Name: Video: Laser-Induced Action Potential-Like Measurements of Cardiomyocytes on Microelectrode Arrays for Increased Predictivity of Safety Pharmacology
Uploaded: 2022-09-13T14:10:00.000+00:00
Duration: 10 min 41 s
Description: 1.9K Views. 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.

The authors would like to thank Foresee Biosystems for lending the IntraCell system during the studies. They would also like to thank Hae In Chang for technical assistance. This work has received funding from the European Union&#39;s Horizon 2020&#39;s research and innovation program under the grant agreement no. 964518 (ToxFree), from the EU Horizon Europe European Innovation Council Programme, project SiMulTox (grant agreement No 101057769), and from the State Ministry of Baden-Wuerttemberg for Economic Affairs, Labour and Tourism.

1. Induced pluripotent stem cell-derived cardiomyocytes preparation
NOTE: iCell cardiomyocytes2 (referred to as induced pluripotent stem cell [iPSCs]-derived cardiomyocytes) were prepared according to the protocol provided by the supplier. The protocol will be briefly summarized in the following section.
<ol>
	<li>Thaw plating and maintenance medium at 4 &#176;C for 24 h prior to use.</li>
	<li>Prepare fibronectin coating by dissolving sterile fibronectin in sterile water at a concentration of 1 mg/mL. Freeze store this stock in aliquots (e.g., 25 &#181;L per aliquot). Dilute the aliquoted stock solution 1:20 in sterile Dulbecco&#39;s balanced salt solution (dPBS).</li>
	<li>Coat the electrode fields of the previously autoclaved MEAs under the laminar flow hood, droplet-wise under sterile conditions with fibronectin using a 10 &#181;L pipette. For this, drop 5 &#181;L of the fibronectin coating solution onto the electrode fields and observe the formation of a droplet on top of the electrode area. Make sure not to touch the sensitive electrodes. 
	NOTE: To maintain sterile conditions, transfer the MEA chip into a sterile Petri dish before removing it from the laminar flow hood.</li>
	<li>Incubate the coated MEAs for 1 h at 37 &#176;C in an incubator preferably.</li>
	<li>Thaw the cryovial containing the cardiomyocytes in a water bath at approximately 37 &#176;C for 2 min until only a little ice crystal is left. Transfer the cell solution gently to a 50 mL tube.</li>
	<li>Add 1 mL of plating medium to the empty cryovial. Transfer the solution dropwise over a time period of 90 s into the 50 mL tube to reduce the osmotic shock. Gently add additional 8 mL of plating medium into the tube.</li>
	<li>Mix the cell suspension carefully using a 10 mL pipette. Calculate the total number of viable cells using automated fluorescence cytometry. The number should be close to the number in the datasheet provided by the manufacturer.</li>
	<li>Spin the cell solution down for 3 min at 200 x g at room temperature. Remove the supernatant by aspiration using a glass pipette attached to a pumping system. Adjust the cell number between 6,000 and 15,000 viable cells/&#181;L. 
	NOTE: The number of cells is usually in the range given above but can vary depending on the respective supplier.</li>
	<li>Remove the coating solution that was applied in step 1.3 from the MEA electrode area using a 10 &#181;L pipette directly before cell seeding. Seed the cells immediately after the removal to avoid drying of the coating. For this, seed the cells dropwise at 4 &#181;L for both 6-well MEAs and 1-well MEAs onto the electrode fields in the same manner as done with the coating.</li>
	<li>Allow the cells to adhere for 1 h in the incubator at 37 &#176;C and 5% CO2 prior to filling the wells with the sterile plating medium heated to approximately 37 &#176;C at 200 &#181;L for 6-well MEA and 1 mL for single well MEA under the laminar flow hood.</li>
	<li>Perform a complete medium change 48 h after plating under the laminar flow hood. For this, remove the plating medium by aspiration using a glass pipette attached to a pumping system. Then, add 200 &#181;L of sterile maintenance medium heated to 37 &#176;C to the wells.</li>
	<li>Perform complete medium changes every other day.</li>
	<li>Start measuring the cells 5-8 days after thawing. Perform a complete medium change 2 h prior to starting the experiments.</li>
</ol>
2. MEA recordings
NOTE: The device used to transform the FP signal to liAP consists of an upright microscope and a 1064 nm laser.
<ol>
	<li>Place the MEA system on top of the device with the MEA chip holder centered over the objective hole. Position the MEA set up so that the objective is directly under the hole of the MEA system to allow the laser to focus on the electrodes.</li>
	<li>Transfer the MEA chip with the cultivated cells from the incubator to the MEA setup 15 min prior to recording, allowing the cells to recover from the mechanical disturbance.</li>
	<li>Clean the contact pads and pins carefully using isopropanol and a cotton swab to decrease noise levels. Place the MEA carefully in the MEA-setup. Position the MEA chip with the logo at the bottom on the top left side (6-well MEA) or with the reference electrode to the left (single-well MEA).</li>
	<li>Set the MEA system-integrated heating to 38 &#176;C. Place a small chamber on top of the MEA chip to constantly perfuse the cells with humidified carbogen (5% CO2 and 95% O2) to recreate incubator conditions and prevent evaporation.</li>
	<li>Close the lid of the device. The integrated safety switch only allows the laser to be activated if the lid is closed over the MEA chip. Set the MEA system filter using the MEA config program to 0.1 Hz or less high-pass and 3,500 Hz low-pass.</li>
	<li>Use the MC_Rack software (recording software) or any alternative software for recording. Adjust the input range according to your needs ensuring that the signal does not saturate the amplifier and the sampling rate (e.g., 20 kHz). Use the long-term display function of the software to check the recording.</li>
</ol>
3. Laser-induced cell poration
<ol>
	<li>After inserting the MEA chip into the MEA setup and setting up the software, initialize the laser mechanics using the FB Alps software (initialization software).</li>
	<li>First, click on the Initialization button. At the end of initialization, the virtual laser point will be in well D for a 6-well MEA and at the bottom left for a single-well MEA, respectively.</li>
	<li>Move the virtual laser point with Ctrl + mouse click into the middle of electrode D5 and adjust the focus. Adjust the focus by Ctrl + scrolling with the mouse wheel.</li>
	<li>Press the button Set P1. The virtual laser point will automatically move into well F. Repeat the process with the electrode F5 and select Set P2.</li>
	<li>After this procedure, the laser point will move into well B. Repeat the same process with electrode B5 and press Set P3 in the software. The system is now aligned.</li>
	<li>Adjust the laser power and process time according to the needs of your cells. Here,&#160;40% power and 25% process time were used.</li>
	<li>To enable the laser, click on the Laser Off button, which will then appear as laser on. Switch to the recording software, choose a filename, and click on the Red Recording button followed by the Play button on top of the window to record the measurement.</li>
	<li>Record a baseline of 60 s prior to opening the cells with the laser. Switch back to the initialization software.</li>
	<li>Deactivate electrodes to be excluded by the laser on the virtual map on the right-side using Ctrl + mouse click. Select the electrodes of interest by activating them on the array representation on the right-hand side of the software window.</li>
	<li>To start the laser, use Alt + mouse click and select the center electrode of this well. This initiates the laser to automatically open the cells on each electrode of this well. Repeat for each well. The laser will then open all the previously activated electrodes of the selected well automatically.</li>
</ol>
4. Drug handling and application
<ol>
	<li>Prepare all substances to be used for drug tests freshly on the day of the measurements. Ensure that the final application concentration is 10 times higher in medium to make a 1:10 dilution in the wells.</li>
	<li>Dissolve Nifedipine, E4031, and Dofetilide first in DMSO in mM concentration, and then further in medium to 10x of the desired concentration. Never exceed a final DMSO concentration of 0.1% in the well.</li>
	<li>Record the baseline activity for 60 s. Start the laser-induced poration as described in step 3 of the protocol, resulting in a transformation of the FP into liAP shape.</li>
	<li>Apply all the drugs as single-concentration-per-well. Remove 20 &#181;L of medium per well for 6-well MEAs and 100 &#181;L for single-well MEAs, respectively. Add 20 &#181;L or 100 &#181;L, depending on the MEA type, of the stock solution of the drug to be measured to the well and carefully pipette up and down 2-3 times. Perform at least three replications of each drug and concentration to achieve statistical relevance.</li>
	<li>Allow the compounds to wash in for 300 s. During this time, the liAP shape may transform back into FP shape. Again, induce laser-induced poration and record possible compound-induced effects on the liAP for an additional 60 s.</li>
</ol>
5. Data export
<ol>
	<li>Select relevant electrodes by replaying the recording in the recording software. Use MC_DataTool to convert MC_Rack files to ASCII .txt files.</li>
	<li>Click on File &#62; Open MCD &#62; Select a MC_Rack file. Click on the txt button (blue text).</li>
	<li>Select an electrode. The selected electrode will be shown in the list on the right side.</li>
	<li>Click on Browse to select the folder and change the name of the new .txt file. Click on Save.</li>
	<li>Unselect the exported electrode and select the next electrode to export. Repeat the procedure for all electrodes of interest.</li>
</ol>
6. Data handling and statistical analysis
<ol>
	<li>Import converted binary traces into R19. Visualize/analyze the data using custom-tailored scripts containing the following packages: dplyr, tidyr, and ggplot220,21,22.</li>
</ol>

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The authors have nothing to disclose.

This innovative method demonstrates a new way to investigate in vitro the pharmacological modulation of the cardiac action potential during the application of cardioactive pharmacological tool compounds.
Classical MEA recordings allow FP recordings, which are the derivative of the cardiac AP14. This indirect recording convolves the time course of the de- and repolarization and thereby eliminates essential characteristics of the AP. Furthermore, although the transcellular voltage change of an AP typically reaches values of approximately 100 mV, the overall FP amplitude remains comparably low, with peak amplitudes between several 100 &#181;V and low single-digit mV values. Due to the recording principle, the repolarizing phase is small; in many cases, it is merely detectable and often of unclear shape, making it difficult to define the end of the FP. The opening of the cell membrane allows us to gain access to the intracellular voltage, thereby uncovering the time course of the cardiac AP. There are multiple advantages of this recording method in comparison to FP recordings. Firstly, the signal amplitude is more prominent, providing a superior signal-to-noise ratio. Secondly, the waveform results in better detection of the repolarization. Thirdly, the shape of the repolarization phase contributes insights into the mode of action of the test compound, provided by the steepness of the signal relaxation. And lastly, this method offers an improved sensitivity to detect critical adverse drug effects, demonstrated by the recording example displayed in Figure 6 for the occurrence of EADs in the liAP but not in the FP.
So far, there are two ways of gaining access to the intracellular AP. The first one is achieved by electroporation26,27. Here, short and strong voltage pulses applied via the recording electrodes can open the cell membrane28. The second possibility is the membrane opening via a laser pulse, making use of a physical phenomenon named surface plasmon resonance, as demonstrated here. One of the advantages compared to electroporation is the increased likelihood of consecutive openings. Due to the highly focused laser spot (1-3 &#181;m) this effect is very locally limited to the electrode of interest. Interestingly, the initiation of the liAP did not alter the signal propagation of the cultivated syncytium. This indicates that, although the cell integrity is damaged, the cardiomyocytes do not seem to depolarize via the hole in the membrane.
There are limitations to this method. Like with electroporation, the membrane opening does, in most cases, not last over the entire experimental course. The minimal power and duration settings of the laser pulse required for stable opening of the specific cell type of interest need to be defined independently prior to the experiments. We found (not shown) that the parameters drastically vary between different cell types (in our case, several hiPS-derived and primary cardiomyocytes). This avoids unnecessary stress on the cells during the compound test experiment and results in more reliable and reproducible data. It is of critical importance to adjust the z-axis to provide a clear focus on the cells and the electrodes. An unfocused camera picture yields in a laser spot located at a suboptimal level, potentially resulting in the inability to open the cell membrane. Even with best-adjusted parameters, the liAP effect is transient, and the amplitude reduces over time. Furthermore, the access to the intracellular space of the cells varies between liAP inductions, both within consecutive openings at the same electrode and between electrodes. This results in high variability of the liAP amplitude. The reason is not yet fully understood. Possible explanations include mechanical issues such as a drift of the laser focus or different subcellular localization of the membrane opening. This makes the analysis of amplitude effects of test compounds complicated at this point in time. Also, recording electrical activity by a MEA system requires high pass filtering to compensate for unavoidable baseline drift. Although in the system used here, this filtering was set to 0.1 Hz (the lowest filter setting available for this system), filtering effects during the plateau phase were still visible, resulting in a slow trend of the voltage deflection toward the baseline during the plateau phase of the cardiac AP. This is especially problematic with extensively long underlying APs like the iPSC-derived iCell cardiomyocytes2 used here, which already generate AP &#62;700 ms under control conditions. The use of systems with lower filtering may better conserve the shape of the AP and allow even better access to the time course of the repolarization phase.

The electrical contribution of a heartbeat results from a complex and precisely timed interplay of many cardiac channels and transporters, as well as the precisely tuned propagation of electrical signals through the myocardium1. Alteration of these closely coordinated mechanisms (e.g., using drugs) can result in severe consequences for the function of the heart (i.e., life-threatening arrhythmia)2,3. Arrhythmias are defined as irregular heartbeats that alter the normal rhythm of the heart, which can have life-threatening consequences. They may be caused either by impaired initiation of a wave of cardiac excitation or by abnormal propagation of cardiac excitation4, which in turn results in a dysfunction of the heart's pumping mechanism.
Many highly potent drug candidates must be excluded from further investigations during the early drug development phase due to their (pro-) arrhythmic potential2,3. They modulate key cardiac channels (e.g., the human ether-a-go-go-related gene channel [hERG]) that are responsible for normal cardiac action potential formation and termination as well as subsequent signal propagation5.
Pharmaceutical companies routinely use patch-clamp measurements or microelectrode arrays (MEA) to investigate potential cardiotoxic off-target effects induced by drug candidates. Patch-clamp recordings allow to decipher the impact of substances on cardiac ion channels and to analyze the transcellular cardiac action potential with high spatio-temporal resolution6,7. However, disadvantages of this technique include low throughput with manual patch-clamp and limited applicability of automation due to the dependence of this method on cells in suspension. Furthermore, chronic effects cannot be investigated due to the invasiveness of the method. Finally, typically only single cells are studied simultaneously rather than the entire cardiac syncytium, making it impossible to address information about signal propagation.
Voltage-sensitive dyes are valuable for noninvasively investigating cardiac action potentials and drug-induced arrhythmias8. They allow the investigation of both single-cell and syncytium activity. Drawbacks of this method are cytotoxic effects of either the dyes per se or of the reaction product during illumination. They are used for acute experiments and are hardly applicable for long-term studies9,10,11. Voltage-sensitive proteins as alternatives have made significant progress over the last couple of years in terms of usability and sensitivity but require genetic modification of the cells of interest and lack high temporal resolution compared to electrophysiological techniques12.
Information from the most recent CiPA initiative13 states that MEAs are widely used in cardiac safety screenings as an alternative electrophysiological approach as they represent a powerful and well-established tool to investigate cardiac function and safety pharmacology. Cardiomyocytes are cultivated as a syncytium directly on top of the chips, and extracellular field potentials (FPs) are recorded noninvasively via substrate-integrated microelectrodes. This recording principle allows conducting increased throughput screenings over several days, which makes them suitable for pharmaceutical research on chronic effects. The resulting FP waveform is a derivative of the intracellular AP14. Parameters such as beat rate, the amplitude of the initial part of the FP, and FP duration are easily accessible15. Other essential criteria such as the differentiation between prolongation and triangulation of the FP (an important marker of proarrhythmia16,17) are inaccessible due to the AC filtering effect of the technique. Furthermore, detecting other small proarrhythmic events such as early and delayed afterdepolarizations (EAD and DAD, respectively) are often easily overlooked due to their small amplitude.
Here we describe a method for gaining access to the intracellular membrane potential by opening the membrane of cardiomyocytes. The IntraCell device (hereafter referred to as intracellular recording device) allows repeated membrane openings of cardiomyocytes cultivated on top of the MEA electrodes using a highly focused nanosecond laser beam via a specific physical phenomenon (surface plasmon resonance)18. As a result, the recording transitions from a regular FP into an intracellular-like AP (laser-induced AP, liAP). The protocol shows how this allows gaining access to kinetic aspects of the waveform that cannot easily be captured by analyzing FPs. This method represents a bridge between traditional intracellular patch-clamp and MEA recordings. The technology is therefore a powerful extension of current cardiac safety assessment methods.

<table><tbody><tr><td>1 well MEA chip</td><td>Multi Channel Systems MCS GmbH</td><td>890301</td><td></td></tr><tr><td>6 well MEA chip</td><td>Multi Channel Systems MCS GmbH</td><td>7600069</td><td></td></tr><tr><td>DMSO</td><td>Merck KGaA</td><td>20-139&lt;br/&gt; Sigma-Aldrich</td><td>solvent for drugs</td></tr><tr><td>Dofetilide</td><td>ALOMONE LABS&lt;br/&gt; ISRAEL HEADQUARTERS</td><td>D-100</td><td>Drug-Measurement</td></tr><tr><td>dPBS</td><td>Fisher Scientific GmbH</td><td>12037539</td><td>Coating</td></tr><tr><td>E4031</td><td>ALOMONE LABS&lt;br/&gt; ISRAEL HEADQUARTERS</td><td>E-500</td><td>Drug-Measurement</td></tr><tr><td>Falcon</td><td>Fisher Scientific GmbH</td><td>10788561</td><td></td></tr><tr><td>FB Alps version 0.5.005</td><td>Foresee Biosystems</td><td></td><td></td></tr><tr><td>Fibronectin</td><td>Merck KGaA</td><td>11051407001</td><td>Coating</td></tr><tr><td>iCell cardiomyocytes</td><td>FUJIFILM Cellular&lt;br/&gt; Dynamics, Inc. (FCDI)</td><td>C1016</td><td></td></tr><tr><td>IntraCell</td><td>Foresee Biosystems</td><td></td><td></td></tr><tr><td>Isopropanol</td><td>Carl Roth GmbH + Co. KG</td><td>CN09.1</td><td>For cleaning of MEA contact pads</td></tr><tr><td>Maintenance Medium</td><td>FUJIFILM Cellular&lt;br/&gt; Dynamics, Inc. (FCDI)</td><td>#M1003</td><td>For cell-culture</td></tr><tr><td>MC_Data Tool</td><td>Multi Channel Systems MCS GmbH</td><td></td><td>Data export</td></tr><tr><td>MC_Rack</td><td>Multi Channel Systems MCS GmbH</td><td></td><td>MEA recording</td></tr><tr><td>MEA 2100 - 2x60 - system</td><td>Multi Channel Systems MCS GmbH</td><td>890485</td><td>For MEA-recordings</td></tr><tr><td>Nifedipine</td><td>Merck KGaA</td><td>N7634&lt;br/&gt; Sigma-Aldrich</td><td>Drug-Measurement</td></tr><tr><td>Plating Medium</td><td>FUJIFILM Cellular&lt;br/&gt; Dynamics, Inc. (FCDI)</td><td>M1001</td><td>For cell-culture</td></tr><tr><td>Tergazyme</td><td>VWR International, LLC</td><td>1304-1</td><td>cleaning of MEAs</td></tr></tbody></table>

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 recording system used to record electrical activity from cultivated cardiomyocytes consisted of a standard MEA system equipped with a heater and a chamber for carbogen attached to a computer. The system was set on top of the intracellular recording device, which in turn was mounted on top of a small anti-vibration unit (Figure 1A-B).
iPSC-derived cardiomyocytes2 started beating spontaneously within 2-3 days after thawing (days in vitro, DIV) and were visible under a microscope. From DIV 4 onward, the beating frequency became regular, and extracellular field potentials (FP) with peak-to-peak amplitudes of the depolarizing component between 1 to 5 mV could be detected on most of the electrodes within the respective wells of the MEA chips. The electrical activity could be detected in more than 95% of the wells under investigation. From DIV 7 onward, the probability of cell detachment increased, making further use of these wells impossible.
The software to control the laser-induced membrane opening allows adjusting both the power and process time of the laser that mediates the opening of the cell membrane only on the electrode under investigation (Figure 1C), whereas other electrodes in the respective well are unaffected. While too conservative settings did not change the waveform of the FP, too high settings resulted in the putative injury of the cardiomyocytes, indicated by vigorous but transient beating or loss of the signal. When adjusted to a setting of 40% power and 25% process time well-tolerated by the cells, triggering of the laser pulse resulted in multiple changes to the recorded waveform (see Figure 1D for an exemplary recording). Under these conditions, no alteration of the electrode material was macroscopically observed. The recorded signal amplitude massively increased by 4.1 &#177; 0.41 (n = 20, range 1.34-8.83) times, analyzed from a randomly picked subset of recordings, resulting in amplitudes between 7 and 22 mV. Furthermore, the waveform transformed from a standard FP shape with rapid, biphasic, and transient voltage deflection at the beginning, followed by a plateau phase back at baseline and a small deflection indicating the end of the FP to a shape that was closer to an intracellularly recorded AP with a rapid rise, extended depolarized plateau phase and a repolarization phase with an undershoot below the baseline (Figure 1E). We defined these voltage deflections as laser-induced AP (liAP). In most cases, the transition was transient and at least partially inverted within 5 min. Signal propagation within the cardiac syncytium remained unaltered after liAP induction (Figure 2), indicating that the remaining syncytium was not affected by potential damage from the laser pulse.
Similarities to intracellularly recorded APs allowed for extracting parameters of the liAP that are not accessible to FPs (for exemplary parameters see e.g., Figure 3A), most prominently the measurement of the duration of the liAP at specific time points (e.g., at 20%, 50%, and 90%) (Figure 3B), analogous to APD20/50/90 commonly used for the description of APs.
We next tested the response of the laser-opened cardiomyocytes to commonly used cardioactive pharmacological tool compounds. An exemplary protocol design can be found in Figure 3C. Since the transformation of the liAP did not always persist throughout the entire experiment, the compound application was performed as a single-concentration-per-well rather than in a cumulative manner to reduce the total recording time. Nevertheless, it was necessary to either reopen the cells or open another electrode area prior to applying the test compound.
The addition of the specific L-type Ca2+ channel blocker, Nifedipine23,24, reduced the plateau phase of the liAP in a concentration-dependent manner and thereby shortened the entire liAP (Figure 4A,B). This shortening was comparable to the analysis obtained from FPs of cardiomyocytes from unmanipulated electrodes (Figure 4C), indicating that this recording method did not have adverse effects compared to classical FP recordings.
E4031 inhibits the repolarization of relevant Kv1.11 (hERG) potassium channel25 and leads to arrhythmic behavior of cardiomyocytes at increased concentrations. Similar to the analysis obtained from FP recordings, E4031 increased the liAP duration in a concentration-dependent manner (Figure 5). Additionally, at concentrations of 0.01 &#181;M and higher, small positive voltage deflections at the end of the liAP were visible. These deflections became more prominent with higher concentrations, indicating a transient new depolarization, unlike in the FPs, where these deflections were virtually invisible (see Figure 5B-C, upper (FP) vs lower (liAP) traces). This behavior is known as early afterdepolarization (EAD). At the highest concentration of 0.1 &#181;M, these EADs escalated over time into ectopic beats, which are premature action potentials (Figure 5C). Both EAD and ectopic beats are key indicators of proarrhythmic activity. At the end of the example shown in Figure 5D, the electrical activity resulted in arrhythmic beating. Also, concentration-response-relationships displayed between FP and liAP recordings were matching (Figure 5E). However, there is more considerable variability in the FP data resulting from the weak repolarization component of the FPs at higher concentrations of the test compound. It seems to be the characteristic nature of iPSC-derived cardiomyocytes2 to tend to generate unphysiologically long APs under control conditions also (AP durations &#62; 700 ms). The MEA system applied an intrinsic 0.1 Hz AC filtering, which in turn resulted in a partially filtered shape of the liAP, although without occluding the qualitative information about the onset and termination of the underlying AP.
It turned out that the initial occurrence of proarrhythmic voltage deflections could be detected at lower concentrations in liAPs in comparison to FP recordings. Shown in Figure 6 is the recording of electrical activity during the application of Dofetilide at a concentration of 3 &#181;M. The recording was obtained in the same well. Although both FP and liAP displayed durations of approximately 2 s, the FP waveform was unobtrusive, presenting regular repolarizing deflections. At the same time, at the end of liAPs, EADs at different magnitudes became visible. This increase in relevant safety-pharmacological sensitivity further supports the finding that liAPs induced by surface plasmon resonance allow improved qualification of the repolarizing phase and thereby help to learn more about the mode of action of the test compounds under investigation.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64355/64355fig1v2.jpg" /> 
Figure 1: The intracellular recording setup and exemplary recordings. (A) Setup overview. (B) Top view of the recording system with open MEA recording amplifier. (C) Initialization software with the virtual MEA map on the right-hand side. 1: anti-vibration table, 2: intracellular recording&#160;system, 3: laser protection lid, 4: humidified carbogen chamber, 5: MEA heating system, 6: MEA interface board, 7: 1-well MEA chip inside the recording amplifier. (D) Recording examples from one electrode before and after induction of liAPs. Top: recording of approximately 6 min. Dotted lines mark the expanded areas shown at the bottom. (E) Magnified FP (top) and liAP (bottom). <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/64355/64355fig1v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/64355/64355fig02.jpg" /> 
Figure 2: Signal propagation pattern remains conserved after liAP induction. False-color coding of the signal propagation of the excitation wave within the syncytium. Blue indicates early (starting at -4 ms); red indicates late time points (+3 ms) from the signal obtained at reference electrode E54 as indicated by the color bar. The signal travels from top right to bottom left of the electrode array. (A) Before liAP induction, (B) 1 min after liAP induction. (C) 4 min after liAP induction. The Flash symbol indicates the laser induction point at electrode 64. Note that no difference in the overall propagation direction is visible. Black rectangles indicate invalid data. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/64355/64355fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/64355/64355fig3v2.jpg" /> 
Figure 3: FP/liAP parameter definition and recording protocol. (A) Parameters that can be extracted from the classical FP. (B) Additional parameters that can be obtained from liAPs. (C) Timeline of drug measurement. From left to right: control recording for 60 s, induction of liAP, recording of liAP for 60 s, drug application, wash-in time 300 s, re-induction of liAP, recording of liAP for 60 s. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/64355/64355fig3v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/64355/64355fig4v2.jpg" /> 
Figure 4: Nifedipine shortens the cardiac liAP in a concentration-dependent manner. (A) Top: FP traces at control (blue) and in the presence of 0.3 &#181;M Nifedipine (red). Traces are displayed with a y-axis offset for better visualization. Bottom: liAP traces from the same MEA recording, resulting in a shortening of the liAP combined with an increase in the beat rate. (B) Superposition of single liAPs during control (blue) and application of different concentrations of Nifedipine (red). a: 0.01 &#181;M, b: 0.1 &#181;M, and c: 0.3 &#181;M. Note the shortening of the liAP duration with increasing concentrations. (C) The concentration-response relationship of signal width obtained from FP (black) and liAP (red) recordings. Data is from n = 3 experiments and normalized to control. Error bars indicate the standard error of the mean. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/64355/64355fig4v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/64355/64355fig5v2.jpg" /> 
Figure 5: E4031 induces (pro-) arrhythmic behavior. (A) FP (top) and liAP (bottom) at control conditions. (B-C) FPs and liAPs were recorded at different time points after applying E4031 (0.1 &#181;M). After 80 s, the first EADs are visible at the end of the liAP (B; marked by a red arrow). EADs convert into ectopic beats after 320 s in the presence of the test compound (C). (D) After 530 s, the cardiac syncytium enters a tachycardic state. Trace depicted from liAP recording. (E) Concentration-response relationship of FP (black) and liAP (red) width. Data from n = 4 experiments, normalized to control. Error bars indicate the standard error of the mean. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/64355/64355fig5v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/64355/64355fig06.jpg" /> 
Figure 6: Detection of proarrhythmic events is more sensitive in liAPs than FPs. (A) FP recording and (B) liAP recording within the same well during application of 3 &#181;M Dofetilide. While at the end of some liAPs, EADs are detectable, they remain undiscoverable in FP recordings. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/64355/64355fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Laser-Induced Action Potential-Like Measurements of Cardiomyocytes on Microelectrode Arrays for Increased Predictivity of Safety Pharmacology at JoVE.com

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

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.

Life-threatening drug-induced cardiac arrhythmia is often preceded by prolonged cardiac action potentials (AP), commonly accompanied by small ...

laser-induced-action-potential-like-measurements-cardiomyocytes-on

Universidad de Cartagena UDEC

Research

JoVE Journal

Medicine

1.9K Views. 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.

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

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 ...

Preclinical Cardiac Electrophysiology Assessment by Dual Voltage and Calcium Optical Mapping of Human Organotypic Cardiac Slices

Human cardiac slice preparations have recently been developed as a platform for human physiology studies and therapy testing to bridge the gap between animal and clinical trials. Numerous animal and cell models have been used to examine the effects of drugs, yet these responses often differ in humans. Human cardiac slices offer an advantage for drug testing in that they are directly derived from viable human hearts. In addition to having preserved multicellular structures, cell-cell coupling, and extracellular matrix environments, human cardiac tissue slices can be used to directly test the effect of innumerable drugs on adult human cardiac physiology. What distinguishes this model from other heart preparations, such as whole hearts or wedges, is that slices can be subjected to longer-term culture. As such, cardiac slices allow for studying the acute as well as chronic effects of drugs. Furthermore, the ability to collect several hundred to a thousand slices from a single heart makes this a high-throughput model to test several drugs at varying concentrations and combinations with other drugs at the same time. Slices can be prepared from any given region of the heart. In this protocol, we describe the preparation of left ventricular slices by isolating tissue cubes from the left ventricular free wall and sectioning them into slices using a high precision vibrating microtome. These slices can then either be subjected to acute experiments to measure baseline cardiac electrophysiological function or cultured for chronic drug studies. This protocol also describes dual optical mapping of cardiac slices for simultaneous recordings of transmembrane potentials and intracellular calcium dynamics to determine the effects of the drugs being investigated.

This protocol describes the procedure for sectioning and culturing human cardiac slices for preclinical drug testing and details the use of optical mapping for recording transmembrane voltage and intracellular calcium signals simultaneously from these slices.

Human cardiac slice preparations have recently been developed as a platform for human physiology studies and therapy testing to bridge the gap between ...

Measuring Fast Calcium Fluxes in Cardiomyocytes

Cardiomyocytes have multiple Ca2+ fluxes of varying duration that work together to optimize function 1,2. Changes in Ca2+ activity in response to extracellular agents is predominantly regulated by the phospholipase C&beta;- G&alpha;q pathway localized on the plasma membrane which is stimulated by agents such as acetylcholine 3,4. We have recently found that plasma membrane protein domains called caveolae5,6 can entrap activated G&alpha;q 7. This entrapment has the effect of stabilizing the activated state of G&alpha;q and resulting in prolonged Ca2+ signals in cardiomyocytes and other cell types8. We uncovered this surprising result by measuring dynamic calcium responses on a fast scale in living cardiomyocytes. Briefly, cells are loaded with a fluorescent Ca2+ indicator. In our studies, we used Ca2+ Green (Invitrogen, Inc.) which exhibits an increase in fluorescence emission intensity upon binding of calcium ions. The fluorescence intensity is then recorded for using a line-scan mode of a laser scanning confocal microscope. This method allows rapid acquisition of the time course of fluorescence intensity in pixels along a selected line, producing several hundreds of time traces on the microsecond time scale. These very fast traces are transferred into excel and then into Sigmaplot for analysis, and are compared to traces obtained for electronic noise, free dye, and other controls. To dissect Ca2+ responses of different flux rates, we performed a histogram analysis that binned pixel intensities with time. Binning allows us to group over 500 traces of scans and visualize the compiled results spatially and temporally on a single plot. Thus, the slow Ca2+ waves that are difficult to discern when the scans are overlaid due to different peak placement and noise, can be readily seen in the binned histograms. Very fast fluxes in the time scale of the measurement show a narrow distribution of intensities in the very short time bins whereas longer Ca2+ waves show binned data with a broad distribution over longer time bins. These different time distributions allow us to dissect the timing of Ca2+fluxes in the cells, and to determine their impact on various cellular events.

We present a method to isolate rapid (microsecond) calcium events from slower fluxes in living cells using laser scanning confocal microscopy. The method measures fluorescence intensity fluctuations of calcium indicators by recording line scans of several hundred pixels in a cell. Histogram analysis allows us to isolate the time scales of different calcium fluxes.

Cardiomyocytes have multiple Ca2+ fluxes of varying duration that work together to optimize function 1,2. Changes in Ca2+ activity in response to ...

Biology

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 ...

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 ...

A Cardiac Microphysiological System for Studying Ca2+ Propagation via Non-genetic Optical Stimulation

In vitro cardiac microphysiological models are highly reliable for scientific research, drug development, and medical applications. Although widely accepted by the scientific community, these systems are still limited in longevity due to the absence of non-invasive stimulation techniques. Phototransducers provide an efficient stimulation method, offering a wireless approach with high temporal and spatial resolution while minimizing invasiveness in stimulation processes. In this manuscript, we present a fully optical method for stimulating and detecting the activity of an in vitro cardiac microphysiological model. Specifically, we fabricated engineered laminar anisotropic tissues by seeding human-induced pluripotent stem cell-derived cardiomyocytes (hiPSC-CMs) generated in a 3D bioreactor suspension culture. We employed a phototransducer, an amphiphilic azobenzene derivative, named Ziapin2, for stimulation and a Ca2+ dye (X-Rhod 1) for monitoring the system&#39;s response. The results demonstrate that Ziapin2 can photomodulate Ca2+ responses in the employed system without compromising tissue integrity, viability, or behavior. Furthermore, we showed that the light-based stimulation approach offers a similar resolution compared to electrical stimulation, the current gold standard. Overall, this protocol opens promising perspectives for the application of Ziapin2 and material-based photostimulation in cardiac research.

Using light to control cardiac cells and tissue enables non-contact stimulation, thereby preserving the natural state and function of the cells, making it a valuable approach for both basic research and therapeutic applications.

In vitro cardiac microphysiological models are highly reliable for scientific research, drug development, and medical applications. Although widely ...

Microelectrode Array Recording of Sinoatrial Node Firing Rate to Identify Intrinsic Cardiac Pacemaking Defects in Mice

The sinoatrial node (SAN), located in the right atrium, contains the pacemaker cells of the heart, and dysfunction of this region can cause tachycardia or bradycardia. Reliable identification of cardiac pacemaking defects requires the measurement of intrinsic heart rates by largely preventing the influence of the autonomic nervous system, which can mask rate deficits. Traditional methods to analyze intrinsic cardiac pacemaker function include drug-induced autonomic blockade to measure in vivo heart rates, isolated heart recordings to measure intrinsic heart rates, and sinoatrial strip or single-cell patch-clamp recordings of sinoatrial pacemaker cells to measure spontaneous action potential firing rates. However, these more traditional techniques can be technically challenging and difficult to perform. Here, we present a new methodology to measure intrinsic cardiac firing rate by performing microelectrode array (MEA) recordings of whole-mount sinoatrial node preparations from mice. MEAs are composed of multiple microelectrodes arranged in a grid-like pattern for recording in vitro extracellular field potentials. The method described herein has the combined advantage of being relatively faster, simpler, and more precise than previous approaches for recording intrinsic heart rates, while also allowing easy pharmacological interrogation.

This protocol aims to describe a new methodology to measure intrinsic cardiac firing rate using microelectrode array recording of the whole sinoatrial node tissue to identify pacemaking defects in mice. Pharmacological agents can also be introduced in this method to study their effects on intrinsic pacemaking.

The sinoatrial node (SAN), located in the right atrium, contains the pacemaker cells of the heart, and dysfunction of this region can cause tachycardia or ...

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 ...