording of APs from a specific subtype (i.e., ventricular-like cells) of CMs generated from human iPSCs. Human iPSC-CMs are an emerging tool to address a huge variety of biological and medical problems, and the differentiation to different CM subtypes is an important source of experimental variability. By using specific promoter elements, the expression of a GEVI is specifically achieved in CMs representing the subtype of interest, which are then optically imaged.
The successful acquisition of APs from single iPSC-CMs depends on the reseeding density after the dissociation. If the cells are reseeded too dense, it is time-consuming to find isolated single iPSC-CMs that are not part of an electrical syncytium. However, too-low seeding densities negatively affect the survival of the cells. Thus, a systematical testing of different seeding densities is recommended, especially before engaging in experiments that are dependent on high-throughput imaging.
During the image acquisition, the VSFP undergoes photobleaching, which might negatively affect the signal-to-noise ratio, especially during longer experiments. If rapid photobleaching is present during the experiment, please ensure that the excitation light shutter is only opened during the image acquisition or during necessary preparation steps (searching for and focusing on VSFP-expressing cells), which should be performed as quickly as possible. In addition, reduce the power of the excitation light to the minimal amount necessary and make use of the dynamic range of the high-sensitivity camera used for image acquisition.
When drawing the ROI during the image analysis, be aware that iPSC-CMs move during contraction and should stay within the ROI in all images of the image stack. Due to the ratiometric nature of the VSFP, any movement of the cell out of the ROI does not lead to artificial signals but might negatively affect the signal quality. However, ROIs that are too large and include unnecessarily-large non-signal-producing areas also negatively affect the signal-to-noise ratio and should be avoided.
For the Representative Results, differentiated iPSC-CMs that have been allowed to mature for at least 30 days after a cardiac induction have been used. After this period of maturation, different CM subtypes, such as nodal-, atrial-, or ventricular-like cells can be distinguished. These cells are still immature and reach only an embryonic-like phenotype in two-dimensional (2-D) culture systems affecting electrical properties as well as calcium handling19. These general limitations of iPSC-CMs need to be considered, as they may confound experimental results independent of the method used for measuring action potentials. More advanced three-dimensional (3-D) culture systems have shown promising results toward further maturation20, and the method described here could be valuable in this context.
The current gold standard for measuring APs of iPSC-CMs is patch clamp recordings, which allow a very detailed analysis of different AP characteristics and even enables measuring single ion channel currents. In addition, absolute values (e.g., the resting membrane potential) can be measured. In contrast, optical AP recordings are based on relative changes of membrane potential. Due to the relatively slow intrinsic kinetics of the VSFP, certain AP characteristics, such as the &#34;overshoot&#34;, might be blunted from the readings, and a meaningful quantitation of fast features, such as the AP upstroke velocity, is not possible. However, in a typical experimental set-up, not the absolute values, but differences in values (e.g., between different cell lines or before and after the application of a drug) are measured. This can be reliably achieved with the optical method described in this manuscript for characteristics like AP duration (e.g., APD90 or APD50) or beating rate, with a much higher throughput as compared to patch clamp10. Furthermore, the non-invasive nature of this technology allows multiple measurements of the same cell over time10. Thus, optical AP recordings and patch clamp should be considered as complementary tools and used depending on the experimental question that needs to be answered.
Optical AP recordings can be achieved either by the expression of a GEVI or by the application of a potentiometric fluorescent dye (e.g., di-8-ANEPPs)21. The fluorescent dyes have usually more favorable fluorescence kinetics but cannot be directed to a certain subtype of cardiomyocytes as they are not genetically-encoded. A number of GEVIs based on different operating principles have been developed22. Here, the FRET-based GEVI VSFP-CR11 is used. The major reason for this choice was the ratiometric nature of this indicator, which reacts to cellular depolarization with an increase in red and a decrease in green fluorescence if the green fluorescent protein is excited. This is advantageous in the field of cardiology because such a ratiometric indicator is less susceptible to artifacts caused by cell movements due to contractions of the cardiomyocytes than indicators that react to depolarization only by a change of their fluorescence intensity.
While only results from experiments targeting ventricular-like iPSC-CMs using the MLC2v enhancer are presented here, it is also possible to target atrial- or nodal-like CMs using different subtype-specific promoter elements10. To describe the method in detail, this manuscript focused on imaging single iPSC-CMs that have been seeded on a glass coverslip, as this enables the analysis of AP properties of single cells and provides the highest signal-to-noise ratio. However, this technology also enables the imaging of CMs that are integrated with other cells in an electrical syncytium or a 3-D structure. This is of special interest as 3-D cell culture models are increasingly used and have been shown to provide properties different from 2-D models20. Depending on the imaging set-up and the definition of the ROIs, optical signals can be analyzed from larger areas of the 3-D tissue or from single cells within the tissue. Moreover, multiple cells can be imaged simultaneously and analyzed separately, leading to a substantial increase in throughput.
Taken together, the method presented here may improve the applicability of iPSC-CMs in the field of arrhythmia research by enabling the rapid optical phenotyping of iPSC-CMs for arrhythmia-related phenotypes.

Cardiomyocytes (CMs) derived from induced pluripotent stem cells (iPSCs) are an emerging tool to dissect molecular mechanisms of heart disease, to investigate novel therapies, and to screen for adverse cardiac drug effects1,2,3. Right from the start, arrhythmogenic diseases such as channelopathies have been an important focus of this research area4. Consequently, methods to investigate electrical phenotypes of CMs, such as arrhythmias or changes in action potential (AP) morphologies, are at the heart of this technology.
An important consideration in the application of iPSC-CMs is that current cardiac differentiation protocols do not result in a homogenous population of cells. Instead, they are rather a mixture of cells resembling sinus node, atrial, and ventricular CMs at different levels of maturation5,6,7,8. This heterogeneity can be a relevant source of experimental variability, especially if parameters such as AP duration (APD) are investigated, which intrinsically differ between CM subtypes (e.g., the APD is shorter in atrial than in ventricular CMs). The conventional approach to address this problem is to investigate single iPSC-CMs using the patch clamp method and to classify each cell as nodal-, atrial-, or ventricular-like, based on its AP morphology9. Any subsequent analysis can be then restricted to the cells representing the CM subtype of interest. The major drawback of this strategy is its limited throughput and lack of scalability. Moreover, the invasive nature of patch clamp electrophysiology does not allow the imaging of the same cells sequentially over extended time periods.
Here, we provide experimental details on a method10 developed to optically image APs in specific subtypes of iPSC-CMs. This overcomes the problem of subtype heterogeneity and dramatically increases the throughput as compared to conventional methods, allowing the rapid phenotyping of iPSC-CMs carrying genetic variants or being exposed to pharmacological agents.
Overview of the subtype-specific optical imaging approach
A genetically-encoded voltage indicator (GEVI), whose fluorescence properties change upon depolarization and repolarization of the cell membrane, is used to optically image changes of the membrane potential of CMs. The GEVI applied here is the voltage-sensing fluorescent protein VSFP-CR11, which consists of a voltage-sensing transmembrane domain fused to a pair of a green (Clover) and a red (mRuby2) fluorescent protein (Figure 1A). Due to the close proximity of the two fluorophores, the excitation of the green fluorescent protein results in a fraction of the excitation energy being transferred to the red fluorescent protein via F&#246;rster resonance energy transfer (FRET). Therefore, the excitation of the green fluorescent protein results in an emission from both the green and the red fluorescent proteins (Figure 1A, upper panel). When the cell depolarizes, a structural rearrangement of the voltage sensor occurs that translates into a reorientation of the two fluorescent proteins, increasing the FRET efficiency. Thus, even more of the excitation energy is transferred from the green to the red fluorescent protein (Figure 1A, lower panel). As a result, in a depolarized cell, the green fluorescence emission is dimmer, and the red fluorescence emission is brighter than in a cell at resting membrane potential (Figure 1B).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58134/58134fig1.jpg" /> 
Figure 1: Optical imaging of membrane potential with VSFP-CR. (A) A schematic depicting the action of the voltage-sensitive fluorescent protein VSFP-CR is shown. Upon the depolarization of the cell membrane, a structural rearrangement in the voltage-sensing transmembrane domain translates into a reorientation of the green (GFP) and red (RFP) fluorescent protein, increasing the efficiency of the intramolecular F&#246;rster resonance energy transfer (FRET). (B) The emission spectra of a VSFP upon excitation of the GFP in cells at the resting membrane potential (upper panel) and in depolarized cells (lower panel) are depicted. The spectral change upon depolarization is exaggerated for clarity. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58134/58134fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
The changes in the FRET efficiency mirroring the fluctuations of the membrane potential are imaged using a fluorescence microscope equipped with an image splitter, which separates red and green fluorescence emissions and projects them onto two adjacent areas of the chip of an sCMOS camera (Figure 2). With this set-up, the fluorescence emission at two different wavelength bands can be recorded simultaneously, which allows the calculation of a ratio of red-to-green fluorescence to reflect the membrane potential in every image of a time-lapse series.
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/58134/58134fig2.jpg" /> 
Figure 2: Configuration of the imaging system. The principal components of the imaging system used to image the spectral changes of the voltage-sensitive fluorescent protein mirroring the membrane potential changes at a high temporal resolution are depicted. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58134/58134fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
The expression of VSFP-CR in CMs is achieved by lentiviral transduction. To direct expression to the CM subtype of interest, the lentivirus contains a promoter element (the MLC2v enhancer) that specifically drives transcription in ventricular-like iPSC-CMs10. When the iPSC-CMs that represent a mixture of atrial-like, nodal-like, and ventricular-like cells are transduced with this lentivirus, VSFP-CR is expressed only in the ventricular-like cells. Since the optical action potential imaging depends on this fluorescent sensor, the recorded action potentials exclusively represent the CM subtype of interest (Figure 3).
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/58134/58134fig3.jpg" /> 
Figure 3: Promoter-driven VSFP expression for subtype-specific membrane potential imaging. (a)&#160;This schematic shows how cardiomyocyte subtype-specific optical action potential recordings are achieved. (b) iPSC-CMs infected with a VSFP under the control of the ventricular-specific MLC2v-enhancer are shown. The expression of the voltage sensor is observed only in ventricular-like CMs in the GFP channel (left panel). The phase contrast (middle panel) and overlay image (right panel) are also provided. The white dotted lines mark cell boundaries. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58134/58134fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table><tbody><tr><td>&amp;szlig;-Mercaptoethanol</td><td>Invitrogen</td><td>21985023</td><td></td></tr><tr><td>DMEM-F12 Medium</td><td>Invitrogen</td><td>21331046</td><td></td></tr><tr><td>FBS (Fetal Bovine Serum)</td><td>Invitrogen</td><td>16141079</td><td></td></tr><tr><td>MEM Non-Essential Amino Acids</td><td>Invitrogen</td><td>11140050</td><td></td></tr><tr><td>GlutaMax-I Supplement</td><td>Invitrogen</td><td>35050061</td><td>alternative L-Glutamine</td></tr><tr><td>Penicillin-Streptomycin</td><td>Invitrogen</td><td>15140122</td><td></td></tr><tr><td>Fibronectin bovine plasma</td><td>Sigma-Aldrich</td><td>F1141&amp;nbsp;</td><td></td></tr><tr><td>Collagenase type II</td><td>Worthington Biochem</td><td>LS004174</td><td></td></tr><tr><td>Hexadimethrine Bromide (Polybrene)</td><td>Sigma-Aldrich</td><td>H9268</td><td>enhancing lentiviral infection</td></tr><tr><td>3.5 cm glass-bottom microdishes</td><td>MatTek corporation, Ashland, MA, USA</td><td>P35G-1.5-14-C</td><td></td></tr><tr><td>Microscope stand</td><td>Leica Microsystems, Wetzlar, Germany</td><td>DMI6000B</td><td></td></tr><tr><td>Microscope objective</td><td>Leica Microsystems, Wetzlar, Germany</td><td>HCX PL APO 63X/1.4-0.6 Oil</td><td></td></tr><tr><td>sCMOS camera</td><td>Andor Technology, Belfast, UK</td><td>Zyla V</td><td></td></tr><tr><td>Microscope filter cube: excitation filter</td><td>Chroma Technology Corp, Bellows Falls, VT, USA</td><td>ET480/40X</td><td>bandpass 480/40</td></tr><tr><td>Microscope filter cube: dichroic mirror</td><td>Chroma Technology Corp, Bellows Falls, VT, USA</td><td>T505lpxr</td><td>longpass 505 nm</td></tr><tr><td>Image splitter</td><td>&amp;nbsp;Cairn Research, Faversham, UK</td><td>OptoSplit II</td><td></td></tr><tr><td>Image splitter filter cube: dichroic mirror</td><td>AHF Analysentechnik GmbH, T&amp;uuml;bigen, Germany</td><td>568LPXR</td><td>longpass 568 nm</td></tr><tr><td>Image splitter filter cube: emission filter 1 (GFP emission)</td><td>AHF Analysentechnik GmbH, T&amp;uuml;bigen, Germany</td><td>520/28 BrightLine HC</td><td>bandpass 520/28 nm</td></tr><tr><td>Image splitter filter cube: emission filter 2 (RFP emission)</td><td>AHF Analysentechnik GmbH, T&amp;uuml;bigen, Germany</td><td>630/75 ET Bandpass</td><td>bandpass 630/75 nm</td></tr><tr><td>Pacing inset</td><td>Warner Instruments, Hamden, CT, USA</td><td>RC-37FS</td><td></td></tr></tbody></table>

subtype-specific optical action potential recordings in human induced pluripotent stem cell-derived ventricular cardiomyocytes

Cardiomyocytes generated from human induced pluripotent stem cells (iPSC-CMs) are an emerging tool in cardiovascular research. Rather than being a homogenous population of cells, the iPSC-CMs generated by current differentiation protocols represent a mixture of cells with ventricular-, atrial-, and nodal-like phenotypes, which complicates phenotypic analyses. Here, a method to optically record action potentials specifically from ventricular-like iPSC-CMs is presented. This is achieved by lentiviral transduction with a construct in which a genetically-encoded voltage indicator is under the control of a ventricular-specific promoter element. When iPSC-CMs are transduced with this construct, the voltage sensor is expressed exclusively in ventricular-like cells, enabling subtype-specific optical membrane potential recordings using time-lapse fluorescence microscopy.

In Figure 4a, a representative single iPSC-CM is depicted with the white dotted lines marking the ROI drawn during the imaging analysis in the RFP (left side) and the GFP (right side) channel. The signal from the RFP channel shows a periodic increase in fluorescent intensity during each action potential (Figure 4b, upper panel). As described in the Introduction, this is due to an increasing FRET caused by changes in the membrane potential (Figure 1). Concordantly, the GFP signal shows a periodic decrease in fluorescent intensity (Figure 4b, middle panel). The RFP/GFP ratio (Figure 4b, lower panel) is the biological signal used in downstream analyses, such as APD50 and APD90 measurements. Using this method, 30-day-old ventricular-like iPSC-CMs paced at 0.5 Hz showed a mean APD50 (the duration from the beginning of the action potential until the repolarization is completed by 50%) of 439 ms (&#177; 46 ms) and a mean APD90 (the duration until the repolarization is completed by 90%) of 520 ms (&#177; 47 ms) (Figure 4c).
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/58134/58134fig4.jpg" /> 
Figure 4: Principles of image analysis and basic action potential characteristics. (a) The simultaneously recorded RFP (right side) and GFP (left side) fluorescence of a single iPSC-CM is shown. The white dotted lines represent the regions of interest (ROI) used for the imaging analysis. (b) Raw traces of background-corrected RFP, GFP, and RFP/GFP signals derived from the ROI are shown. (c) The means and SEM of APD90 and APD50 of single ventricular-like iPSC-CMs at room temperature and with 0.5-Hz pacing are shown. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58134/58134fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
To demonstrate that the method is also capable of capturing changes in action potential duration, iPSC-CMs were electrically stimulated at increasing stimulation rates ranging from 0.4 to 2.5 Hz, increasing the rate every five beats (Figure 5). A typical optically-recorded membrane potential signal from one iPSC-CM is shown in Figure 5a. The first four action potentials at each beating rate were averaged, demonstrating a progressive shortening of the action potentials at increased beating rates (Figure 5b). Averaging subsequent paced beats is an effective method of reducing the noise inherent to optically-recorded action potentials10. It should be mentioned that the depicted experiment might not capture the full amount of rate-dependent action potential shortening possible, given that we paced the cell for only five beats at each pacing rate, which might not be sufficient to reach equilibrium.
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/58134/58134fig5.jpg" /> 
Figure 5: Effect of the pacing frequency on the action potential duration. (a) An iPSC-CM was paced at increasing frequencies ranging from 0.4 to 2.5 Hz as indicated (the single electrical pacing pulses are represented by arrows). (b) For each pacing frequency, an averaged AP was calculated from the first four APs at this frequency. The averaged APs are plotted as an overlay, demonstrating the AP shortening with increasing pacing rates. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58134/58134fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
The method can also be used to investigate electrophysiological effects of drugs applied to the iPSC-CMs (Figure 6). Here, isoproterenol, a non-selective &#946;-adrenoreceptor agonist, was applied to spontaneously-beating iPSC-CMs as indicated, leading to a prompt increase in the beating rate (Figure 6a). A dose-response curve showing the effect of different Isoproterenol concentrations on the beating frequency of iPSC-CMs is shown in Figure 6b.
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/58134/58134fig6.jpg" /> 
Figure 6: Effect of isoproterenol on the beating frequency. (a) The representative raw optical action potential tracing of a spontaneously-beating single iPSC-CM is shown. Isoproterenol (with a final concentration of 1 &#181;M) was added to iPSC-CMs as indicated by the bar. (b) This dose-response curve shows the effect of different isoproterenol concentrations on the beating frequency of iPSC-CMs. Error bars represent standard deviation. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58134/58134fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Subtype-specific Optical Action Potential Recordings in Human Induced Pluripotent Stem Cell-derived Ventricular Cardiomyocytes at JoVE.com

Subtype-specific Optical Action Potential Recordings in Human Induced Pluripotent Stem Cell-derived Ventricular Cardiomyocytes

Here we present a method to optically image action potentials, specifically in ventricular-like induced pluripotent stem cell-derived cardiomyocytes. The method is based on the promoter-driven expression of a voltage-sensitive fluorescent protein.

Cardiomyocytes generated from human induced pluripotent stem cells (iPSC-CMs) are an emerging tool in cardiovascular research. Rather than being a ...

subtype-specific-optical-action-potential-recordings-human-induced

Nanyang Technological University

Research

JoVE Journal

Biology

7.1K Views.  Technical University of Munich. Cardiomyocyten gegenereerd uit geïnduceerde Pluripotente Stam of IPS cellen worden steeds vaker gebruikt om een verscheidenheid aan biologische vragen op het gebied van cardiale elektrofysiologie te beantwoorden. De belangrijkste voordelen van deze technologie zijn dat het een dramatische toename van de doorvoer mogelijk maakt, evenals subtypespecifieke actie potentiële opnames in ventriculaire zoals myocyten, met behulp van een ventriculaire specifieke promotor element. De introductie van ...

Video: Subtype-specific Optical Action Potential Recordings in Human Induced Pluripotent Stem Cell-derived Ventricular Cardiomyocytes

Electrophysiological Analysis of human Pluripotent Stem Cell-derived Cardiomyocytes (hPSC-CMs) Using Multi-electrode Arrays (MEAs)

Cardiomyocytes can now be derived with high efficiency from both human embryonic and human induced-Pluripotent Stem Cells (hPSC). hPSC-derived cardiomyocytes (hPSC-CMs) are increasingly recognized as having great value for modeling cardiovascular diseases in humans, especially arrhythmia syndromes. They have also demonstrated relevance as in vitro systems for predicting drug responses, which makes them potentially useful for drug-screening and discovery, safety pharmacology and perhaps eventually for personalized medicine. This would be facilitated by deriving hPSC-CMs from patients or susceptible individuals as hiPSCs. For all applications, however, precise measurement and analysis of hPSC-CM electrical properties are essential for identifying changes due to cardiac ion channel mutations and/or drugs that target ion channels and can cause sudden cardiac death. Compared with manual patch-clamp, multi-electrode array (MEA) devices offer the advantage of allowing medium- to high-throughput recordings. This protocol describes how to dissociate 2D cell cultures of hPSC-CMs to small aggregates and single cells and plate them on MEAs to record their spontaneous electrical activity as field potential. Methods for analyzing the recorded data to extract specific parameters, such as the QT and the RR intervals, are also described here. Changes in these parameters would be expected in hPSC-CMs carrying mutations responsible for cardiac arrhythmias and following addition of specific drugs, allowing detection of those that carry a cardiotoxic risk.

Electrophysiological Analysis of human Pluripotent Stem Cell-derived Cardiomyocytes hPSC-CMs Using Multi-electrode Arrays MEAs

Electrophysiological characterization of cardiomyocytes derived from human Pluripotent Stem Cells (hPSC-CMs) is crucial for cardiac disease modeling and for determining drug responses. This protocol provides the necessary information to dissociate and plate hPSC-CMs on multi-electrode arrays, measure their field potential, and a method for analyzing QT and RR intervals.

Cardiomyocytes can now be derived with high efficiency from both human embryonic and human induced-Pluripotent Stem Cells (hPSC). hPSC-derived ...

Developmental Biology

Human iPSC-Derived Cardiomyocyte Networks on Multiwell Micro-electrode Arrays for Recurrent Action Potential Recordings

Cardiac safety screening is of paramount importance for drug discovery and therapeutics. Therefore, the development of novel high-throughput electrophysiological approaches for hiPSC-derived cardiomyocyte (hiPSC-CM) preparations is much needed for efficient drug testing. Although multielectrode arrays (MEAs) are frequently employed for field potential measurements of excitable cells, a recent publication by Joshi-Mukherjee and colleagues described and validated its application for recurrent action potential (AP) recordings from the same hiPSC-CM preparation over days. The aim here is to provide detailed step-by-step methods for seeding CMs and for measuring AP waveforms via electroporation with high precision and a temporal resolution of 1 &#181;s. This approach addresses the lack of easy-to-use methodology to gain intracellular access for high-throughput AP measurements for reliable electrophysiological investigations. A detailed work flow and methods for plating of hiPSC-CMs on multiwell MEA plates are discussed emphasizing critical steps wherever relevant. In addition, a custom-built MATLAB script for rapid data handling, extraction and analysis is reported for comprehensive investigation of the waveform analysis to quantify subtle differences in morphology for various AP duration parameters implicated in arrhythmia and cardiotoxicity.

This article contains a set of protocols for the development of human induced pluripotent stem cell-derived cardiomyocyte (hiPSC-CM) networks cultured on multiwell MEA plates to reversibly electroporate the cell membrane for action potential measurements. High-throughput recordings are obtained from the same cell sites repeatedly over days.

Cardiac safety screening is of paramount importance for drug discovery and therapeutics. Therefore, the development of novel high-throughput ...

Bioengineering

Optical Imaging of Isolated Murine Ventricular Myocytes

The ability to isolate adult cardiac myocytes has permitted researchers to study a variety of cardiac pathologies at the single cell level. While advances in calcium sensitive dyes have permitted the robust optical recording of single cell calcium dynamics, recording of robust transmembrane optical voltage signals has remained difficult. Arguably, this is because of the low single to noise ratio, phototoxicity, and photobleaching of traditional potentiometric dyes. Therefore, single cell voltage measurements have long been confined to the patch clamp technique which while the gold standard, is technically demanding and low throughput. However, with the development of novel potentiometric dyes, large, fast optical responses to changes in voltage can be obtained with little to no phototoxicity and photobleaching. This protocol describes in detail how to isolate adult murine myocytes which can be used for cellular shortening, calcium, and optical voltage measurements. Specifically, the protocol describes how to use a ratiometric calcium dye, a single-excitation calcium dye, and a single excitation voltage dye. This approach can be used to assess the cardiotoxicity and arrhythmogenicity of various chemical agents. While phototoxicity is still an issue at the single cell level, methodology is discussed on how to reduce it.

We present the methodology for the isolation of murine myocytes and how to obtain voltage or calcium traces simultaneously with sarcomere shortening traces using fluorescence photometry with simultaneous digital cell geometry measurements.

The ability to isolate adult cardiac myocytes has permitted researchers to study a variety of cardiac pathologies at the single cell level. While advances ...

Single-Cell Optical Action Potential Measurement in Human Induced Pluripotent Stem Cell-Derived Cardiomyocytes

Conventional intracellular microelectrode techniques to quantify cardiomyocyte electrophysiology are extremely complex, labor intensive, and typically carried out in low throughput. Rapid and ongoing expansion of induced pluripotent stem cell (iPSC) technology presents a new standard in cardiovascular research and alternate methods are now necessary to increase throughput of electrophysiological data at a single cell level. VF2.1Cl is a recently derived voltage sensitive dye which provides a rapid single channel, high magnitude response to fluctuations in membrane potential. It possesses kinetics superior to those of other existing voltage indicators and makes available functional data equivalent to that of traditional microelectrode techniques. Here, we demonstrate simplified, non-invasive action potential characterization in externally paced human iPSC derived cardiomyocytes using a modular and highly affordable photometry system.

Here we describe optical acquisition and characterization of action potentials from induced pluripotent stem cell derived cardiomyocytes using a high-speed modular photometry system.

Conventional intracellular microelectrode techniques to quantify cardiomyocyte electrophysiology are extremely complex, labor intensive, and typically ...

Murine Short Axis Ventricular Heart Slices for Electrophysiological Studies

Murine cardiomyocytes have been extensively used for in vitro studies of cardiac physiology and new therapeutic strategies. However, multicellular preparations of dissociated cardiomyocytes are not representative of the complex in vivo structure of cardiomyocytes, non-myocytes and extracellular matrix, which influences both mechanical and electrophysiological properties of the heart. Here we describe a technique to prepare viable ventricular slices of adult mouse hearts with a preserved in vivo like tissue structure, and demonstrate their suitability for electrophysiological recordings. After excision of the heart, ventricles are separated from the atria, perfused with Ca2+-free solution containing 2,3-butanedione monoxime and embedded in a 4% low-melt agarose block. The block is placed on a microtome with a vibrating blade, and tissue slices with a thickness of 150-400 &#181;m are prepared keeping the vibration frequency of the blade at 60-70 Hz and moving the blade forward as slowly as possible. Thickness of the slices depends on the further application. Slices are stored in ice cold Tyrode&#39;s solution with 0.9 mM Ca2+ and 2,3-butanedione monoxime (BDM) for 30 min. Afterwards, slices are transferred to 37 &#176;C DMEM for 30 min to wash out the BDM. Slices can be used for electrophysiological studies with sharp electrodes or micro electrode arrays, for force measurements to analyze contractile function or to investigate the interaction of transplanted stem cell-derived cardiomyocytes and host tissue. For sharp electrode recordings, a slice is placed into a 3 cm cell culture dish on the heating plate of an inverted microscope. The slice is stimulated with a unipolar electrode, and intracellular action potentials of cardiomyocytes within the slice are recorded with a sharp glass electrode.

Here, we describe the preparation of viable ventricular slices from adult mice and their use for sharp electrode action potential recordings. These multicellular preparations provide a preserved in vivo like tissue structure, which makes them a valuable model for electrophysiological and pharmacological studies in vitro.

Murine cardiomyocytes have been extensively used for in vitro studies of cardiac physiology and new therapeutic strategies. However, multicellular ...

Medicine

High-Throughput Cardiotoxicity Screening Using Mature Human Induced Pluripotent Stem Cell-Derived Cardiomyocyte Monolayers

Human induced stem cell-derived cardiomyocytes (hiPSC-CMs) are used to replace and reduce the dependence on animals and animal cells for preclinical cardiotoxicity testing. In two-dimensional monolayer formats, hiPSC-CMs recapitulate the structure and function of the adult human heart muscle cells when cultured on an optimal extracellular matrix (ECM). A human perinatal stem cell-derived ECM (maturation-inducing extracellular matrix-MECM) matures the hiPSC-CM structure, function, and metabolic state in 7 days after plating. 
Mature hiPSC-CM monolayers also respond as expected to clinically relevant medications, with a known risk of causing arrhythmias and cardiotoxicity. The maturation of hiPSC-CM monolayers was an obstacle to the widespread adoption of these valuable cells for regulatory science and safety screening, until now. This article presents validated methods for the plating, maturation, and high-throughput, functional phenotyping of hiPSC-CM electrophysiological and contractile function. These methods apply to commercially available purified cardiomyocytes, as well as stem cell-derived cardiomyocytes generated in-house using highly efficient, chamber-specific differentiation protocols. 
High-throughput electrophysiological function is measured using either voltage-sensitive dyes (VSDs; emission: 488 nm), calcium-sensitive fluorophores (CSFs), or genetically encoded calcium sensors (GCaMP6). A high-throughput optical mapping device is used for optical recordings of each functional parameter, and custom dedicated software is used for electrophysiological data analysis. MECM protocols are applied for medication screening using a positive inotrope (isoprenaline) and human Ether-a-go-go-related gene (hERG) channel-specific blockers. These resources will enable other investigators to successfully utilize mature hiPSC-CMs for high-throughput, preclinical cardiotoxicity screening, cardiac medication efficacy testing, and cardiovascular research.

Human induced pluripotent stem cell-derived cardiomyocytes (hiPSC-CMs) offer an alternative to using animals for preclinical cardiotoxicity screening. A limitation to the widespread adoption of hiPSC-CMs in preclinical toxicity screening is the immature, fetal-like phenotype of the cells. Presented here are protocols for robust and rapid maturation of hiPSC-CMs.

Human induced stem cell-derived cardiomyocytes (hiPSC-CMs) are used to replace and reduce the dependence on animals and animal cells for preclinical ...

Generation of Human Cardiomyocytes: A Differentiation Protocol from Feeder-free Human Induced Pluripotent Stem Cells

In order to investigate the events driving heart development and to determine the molecular mechanisms leading to myocardial diseases in humans, it is essential first to generate functional human cardiomyocytes (CMs). The use of these cells in drug discovery and toxicology studies would also be highly beneficial, allowing new pharmacological molecules for the treatment of cardiac disorders to be validated pre-clinically on cells of human origin. Of the possible sources of CMs, induced pluripotent stem (iPS) cells are among the most promising, as they can be derived directly from readily accessible patient tissue and possess an intrinsic capacity to give rise to all cell types of the body 1. Several methods have been proposed for differentiating iPS cells into CMs, ranging from the classical embryoid bodies (EBs) aggregation approach to chemically defined protocols 2,3. In this article we propose an EBs-based protocol and show how this method can be employed to efficiently generate functional CM-like cells from feeder-free iPS cells.

Pluripotent stem cells, either embryonic or induced pluripotent stem (iPS) cells, constitute a valuable source of human differentiated cells, including cardiomyocytes. Here, we will focus on cardiac induction of iPS cells, showing how to use them to obtain functional human cardiomyocytes through an embryoid bodies-based protocol.

In order to investigate the events driving heart development and to determine the molecular mechanisms leading to myocardial diseases in humans, it is ...

Electromechanical Assessment of Optogenetically Modulated Cardiomyocyte Activity

Over the past two decades, optogenetic tools have been established as potent means to modulate cell-type specific activity in excitable tissues, including the heart. While Channelrhodopsin-2 (ChR2) is a common tool to depolarize the membrane potential in cardiomyocytes (CM), potentially eliciting action potentials (AP), an effective tool for reliable silencing of CM activity has been missing. It has been suggested to use anion channelrhodopsins (ACR) for optogenetic inhibition. Here, we describe a protocol to assess the effects of activating the natural ACR GtACR1 from Guillardia theta in cultured rabbit CM. Primary readouts are electrophysiological patch-clamp recordings and optical tracking of CM contractions, both performed while applying different patterns of light stimulation. The protocol includes CM isolation from rabbit heart, seeding and culturing of the cells for up to 4 days, transduction via adenovirus coding for the light-gated chloride channel, preparation of patch-clamp and carbon fiber setups, data collection and analysis. Using the patch-clamp technique in whole-cell configuration allows one to record light-activated currents (in voltage-clamp mode, V-clamp) and AP (current-clamp mode, I-clamp) in real time. In addition to patch-clamp experiments, we conduct contractility measurements for functional assessment of CM activity without disturbing the intracellular milieu. To do so, cells are mechanically preloaded using carbon fibers and contractions are recorded by tracking changes in sarcomere length and carbon fiber distance. Data analysis includes assessment of AP duration from I-clamp recordings, peak currents from V-clamp recordings and force calculation from carbon fiber measurements. The described protocol can be applied to the testing of biophysical effects of different optogenetic actuators on CM activity, a prerequisite for the development of a mechanistic understanding of optogenetic experiments in cardiac tissue and whole hearts.

We present a protocol for evaluating the electromechanical effects of GtACR1 activation in rabbit cardiomyocytes. We provide detailed information on cell isolation, culturing and adenoviral transduction, and on functional experiments with the patch-clamp and carbon-fiber techniques.

Over the past two decades, optogenetic tools have been established as potent means to modulate cell-type specific activity in excitable tissues, including ...

Technical Applications of Microelectrode Array and Patch Clamp Recordings on Human Induced Pluripotent Stem Cell-Derived Cardiomyocytes

Drug-induced cardiotoxicity is the leading cause of drug attrition and withdrawal from the market. Therefore, using appropriate preclinical cardiac safety assessment models is a critical step during drug development. Currently, cardiac safety assessment is still highly dependent on animal studies. However, animal models are plagued by poor translational specificity to humans due to species-specific differences, particularly in terms of cardiac electrophysiological characteristics. Thus, there is an urgent need to develop a reliable, efficient, and human-based model for preclinical cardiac safety assessment. Human-induced pluripotent stem cell-derived cardiomyocytes (hiPSC-CMs) have emerged as an invaluable in vitro model for drug-induced cardiotoxicity screening and disease modeling. hiPSC-CMs can be obtained from individuals with diverse genetic backgrounds and various diseased conditions, making them an ideal surrogate to assess drug-induced cardiotoxicity individually. Therefore, methodologies to comprehensively investigate the functional characteristics of hiPSC-CMs need to be established. In this protocol, we detail various functional assays that can be assessed on hiPSC-CMs, including the measurement of contractility, field potential, action potential, and calcium handling. Overall, the incorporation of hiPSC-CMs into preclinical cardiac safety assessment has the potential to revolutionize drug development.

Human-induced pluripotent stem cell-derived cardiomyocytes (hiPSC-CMs) have emerged as a promising in vitro model for drug-induced cardiotoxicity screening and disease modeling. Here, we detail a protocol for measuring the contractility and electrophysiology of hiPSC-CMs.

Drug-induced cardiotoxicity is the leading cause of drug attrition and withdrawal from the market. Therefore, using appropriate preclinical cardiac safety ...

Laser-Induced Action Potential-Like Measurements of Cardiomyocytes on Microelectrode Arrays for Increased Predictivity of Safety Pharmacology

Life-threatening drug-induced cardiac arrhythmia is often preceded by prolonged cardiac action potentials (AP), commonly accompanied by small proarrhythmic membrane potential fluctuations. The shape and time course of the repolarizing fraction of the AP can be pivotal for the presence or absence of arrhythmia.
Microelectrode arrays (MEA) allow easy access to cardiotoxic compound effects via extracellular field potentials (FP). Although a powerful and well-established tool in research and cardiac safety pharmacology, the FP waveform does not allow to infer the original AP shape due to the extracellular recording principle and the resulting intrinsic alternating current (AC) filtering.
A novel device, described here, can repetitively open the membrane of cardiomyocytes cultivated on top of the MEA electrodes at multiple cultivation time points, using a highly focused nanosecond laser beam. The laser poration results in transforming the electrophysiological signal from FP to intracellular-like APs (laser-induced AP, liAP) and enables the recording of transcellular voltage deflections. This intracellular access allows a better description of the AP shape and a better and more sensitive classification of proarrhythmic potentials than regular MEA recordings. This system is a revolutionary extension to the existing electrophysiological methods, permitting accurate evaluation of cardiotoxic effect with all advantages of MEA-based recordings (easy, acute, and chronic experiments, signal propagation analysis, etc.).

The combination of laser poration and microelectrode arrays (MEA) allows action potential-like recordings of cultivated primary and stem cell-derived cardiomyocytes. The waveform shape provides superior insight into test compounds' mode of action than standard recordings. It links patch-clamp and MEA readout to further optimize cardio safety research in the future.

Subtype-specific Optical Action Potential Recordings in Human Induced Pluripotent Stem Cell-derived Ventricular Cardiomyocytes

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