1. Preparation of solutions and materials (see Table of Materials)
<ol>
	<li>6-well tissue-culture plate substrate-coating
	<ol>
		<li>Thaw the coating substrate on ice or at 4 &#176;C.</li>
		<li>Prepare a 1:100 coating substrate dilution in cold DMEM/F12 medium. Mix the solution by slow pipetting.</li>
		<li>Transfer 2 mL of the coating substrate solution (step 1.1.2) per well of a 6-well tissue culture plate.</li>
		<li>Immediately place the coated tissue culture plate in the cell culture incubator at 37 &#176;C and 5% CO2 for at least 7 h and use within 7 days.</li>
	</ol>
	</li>
	<li>hiPSC-CM culture medium
	<ol>
		<li>Add 10 mL of CM media supplement thawed at 4 &#176;C to 500 mL of CM base medium. Store at 4 &#176;C for up to 2 weeks.</li>
		<li>Aliquot required amount of media for the day and bring to room temperature before use.</li>
	</ol>
	</li>
	<li>hiPSC-CM thawing medium: Prepare fresh by mixing hiPSC-CM culture medium (step 1.2) with 10% fetal bovine serum (FBS). Bring the media to room temperature prior to thawing of CMs for suspension.</li>
	<li>Fibronectin 1 mg/mL stock solution: Aliquot 200 &#181;L into 1.5 mL sterile microfuge tubes and stored at 4 &#176;C for later use. Prepare working solution of 50 &#181;g/mL concentration freshly on ice.</li>
	<li>Multiwell cleaning solution: Combine 0.5 g of enzymatic detergent with 50 mL of sterile double distilled water (ddH2O). Vortex to mix the contents. Filter and store at 4 &#176;C for up to a week.</li>
</ol>
2. Pre-plating of cryopreserved hiPSC-CM for maturation (Figure 1)
NOTE: This section is intended for thawing and culturing hiPSC-CMs that were differentiated using the feeder-free monolayer method3,16 and cryopreserved in liquid nitrogen 10 days post-differentiation at 1-2 million cells/vial. Cells from one vial are plated into two substrate-coated wells of a 6-well tissue culture plate. Cardiomyocytes tend to settle at the bottom of the tube so gentle mixing at the time of pre-plating is important for achieving even cell density across wells.
<ol>
	<li>Aliquot 2 mL of FBS per vial of hiPSC-CMs being thawed into a 15 mL conical tube and bring to room temperature.</li>
	<li>Thaw vials of cryopreserved hiPSC-CMs by placing them in a 37 &#176;C water bath and swirl gently for even thawing for no more than 3 min.</li>
	<li>Immediately transfer the vial contents to the tube containing FBS (see step 2.1), mix by swirling and centrifuge at 200 x g for 5 min.</li>
	<li>Aspirate the supernatant and resuspend the cell pellet in 1 mL of hiPSC-CM thawing medium (see step 1.3) per vial thawed. Use a transfer pipette to resuspend the pellet by gentle trituration. Assess cell viability.</li>
	<li>Add additional 3 mL of thawing medium per vial of hiPSC-CM thawed in step 2.4 and suspend gently using a transfer pipette to further dissociate cell clumps.</li>
	<li>Dispense 2 mL of cell suspension gently into each well of substrate-coated 6-well plates (see step 1.1). Place in cell culture incubator at 37 &#176;C and 5% CO2.</li>
	<li>Replace with fresh hiPSC-CM culturing medium after 24 h and 3 times weekly thereafter for 20 days. 
	NOTE: Cells should adhere to the substrate coating by 24 h and beat spontaneously at 48 h post-plating (See Video 1 and Video 2).</li>
</ol>
3. Multiwell MEA plate sterilization and coating (Figure 2 and Figure 3)
NOTE: The protocol described here is for preparing 24-well MEA plates with 12 micro gold PEDOT-coated electrodes on glass for hiPSC-CM plating. Avoid touching the bottom of the plate as this may damage the electrodes.
<ol>
	<li>Two days prior to cell plating, add 0.5 mL of hiPSC-CM culture medium (see step 1.2) to each well and perform a baseline recording to verify signal-to-noise ratio for quality check of the MEAs.</li>
	<li>Aspirate the media, rinse with sterile ddH2O and sterilize under UV light inside a laminar flow hood overnight.</li>
	<li>The day prior to cell plating, add 0.1 mL of FBS to each well for hydrophilic treatment of MEA surfaces. Incubate for 30 min at room temperature. This step is necessary for cell attachment.</li>
	<li>Aspirate the FBS and rinse with 0.5 mL of sterile ddH2O per well. Repeat one more time.</li>
	<li>Leave the plate to dry in the laminar flow hood overnight.</li>
	<li>Prepare a working dilution of 50 &#181;g/mL fibronectin in cold DMEM/F12 medium from the stock (see step 1.4). Keep the solution on ice.</li>
	<li>Pipette 5 &#181;L of the working fibronectin dilution (see 3.6) and carefully dispense the droplet to the center of each well to cover all 12 electrodes. It is important to work swiftly across the 24 wells to prevent the droplet from drying.</li>
	<li>Immediately place the fibronectin-coated multiwell MEA plate on a raised surface inside a humidifying chamber containing sterile ddH2O to cover the entire dish surface. Place the chamber with the multiwell MEA plate for 3 h in the cell culture incubator. 
	NOTE: Placing the 24 well MEA plate in a humidifying chamber is critical to prevent fibronectin droplets from drying out during the incubation period.</li>
</ol>
4. hiPSC-CM dissociation and plating on Multiwell MEA plate (Figure 3)
NOTE: Start this step about 1 h prior to MEA fibronectin incubation is complete. Ensure that the cell dissociation solution is at 37 &#176;C and the iPSC-CM thawing medium is at room temperature. Dissociation methods have been optimized for 30 days post-differentiated hiPSC-CMs cultured on substrate-coated 6-well plates (see step 2) to obtain about 90% viable CMs for MEA plating. Care should be taken not to introduce air bubbles while trituration to prevent cell death.
<ol>
	<li>Aspirate the culture medium from each well of the 6-well tissue culture dish with 30 day post-differentiated hiPSC-CM culture (see step 2.7) and wash with 2 mL of sterile D-PBS per well.</li>
	<li>Add 1 mL of pre-warmed cell dissociation solution (see Table of Materials) per well and incubate for 4 min at 37 &#176;C. Use a transfer pipette to gently triturate to loosen cells for dissociation. If most of the cells are still adherent, incubate for another 3 min at 37 &#176;C and triturate again. This additional incubation should help for maximal cell recovery. Do not incubate for more than 7 min as this might result in low cell viability.</li>
	<li>Using a transfer pipette, pool all the dissociated cells into a conical tube containing hiPSC-CM thawing medium (see step 1.3). It is recommended to suspend the cells in at least twice the volume of media (2 mL per well being harvested) to block the activity of the cell dissociation solution.</li>
	<li>Centrifuge at 200 x g for 5 min.</li>
	<li>Aspirate the solution carefully as the pellet is loosely attached to the surface and resuspend in 0.1 mL of hiPSC-CM thawing medium. Use a transfer pipette to resuspend the pellet a few times by gentle trituration.</li>
	<li>Aliquot 2 &#181;L of dissociated cells and dilute with 18 &#181;L of media in a 1.5 mL microfuge tube. Add 20 &#181;L of Trypan Blue and mix for cell count and viability assessment.</li>
	<li>Adjust the cell density to 6,000 cells/&#181;L by adding the appropriate volume of hiPSC-CM thawing medium. Suspend the pellet by gentle flicking a few times. The hiPSC-CMs easily detach from glass surfaces especially when plated at high densities. We have optimized the seeding density and plating conditions to achieve 15 days of electrical recordings.</li>
	<li>When ready, bring the multiwell MEA plate into the laminar flow hood for cell seeding. It is critical to perform the following two steps one well at a time to prevent the fibronectin from drying.
	<ol>
		<li>Carefully remove the fibronectin droplet using a P10 pipette without touching the electrodes.</li>
		<li>Immediately dispense a 5 &#181;L cell droplet (30,000 cells) to the center of the well of the MEA plate covering all 12 electrodes. Repeat the step until all the 24 wells are plated. Intermittent mixing of cell suspension by flicking is recommended.</li>
	</ol>
	</li>
	<li>Place the multiwell MEA plate back in the loosely covered humidifying chamber and return to the cell culture incubator at 37 &#176;C and 5% CO2 for 3 h for cell attachment.</li>
	<li>Carefully add 200 &#181;L of hiPSC-CM thawing medium to each well without disturbing the cells&#160;using a P200 pipette. Add the media dropwise to the side of the well.</li>
	<li>Place the multiwell MEA plate back in the cell culture incubator.</li>
	<li>Replace with fresh hiPSC-CM culture medium at 24 h post plating. Spontaneous beating of cardiomyocytes can be observed at this point (See Video 3).</li>
	<li>Change the media every 2 days until the end of experiments.</li>
</ol>
5. hiPSC-CM electroporation and signal acquisition (Figures 4 &#8211; 6)
NOTE: This protocol is for simultaneous recording of high-throughput electrode signals (12 sites for each of the 24 wells). The 24-well multiwell MEA system is used with the acquisition software (see Table of Materials). All MEA recordings are conducted at 37 &#176;C.
<ol>
	<li>Turn on the interface board and initiate the acquisition software. Allow enough time for the multiwell MEA headstage to reach 37 &#176;C temperature (see arrow no. 1 in Figure 4).</li>
	<li>Insert multiwell MEA plate from step 4 into the multiwell MEA headstage by placing it covered over the recording platform and click the Insert button (see arrow no. 2 in Figure 4). Allow for temperature to stabilize before starting recordings.</li>
	<li>Adjust acquisition and electroporation settings
	<ol>
		<li>Click on the Define Experimental Flow icon (see arrow no. 3 in Figure 4) and set recording time to 2 min or as desired.</li>
		<li>Click on the Data Acquisition Setup icon (see arrow no. 4 in Figure 4) and set sampling rate to 20 kHz, high-pass filter to 0.1 Hz and low-pass filter to 3500 Hz.</li>
		<li>Click on the Stimulator Settings icon (see Figure 5). Under the Stimulus Definition tab, define stimulation as biphasic symmetric voltage pulses of 1 mV, 1 ms and 1 Hz. Under the Stimulation Electrodes tab, select electroporation sites by highlighting all relevant electrodes.</li>
	</ol>
	</li>
	<li>Click the Explore button to visualize signals in all wells. Verify signal quality and steady state conditions. Take notes on electrodes with FP signals in the mV range. Click the same button to stop the exploration. No data has been recorded up to this point.</li>
	<li>Start recording by clicking on the Go! button. The electrodes in each well will show FP signals in the raw data window (Figure 6). After 30 s of recording, click on the Stimulate button and allow electroporation to take place on the selected sites for 30 s; then, click on the same button to stop stimulation and continue recording for the remaining 60 s. 
	NOTE: The recorded file can be played back by switching to &#8216;Replayer Mode&#8217; in the &#8216;Application&#8217; dropdown menu.</li>
</ol>
6. Multiwell MEA plate cleaning for reuse
<ol>
	<li>After final experiments, clean all the wells in the multiwell plate by aspirating all media contents from each well and carefully avoiding touching the electrode surface.</li>
	<li>Add 1 mL of sterile ddH2O per well. Aspirate and repeat once.</li>
	<li>Add 0.3 mL of multiwell cleaning solution (see step 1.5) per well. Incubate overnight at room temperature to dislodge cells and debris.</li>
	<li>The following morning, aspirate the solution and rinse with 1 mL of sterile ddH2O. Incubate for 5 - 7 min and aspirate. Repeat 5 times.</li>
	<li>Add 0.5 mL of sterile ddH2O per well. Record the baseline of the cleaned multiwell plate for quality check of the cleaned MEAs (Figure 7).</li>
	<li>Store at 4 &#176;C until ready to use.</li>
</ol>
7. Data file conversion and export
NOTE: Four data files will be generated for every recording: MWR, MWC, MWD, and MWS files. Using the converter software, the MWD file can be converted to H5 file for subsequent analysis using custom-built script (see Supplementary File 1).
<ol>
	<li>Initiate the converter software (see Table of Materials).</li>
	<li>Select Set Input Path from the File menu. Select folder containing data files of interest.</li>
	<li>Select Set Output Path from the File menu. Select folder where converted files are to be saved.</li>
	<li>Highlight MWD file of interest.</li>
	<li>Click the EXPORT to HDF5 button.</li>
</ol>
8. Data segmentation and analysis (Figures 8-10)
NOTE: Matlab-based custom software is used to segment and extract various FP and AP data parameters. Software is available on demand.
<ol>
	<li>Run the Waveform Analysis code using Matlab (see Figure 8 for a view of the GUI&#8217;s main window).</li>
	<li>Click on File and select Process .h5.</li>
	<li>Find and select the mwd.h5 file created according to step 7 above.</li>
	<li>Click on the Save Directory button to change the storage location of the output files.</li>
	<li>Create a signal processing queue by selecting electrode/well combinations of interest and then clicking on the Queue button. Repeat this step to append more electrode/well combinations to be processed to the queue.</li>
	<li>Edit the queue by clicking directly on Med Name / Med Concentration if cells were treated with drugs (Figure 8).</li>
	<li>Once the queue is final, click the Initialize Waveforms button. This will start the preliminary processing in which signals are identified and extracted for segmentation.</li>
	<li>Click on the Zoom In button and select the action potential area of interest with the cursor (Figure 9).</li>
	<li>Click the Keep button and review the panels. Peaks (red &#8216;x&#8217;) and troughs (yellow circles) are detected for every waveform and the normalized action potentials are superimposed. Click the Keep button and move on to the next trace in the queue (Figure 10).</li>
	<li>Repeat steps 8.8-8.9 for the rest of the electrode/well combination signals in the queue. 
	NOTE: A .csv file will be generated with APD parameters measured for every waveform. A .mat file for each .h5 file is also saved to allow additional processing of segmented data.</li>
</ol>

The authors have nothing to disclose.

Over the years, the application of MEAs has been limited to carrying out FP measurements of excitable cells to study their electrophysiological properties36,37,38,39. Only a few groups have reported AP traces from electrogenic cells using custom MEA based technology18,29,30. However, these approaches have not been investigated for repeated recordings from the same preparations. We developed an innovative and accurate methodology for studying APs from the same cell site over days in multiple hiPSC-CM networks simultaneously3. In our published study, a multiwell micro-gold MEA platform was employed to generate AP waveform libraries from multiple batches of hiPSC-CM cultures with high precision and with a temporal resolution of 1 &#181;s. The protocol described here explains seeding of hiPSC-CMs on the array for efficient development of syncytial CM networks for high-throughput AP recordings. Several critical steps in the protocol are: 1) production of multiple high-purity batches of quality-controlled CMs for cryopreservation banking, 2) highly viable post-thaw CMs for pre-plating and maturation, 3) treatment of the multiwell MEA plate for CM seeding, 4) hiPSC-CM culture dissociation at 30 days post-differentiation for MEA plating, and 5) restoration of the MEAs for multiple reuse.
It is important to note that batch-to-batch variation in hiPSC differentiation might affect experimental outcomes. The monolayer method of differentiation was optimized in-house for high percent cardiomyocyte production3,40. The FACS analysis of MLC2v and TNNT2 markers of our cultures demonstrate a &#8805;90% ventricular-like phenotype3. These quality-controlled cultures are cryopreserved for experimental studies. The current differentiation approaches yield a heterogeneous mix of nodal-, atrial- and ventricle-like cells3,16,17,41. Therefore, strategies employed for CM subtype population enrichment can further improve the specificity of the cultures. Additionally, tissue engineering approaches can be employed to enhance their maturation. The methods proposed here can easily be implemented for other CM sources.
The AP waveforms recorded using MEA were similar to those recorded from networks of cardiomyocytes by optical mapping42,43, complementary metal oxide semiconductor-based MEA18,21, and simulated AP using FP recordings20. To address the mechanism of AP measurements via MEA Hai and Spira25 demonstrated that electropore-electrode interface mimic the established sharp glass microelectrode technique. However, the resting membrane potential and true amplitude values in our study cannot be established given that the electropore-electrode interface in MEA systems is not calibrated, and that the amplitude is a function of the sensitivity and resolution of the technique. Our approach shares similar limitations to optical mapping when it comes to AP amplitude.
The multiwell MEA-based FP/AP readouts reported here open new possibilities for drug safety assessment. Although spontaneous, these hiPSC-CM monolayers beat at constant rates. Analysis of APD parameters across multiple networks provide insight on electrical heterogeneity (Figure 13). However,comprehensive APD restitution analyses must incorporate preceding diastolic intervals. Furthermore, high-quality AP waveforms recorded from the same cell site over 96 h (Figure 11B) is the first report to track membrane electrodynamics over time which will be of value in development and in disease.
The protocol described here for quantifying AP parameters can be used to generate dose-response curves to test compounds. As recently reported by Edwards et al.3, dose response of norepinephrine, isoproterenol and E 4031 are plotted for APD at various repolarization phases. The published study demonstrated the accuracy and reliability of the approach for identification of the dose-dependent subtle changes in the AP waveforms in real time. This technique could easily be extended for other compounds or small molecule libraries for understanding various electrophysiological responses.
The MEA based approach for AP measurements presented in this study will be of interest not only to electrophysiologists but also to cell biologists and in-silico modelers. Furthermore, FP/AP recordings from the same cell site on hiPSC-CMs will enable researchers to generate bioelectric data libraries of wide array of excitable cellular networks within a short period of time. Availability of these resources will be valuable for drug discoveries and disease modeling.

Human induced pluripotent stem cell-derived cardiomyocytes (hiPSC-CMs) are the gold standard for an increasing number of laboratories1,2,3,4,5,6,7,8,9,10. Beating embryoid bodies11,12,13 and monolayer3,7,10,11,12,13,14,15,16,17 differentiation are the preferred methods for cardiomyocyte production and the multielectrode array (MEA) has become a common modality for monitoring the electrodynamics of these networks18,19,20. While parameters that can be extracted from field potentials (FPs) such as beating rate, amplitude, duration and RR intervals are baseline electrophysiological responses of spontaneously beating monolayers18,21,22,23, the action potential (AP) components underlying these extracellular FP signals are difficult to extrapolate24. Our recent publication on the discovery of an application of MEAs for direct recurrent AP measurements provides proof of methodology for exemplary intracellular AP readouts with an extensive waveform analysis at various repolarization phases across multiple batches of hiPSC-derived cardiomyocyte networks3. In the study we demonstrated that the delivery of electroporating pulses to networks of hiPSC-derived cardiomyocytes enables intracellular access for AP recordings. These transient AP recordings are dependent on transmembrane potential recoveries observed through the injury site3,25,26. Waveforms recorded via MEA and patch- clamp in our study showed similar AP morphologies thus validating the reliability of the approach3.
A few laboratories have reported measuring APs from various electrogenic cells using custom-built MEAs18,21,26,27,28,29,30, but the reliability of using MEAs for consistent and recurrent AP measurements was not assessed. Currently, the gold standard patch-clamp technique is limited to terminal recordings7,31 whereas, MEA-based AP measurements are transient and therefore can be conducted multiple times on the same cell. We also show that one can easily record high-quality AP signals in the millivolt range requiring minimal filtering. Researchers can therefore conduct not only acute but also chronic drug studies in the same preparations using MEAs. Additionally, this technology permits simultaneous FP/AP measurement generating electro-biome libraries in a short period of time. Given the growing emphasis on arrhythmia prediction and drug associated cardiotoxicity24,32,33,34,35, integration of AP measurement approaches will enhance drug safety and efficacy assessments.
Here, we present protocols for 1) pre-plating of cryopreserved hiPSC-CMs for maturation, 2) dissociating and plating of hiPSC-CMs on multiwell MEAs, 3) recording of FPs and APs from hiPSC-CM networks, 4) segmenting and extracting the data for analysis, and 5) restoring the arrays for multiple reuse. Each step has been optimized emphasizing critical steps wherever relevant. Requirements for cell attachment to ensure a beating syncytial monolayer are discussed and procedures for multiwell MEA restoration for repetitive electrophysiological studies are explained. Finally, a custom GUI developed in the laboratory is presented for AP signal extraction, quality assurance, and segmentation workflow to quantify and analyze AP parameters.

<table>
	<tbody>
		<tr>
			<td>Accutase</td>
			<td>Sigma Aldrich</td>
			<td>A6964-100ML</td>
			<td>cell dissociation solution</td>
		</tr>
		<tr>
			<td>Acquisition software</td>
			<td>Multichannel Systems</td>
			<td>Multiwell-Screen v 1.9.2.0</td>
			<td></td>
		</tr>
		<tr>
			<td>B27 Supplement</td>
			<td>ThermoFisher</td>
			<td>17504-044</td>
			<td>CM media supplement</td>
		</tr>
		<tr>
			<td>Converter software</td>
			<td>Multichannel Systems</td>
			<td>MultiChannel DataManager</td>
			<td></td>
		</tr>
		<tr>
			<td>DMEM/F12</td>
			<td>ThermoFisher</td>
			<td>11330-032</td>
			<td></td>
		</tr>
		<tr>
			<td>D-PBS</td>
			<td>ThermoFisher</td>
			<td>14190-250</td>
			<td></td>
		</tr>
		<tr>
			<td>FBS</td>
			<td>Fisher Scientific</td>
			<td>SH3007103HI</td>
			<td></td>
		</tr>
		<tr>
			<td>Fibronectin</td>
			<td>Sigma Aldrich</td>
			<td>F1141-5MG</td>
			<td></td>
		</tr>
		<tr>
			<td>Geltrex</td>
			<td>ThermoFisher</td>
			<td>A1413202</td>
			<td>coating substrate</td>
		</tr>
		<tr>
			<td>Interface board</td>
			<td>Multichannel Systems</td>
			<td>MCS-IFB 3.0 Multiboot Interface Board</td>
			<td></td>
		</tr>
		<tr>
			<td>Multiwell MEA Plate</td>
			<td>Multichannel Systems</td>
			<td>24W300/30G-288</td>
			<td></td>
		</tr>
		<tr>
			<td>RPMI 1640</td>
			<td>ThermoFisher</td>
			<td>11875-093</td>
			<td>CM base medium</td>
		</tr>
		<tr>
			<td>Terg-a-zyme</td>
			<td>Sigma Aldrich</td>
			<td>Z273287-1EA</td>
			<td>enzymatic detergent</td>
		</tr>
		<tr>
			<td>Transfer pipettes, individually wrapped</td>
			<td>Fisher Scientific</td>
			<td>1371148</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypan Blue</td>
			<td>Sigma Aldrich</td>
			<td>T8154-100ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultrapure sterile water</td>
			<td>ThermoFisher</td>
			<td>10977-023</td>
			<td></td>
		</tr>
		<tr>
			<td>6-well tissue-culture treated plates</td>
			<td>Fisher Scientific</td>
			<td>08-772-1B</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

The viability and plating density of post-thawed hiPSC-CMs is critical for multiwell MEA culture. Pre-plating of 1-2 million hiPSC-CMs/vial into two wells of a 6-well tissue culture plate with 50% or greater viability will produce a healthy monolayer culture with spontaneous beating at 48 h. Poor viability of CMs will result in cultures with a high percentage of non-myocyte populations. These monolayers when dissociated for multiwell MEA plating generally produce inconsistent results and bad quality signals and therefore should be discarded. Figure 1 shows examples of optimal vs. sub-optimal hiPSC-CMs cultures at 48 h post plating. Thawing the CMs on substrate-coated tissue culture plates rather than directly on multiwell MEAs, allows for cell recovery and maturation3. Direct plating of cryopreserved CMs on the array is not recommended as it produced inconsistent results.
In addition to the quality of the dissociated CMs, cell attachment on multiwell MEA is highly dependent on cell density and the fibronectin coating technique. The fibronectin droplet size is critical as the CMs will conform to the boundaries of the fibronectin-coated area. For this reason, only 5 &#181;L of the fibronectin solution are dispensed directly over the electrode array area. To ensure that the droplet does not disperse, the well surface must be completely dry at the time of coating. Figure 2 shows the layout of the multiwell MEA plate with schematics of step-by-step pretreatment for optimal preparation. Additionally, to prevent the fibronectin from drying the multiwell MEA plates must be placed inside a humidifying chamber during the incubation period lasting no more than 3 h (see step 3.8). Once the incubation period is complete, it is important to remove the fibronectin droplet from each well just before CM plating and only then proceed to next well plating. Working swiftly and carefully dispensing of the CMs is the key to successful cell attachment.
hiPSC-CM cultures at 30 days post-differentiation are dissociated for multiwell MEA plating using the enzymatic cell dissociation method (see step 4). CMs will attach to the fibronectin-coated MEA surfaces by 3 h and a monolayer covering the arrays will be visible after 24 h post-plating (Figure 3). Synchronous beating of the monolayer will be observed at 24-48 h. Cell droplet dispersion will affect the culture density or even lead to drying and cell death. Precise cell placement directly on the array is of utmost importance and therefore the technique must be practiced for optimal plating. Cell adhesion to the reference electrode will hinder electrical signal production. See Figure 3 for images of optimal CM placement, and culture after 24 h.
The CMs cultured on multiwell MEAs are subjected to quality check for electrical activity at 48 h post-plating. Typically, FP signal amplitude increases from the &#181;V range to mV in approximately 4 days3. If 50% of the electrodes within a network and 70% of the total networks do not produce FP signals, then the network or the culture are suboptimal and should be discarded. Only cultures that pass the quality check are processed for FP and AP analysis. Figure 6 shows examples of good and sub-standard FP signals.
Electroporation-mediated AP recordings can be obtained multiple times from cultures 48 h post-MEA plating. Employing electroporation, we gained intracellular access to record high-resolution APs from multiple hiPSC-derived cardiomyocyte networks. Low-voltage pulses (1 V, 1 ms, 1 Hz) for 30 s were delivered for transient, reversible transformation of FP to AP. The electroporation allows successful intracellular access for AP measurement in approximately 75% of the electrodes. Electrical signals are recorded for 2 min that include 30 s pre-electroporation, 30 s during and 1 min post-electroporation. A train of 10 s AP waveforms 10 s post-electroporation are evaluated across all sites for signal quality and analysis. Any trace not conforming to pure AP signal are discarded. To investigate if AP amplitudes correlate to FP signal we electroporated all 288 sites to simultaneously record waveforms. Representative FP and AP signals recorded from the same cell site from two different electrodes are shown in Figure 11A. We observed no correlation between FP amplitudes and post electroporation AP amplitudes recorded from the same cell site. Additionally, multiple electroporations of the same cell site at 0, 24, 48, 72 and 96 h had no significant effect on the AP shape over time (Figure 11B).
Given the high-throughput nature of the system, a manual technique to extract and quantify parameters of interest such as RR interval, instantaneous frequency and differential action potential duration is inefficient and time consuming. A custom-built MATLAB script available to the research community upon request is employed to perform waveform measurements with 1 &#181;s resolution. Electroporation time points are overlaid with the extracted signal to identify 10 s of AP post-electroporation to conduct signal extraction, quality assurance, and segmentation workflow (Figure 8, Figure 9, Figure 10). The user interface allows for selection of the desired segment using the overlaid electroporation indicators as a guide. The segmented waveform is processed by subroutines to further identify individual AP waveforms. This is completed through peak detection, where the highest and lowest voltage is identified for each cycle. Once this process is completed, the amplitudes are normalized, and the associating time vectors are shifted to define time zero at a peak value of 1. Interpolation of intersection points along the individual cycles was used to determine APD measurements. Thus, partial automation workflow for AP waveform segmentation allows efficient data analysis for various APD parameters across multiple batches of cultures in a short period of time. Further automation of inclusion and exclusion criteria for FPs and APs is ongoing for real-time data analysis.
A significant advantage of the multiwell MEA plate is that it can be reused multiple times. This restoration enables repetitive electrophysiological studies for cost-effective and consistent data collection. Recordings of APs from the same array after 6 restorations are shown in Figure 12. Signal-to-noise ratio is similar across multiple reuses. To demonstrate the reliability of the array for repetitive electrophysiological studies, a total of 3815 AP waveforms are pooled from three restoration batches and AP duration data is extracted to examine the repeatability of the results. Distribution plots for individual waveform APD30, APD80, triangulation (APD80&#8212;APD30) and fractional shortening ((APD80&#8212;APD30)/(APD80)) are displayed (Figure 13).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59906/59906fig1.jpg" /> 
Figure 1: Pre-plating of cryopreserved hiPSC-CM for maturation. (A) Cell processing for pre-plating 1 vial of 10 days post-differentiation cryopreserved hiPSC-CMs. (B) Phase contrast images of successful (left) and unsuccessful (right) hiPSC cultures. Scale bar: 275 &#181;m. See Video 1 and Video 2 for successful 14 and 24 days post-differentiation culture examples. <a href="https://www.jove.com/files/ftp_upload/59906/59906fig1large.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/59906/59906fig2.jpg" /> 
Figure 2: Multiwell MEA plate setup and preparation. (A) Multiwell MEA plate schematics: The plate consists of 24 wells (A1 through D6) each containing 12 microelectrode arrays and 4 peripheral reference electrodes. Electrode diameter: 30 &#181;m / Inter-electrode distance: 300 &#181;m. Recordings can be obtained from the 288 electrodes simultaneously. (B) Sterilization and hydrophilic treatment steps to be conducted prior to hiPSC-CM plating. <a href="https://www.jove.com/files/ftp_upload/59906/59906fig2large.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/59906/59906fig3.jpg" /> 
Figure 3: hiPSC-CM dissociation and plating on Multiwell MEA plate. (A) Schematics of hiPSC-CM MEA plating steps for each well. (B) Microscopic image illustrating correct cell droplet placement covering all 12 electrodes without spreading to the 4 reference electrodes. (C) Phase contrast microscopic images of an exemplary (left) and suboptimal (right) hiPSC-CM platting on MEA at 24 h post-plating. Scale bar= 275 &#181;m. See Video 3 for successful MEA plating example. <a href="https://www.jove.com/files/ftp_upload/59906/59906fig3large.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/59906/59906fig4.jpg" /> 
Figure 4: Multiwell-screen acquisition software. Arrows indicate the location of key features and functions referenced in the text: Temperature Control (1) panel allows for real time temperature monitoring throughout the experiment. Insert/Eject (2) button engage and release the Multiwell MEA Plate. Define Experimental Flow (3) function allows the user to set the duration of the recording. Data Acquisition Setup (4) function allows the user to set the sampling rate and acquisition filter settings. <a href="https://www.jove.com/files/ftp_upload/59906/59906fig4large.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/59906/59906fig5.jpg" /> 
Figure 5: hiPSC-CM electroporation and signal acquisition. Stimulus Definition tab allows the user to define the electroporating pulse parameters. Stimulation Electrodes tab allows the user to select the electroporating electrodes. Any combination of the 288 electrodes can be selected. <a href="https://www.jove.com/files/ftp_upload/59906/59906fig5large.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/59906/59906fig6.jpg" /> 
Figure 6: Quality check of Multiwell MEAs for electrical activity. Multiwell-Screen acquisition software showing raw data windows with representative examples of optimal (A) and sub-standard (B) FP signals. <a href="https://www.jove.com/files/ftp_upload/59906/59906fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/59906/59906fig7.jpg" /> 
Figure 7: FP and AP signals from the new and restored array. Multiwell MEA enzymatic cleaning steps (A). The baseline signal of the new array shows minimal signal to noise ratio (B) and FP signals show the electrical activity of the network (C). <a href="https://www.jove.com/files/ftp_upload/59906/59906fig7large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/59906/59906fig8.jpg" /> 
Figure 8: Data segmentation and analysis. View of GUI&#8217;s main window for Waveform Analysis. <a href="https://www.jove.com/files/ftp_upload/59906/59906fig8large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 9" class="xfigimg" src="/files/ftp_upload/59906/59906fig9.jpg" /> 
Figure 9: Data segmentation and analysis. Initialize Waveforms button to identify and extract AP waveforms for segmentation and to start the preliminary processing by zooming in and selecting the action potential area of interest. Red circles are the electroporation indicators. <a href="https://www.jove.com/files/ftp_upload/59906/59906fig9large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 10" class="xfigimg" src="/files/ftp_upload/59906/59906fig10.jpg" /> 
Figure 10: Data segmentation and analysis. Peaks (red &#8216;x&#8217;) and troughs (yellow circles) are detected for every waveform and the normalized APs are superimposed for a quality check of the waveforms. <a href="https://www.jove.com/files/ftp_upload/59906/59906fig10large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 11" class="xfigimg" src="/files/ftp_upload/59906/59906fig11.jpg" /> 
Figure 11: AP Amplitude dependence on the FP signal for multiple recordings from the same cell site. FP amplitude in &#181;V ranges (A, top left panel) or mV ranges (A, top right panel) recorded from two independent electrodes produce AP amplitude in mV range (A, bottom left and right panels) showing no correlation between FP amplitudes and post-electroporation AP amplitudes. The normalized AP waveforms for each recording are superimposed as shown for each recording. Multiple electroporations of the same cell site at 0 to 96 h produced high quality AP waveforms allowing tracking of membrane electrodynamics (B). <a href="https://www.jove.com/files/ftp_upload/59906/59906fig11large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 12" class="xfigimg" src="/files/ftp_upload/59906/59906fig12.jpg" /> 
Figure 12: AP recordings after six restorations. AP waveforms recorded simultaneously 10 s post-electroporation across 12 electrodes from the same well are displayed. <a href="https://www.jove.com/files/ftp_upload/59906/59906fig12large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 13" class="xfigimg" src="/files/ftp_upload/59906/59906fig13.jpg" /> 
Figure 13: APD parameter histograms from multiple restorations. Distribution plots for individual waveform APD30 (A), APD80 (B), triangulation (APD80&#8212;APD30) (C) and fractional shortening ((APD80&#8212;APD30)/(APD80)) (D) are displayed. <a href="https://www.jove.com/files/ftp_upload/59906/59906fig13large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary files. Videos 1-3. <a href="https://www.jove.com/files/ftp_upload/59906/Suppl_Files.zip">Please click here to download this file.</a>

Watch this Scientific Journal Video about Human iPSC-Derived Cardiomyocyte Networks on Multiwell Micro-electrode Arrays for Recurrent Action Potential Recordings at JoVE.com

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

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 ...

human-ipsc-derived-cardiomyocyte-networks-on-multiwell-micro

Research

JoVE Journal

Bioengineering

11.4K Views.  Advocate Aurora Health Care. 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.

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

Plasma Lithography Surface Patterning for Creation of Cell Networks

Systematic manipulation of a cell microenvironment with micro- and nanoscale resolution is often required for deciphering various cellular and molecular phenomena. To address this requirement, we have developed a plasma lithography technique to manipulate the cellular microenvironment by creating a patterned surface with feature sizes ranging from 100 nm to millimeters. The goal of this technique is to be able to study, in a controlled way, the behaviors of individual cells as well as groups of cells and their interactions.
This plasma lithography method is based on selective modification of the surface chemistry on a substrate by means of shielding the contact of low-temperature plasma with a physical mold. This selective shielding leaves a chemical pattern which can guide cell attachment and movement. This pattern, or surface template, can then be used to create networks of cells whose structure can mimic that found in nature and produces a controllable environment for experimental investigations. The technique is well suited to studying biological phenomenon as it produces stable surface patterns on transparent polymeric substrates in a biocompatible manner. The surface patterns last for weeks to months and can thus guide interaction with cells for long time periods which facilitates the study of long-term cellular processes, such as differentiation and adaption. The modification to the surface is primarily chemical in nature and thus does not introduce topographical or physical interference for interpretation of results. It also does not involve any harsh or toxic substances to achieve patterning and is compatible for tissue culture. Furthermore, it can be applied to modify various types of polymeric substrates, which due to the ability to tune their properties are ideal for and are widely used in biological applications. The resolution achievable is also beneficial, as isolation of specific processes such as migration, adhesion, or binding allows for discrete, clear observations at the single to multicell level.
This method has been employed to form diverse networks of different cell types for investigations involving migration, signaling, tissue formation, and the behavior and interactions of neurons arraigned in a network.

A versatile plasma lithography technique has been developed to generate stable surface patterns for guiding cellular attachment. This technique can be applied to create cell networks including those that mimic natural tissues and has been used for studying several, distinct cell types.

Systematic manipulation of a cell microenvironment with micro- and nanoscale resolution is often required for deciphering various cellular and molecular ...

Mass Cytometry: Protocol for Daily Tuning and Running Cell Samples on a CyTOF Mass Cytometer

In recent years, the rapid analysis of single cells has commonly been performed using flow cytometry and fluorescently-labeled antibodies. However, the issue of spectral overlap of fluorophore emissions has limited the number of simultaneous probes. In contrast, the new CyTOF mass cytometer by DVS Sciences couples a liquid single-cell introduction system to an ICP-MS.1 Rather than fluorophores, chelating polymers containing highly-enriched metal isotopes are coupled to antibodies or other specific probes.2-5 Because of the metal purity and mass resolution of the mass cytometer, there is no "spectral overlap" from neighboring isotopes, and therefore no need for compensation matrices. Additionally, due to the use of lanthanide metals, there is no biological background and therefore no equivalent of autofluorescence. With a mass window spanning atomic mass 103-203, theoretically up to 100 labels could be distinguished simultaneously. Currently, more than 35 channels are available using the chelating reagents available from DVS Sciences, allowing unprecedented dissection of the immunological profile of samples.6-7
Disadvantages to mass cytometry include the strict requirement for a separate metal isotope per probe (no equivalent of forward or side scatter), and the fact that it is a destructive technique (no possibility of sorting recovery). The current configuration of the mass cytometer also has a cell transmission rate of only ~25%, thus requiring a higher input number of cells.
Optimal daily performance of the mass cytometer requires several steps. The basic goal of the optimization is to maximize the measured signal intensity of the desired metal isotopes (M) while minimizing the formation of oxides (M+16) that will decrease the M signal intensity and interfere with any desired signal at M+16. The first step is to warm up the machine so a hot, stable ICP plasma has been established. Second, the settings for current and make-up gas flow rate must be optimized on a daily basis. During sample collection, the maximum cell event rate is limited by detector efficiency and processing speed to 1000 cells/sec. However, depending on the sample quality, a slower cell event rate (300-500 cells/sec) is usually desirable to allow better resolution between cells events and thus maximize intact singlets over doublets and debris. Finally, adequate cleaning of the machine at the end of the day helps minimize background signal due to free metal.

The steps necessary for daily tuning and optimization of the performance of a CyTOF mass cytometer are described. Comments on optimal sample preparation and flow rate are discussed

In recent years, the rapid analysis of single cells has commonly been performed using flow cytometry and fluorescently-labeled antibodies. However, the ...

Production, Characterization and Potential Uses of a 3D Tissue-engineered Human Esophageal Mucosal Model

The incidence of both esophageal adenocarcinoma and its precursor, Barrett&#8217;s Metaplasia, are rising rapidly in the western world. Furthermore esophageal adenocarcinoma generally has a poor prognosis, with little improvement in survival rates in recent years. These are difficult conditions to study and there has been a lack of suitable experimental platforms to investigate disorders of the esophageal mucosa.
A model of the human esophageal mucosa has been developed in the MacNeil laboratory which, unlike conventional 2D cell culture systems, recapitulates the cell-cell and cell-matrix interactions present in vivo and produces a mature, stratified epithelium similar to that of the normal human esophagus. Briefly, the model utilizes non-transformed normal primary human esophageal fibroblasts and epithelial cells grown within a porcine-derived acellular esophageal scaffold. Immunohistochemical characterization of this model by CK4, CK14, Ki67 and involucrin staining demonstrates appropriate recapitulation of the histology of the normal human esophageal mucosa.
This model provides a robust, biologically relevant experimental model of the human esophageal mucosa. It can easily be manipulated to investigate a number of research questions including the effectiveness of pharmacological agents and the impact of exposure to environmental factors such as alcohol, toxins, high temperature or gastro-esophageal refluxate components. The model also facilitates extended culture periods not achievable with conventional 2D cell culture, enabling, inter alia, the study of the impact of repeated exposure of a mature epithelium to the agent of interest for up to 20 days. Furthermore, a variety of cell lines, such as those derived from esophageal tumors or Barrett&#8217;s Metaplasia, can be incorporated into the model to investigate processes such as tumor invasion and drug responsiveness in a more biologically relevant environment.

This manuscript describes the production, characterization and potential uses of a tissue engineered 3D esophageal construct prepared from normal primary human esophageal fibroblast and squamous epithelial cells seeded within a de-cellularized porcine scaffold. The results demonstrate the formation of a mature stratified epithelium similar to the normal human esophagus.

The incidence of both esophageal adenocarcinoma and its precursor, Barrett&#8217;s Metaplasia, are rising rapidly in the western world. Furthermore ...

Easy and Accurate Mechano-profiling on Micropost Arrays

Cell culture substrates with integrated flexible microposts enable a user to study the mechanical interactions between cells and their immediate surroundings. Particularly, cell-substrate interactions are the main interest. Today micropost arrays are a well-characterized and established method with a broad range of applications that have been published over the last decade. However, there seems to be a reservation among biologists to adapt the technique due to the lengthy and challenging process of micropost manufacture along with the lack of easily approachable software for analyzing images of cells interacting with microposts. 
The force read-out from microposts is surprisingly easy. A micropost acts like a spring with the cell ideally attached at its tip. Depending on size a cell applies force from its cytoskeleton through one or multiple focal adhesion points to the micropost, thus deflecting the micropost. The amount of deflection correlates directly to the applied force in direction and in magnitude. The number of microposts covered by a cell and the post deflection patterns are characteristic and allow determination of values like force per post and many biologically relevant parameters that allow &#8220;mechano-profiling&#8221; of cell phenotypes.
A convenient method for mechano-profiling is described here combining the first generation of ready-to-use commercially available microposts with an in-house developed software package that is now accessible to all researchers. As a demonstration of typical application, single images of bone cancer cells were taken in bright-field microscopy for mechano-profiling of cell line models of metastasis. This combination of commercial traction force sensors and open source software for analysis allows for the first time a rapid implementation of the micropost array technique into routine lab work done by non-expert users. Furthermore, a robust and streamlined analysis process enables a user to analyze a large number of micropost images in a highly time-efficient manner.

Described are protocols for quantifying mechanical interactions between adherent cells and microstructured substrates. These interactions are closely linked to essential cell behaviors including migration, proliferation, differentiation, and apoptosis. The protocols present an open-source image analysis software called MechProfiler, which enables determination of involved forces for each micropost.

Cell culture substrates with integrated flexible microposts enable a user to study the mechanical interactions between cells and their immediate ...

Creating a Structurally Realistic Finite Element Geometric Model of a Cardiomyocyte to Study the Role of Cellular Architecture in Cardiomyocyte Systems Biology

With the advent of three-dimensional (3D) imaging technologies such as electron tomography, serial-block-face scanning electron microscopy and confocal microscopy, the scientific community has unprecedented access to large datasets at sub-micrometer resolution that characterize the architectural remodeling that accompanies changes in cardiomyocyte function in health and disease. However, these datasets have been under-utilized for&#160;investigating the role of cellular architecture remodeling in cardiomyocyte function. The purpose of this protocol is to outline how to create an accurate finite element model of a cardiomyocyte using high resolution electron microscopy and confocal microscopy images. A detailed and accurate model of cellular architecture has significant potential to provide new insights into cardiomyocyte biology, more than experiments alone can garner. The power of this method lies in its ability to computationally fuse information from two disparate imaging modalities of cardiomyocyte ultrastructure to develop one unified and detailed model of the cardiomyocyte. This protocol outlines steps to integrate electron tomography and confocal microscopy images of adult male Wistar (name for a specific breed of albino rat) rat cardiomyocytes to develop a half-sarcomere finite element model of the cardiomyocyte. The procedure generates a 3D finite element model that contains an accurate, high-resolution depiction (on the order of ~35 nm) of the distribution of mitochondria, myofibrils and ryanodine receptor clusters that release the necessary calcium for cardiomyocyte contraction from the sarcoplasmic reticular network (SR) into the myofibril and cytosolic compartment. The model generated here as an illustration does not incorporate details of the transverse-tubule architecture or the sarcoplasmic reticular network and is therefore a minimal model of the cardiomyocyte. Nevertheless, the model can already be applied in simulation-based investigations into the role of cell structure in calcium signaling and mitochondrial bioenergetics, which is illustrated and discussed using two case studies that are presented following the detailed protocol.

This protocol outlines a novel method to create a spatially detailed finite element model of the intracellular architecture of cardiomyocytes from electron microscopy and confocal microscopy images. The power of this spatially detailed model is demonstrated using case studies in calcium signaling and bioenergetics.

With the advent of three-dimensional (3D) imaging technologies such as electron tomography, serial-block-face scanning electron microscopy and confocal ...

Standardized and Scalable Assay to Study Perfused 3D Angiogenic Sprouting of iPSC-derived Endothelial Cells In Vitro

Pre-clinical drug research of vascular diseases requires in vitro models of vasculature that are amendable to high-throughput screening. However, current in vitro screening models that have sufficient throughput only have limited physiological relevance, which hinders the translation of findings from in vitro to in vivo. On the other hand, microfluidic cell culture platforms have shown unparalleled physiological relevancy in vitro, but often lack the required throughput, scalability and standardization. We demonstrate a robust platform to study angiogenesis of endothelial cells derived from human induced pluripotent stem cells (iPSC-ECs) in a physiological relevant cellular microenvironment, including perfusion and gradients. The iPSC-ECs are cultured as 40 perfused 3D microvessels against a patterned collagen-1 scaffold. Upon the application of a gradient of angiogenic factors, important hallmarks of angiogenesis can be studied, including the differentiation into tip- and stalk cell and the formation of perfusable lumen. Perfusion with fluorescent tracer dyes enables the study of permeability during and after anastomosis of the angiogenic sprouts. In conclusion, this method shows the feasibility of iPSC-derived ECs in a standardized and scalable 3D angiogenic assay that combines physiological relevant culture conditions in a platform that has the required robustness and scalability to be integrated within the drug screening infrastructure.

This method describes the culture of iPSC-derived endothelial cells as 40 perfused 3D microvessels in a standardized microfluidic platform. This platform enables the study of gradient-driven angiogenic sprouting in 3D, including anastomosis and stabilization of the angiogenic sprouts in a scalable and high-throughput manner.

Pre-clinical drug research of vascular diseases requires in vitro models of vasculature that are amendable to high-throughput screening. However, current ...

Fabricating Multi-Component Lipid Nanotube Networks Using the Gliding Kinesin Motility Assay

Lipid nanotube (LNT) networks represent an in vitro model system for studying molecular transport and lipid biophysics with relevance to the ubiquitous lipid tubules found in eukaryotic cells. However, in vivo LNTs are highly non-equilibrium structures that require chemical energy and molecular motors to be assembled, maintained, and reorganized. Furthermore, the composition of in vivo LNTs is complex, comprising of multiple different lipid species. Typical methods to extrude LNTs are both time- and labor-intensive, and they require optical tweezers, microbeads, and micropipettes to forcibly pull nanotubes from giant lipid vesicles. Presented here is a protocol for the gliding motility assay (GMA), in which large scale LNT networks are rapidly generated from giant unilamellar vesicles (GUVs) using kinesin-powered microtubule motility. Using this method, LNT networks are formed from a wide array of lipid formulations that mimic the complexity of biological LNTs, making them increasingly useful for in vitro studies of lipid biophysics and membrane-associated transport. Additionally, this method is capable of reliably producing LNT networks in a short time (&#60;30 min) using commonly used laboratory equipment. LNT network characteristics such as length, width, and lipid partitioning are also tunable by altering the lipid composition of the GUVs used for fabricating the networks.

This protocol describes a process for fabricating lipid nanotube networks using gliding kinesin motility in conjunction with giant unilamellar lipid vesicles.

Lipid nanotube (LNT) networks represent an in vitro model system for studying molecular transport and lipid biophysics with relevance to the ubiquitous ...

Fabrication of 3D Cardiac Microtissue Arrays using Human iPSC-Derived Cardiomyocytes, Cardiac Fibroblasts, and Endothelial Cells

Generation of human cardiomyocytes (CMs), cardiac fibroblasts (CFs), and endothelial cells (ECs) from induced pluripotent stem cells (iPSCs) has provided a unique opportunity to study the complex interplay among different cardiovascular cell types that drives tissue development and disease. In the area of cardiac tissue models, several sophisticated three-dimensional (3D) approaches use induced pluripotent stem cell-derived cardiomyocytes (iPSC-CMs) to mimic physiological relevance and native tissue environment with a combination of extracellular matrices and crosslinkers. However, these systems are complex to fabricate without microfabrication expertise and require several weeks to self-assemble. Most importantly, many of these systems lack vascular cells and cardiac fibroblasts that make up over 60% of the nonmyocytes in the human heart. Here we describe the derivation of all three cardiac cell types from iPSCs to fabricate cardiac microtissues. This facile replica molding technique allows cardiac microtissue culture in standard multi-well cell culture plates for several weeks. The platform allows user-defined control over microtissue sizes based on initial seeding density and requires less than 3 days for self-assembly to achieve observable cardiac microtissue contractions. Furthermore, the cardiac microtissues can be easily digested while maintaining high cell viability for single-cell interrogation with the use of flow cytometry and single-cell RNA sequencing (scRNA-seq). We envision that this in vitro model of cardiac microtissues will help accelerate validation studies in drug discovery and disease modeling.

Here, we describe an easy-to-use methodology to generate 3D self-assembled cardiac microtissue arrays composed of pre-differentiated human-induced pluripotent stem cell-derived cardiomyocytes, cardiac fibroblasts, and endothelial cells. This user-friendly and low cell requiring technique to generate cardiac microtissues can be implemented for disease modeling and early stages of drug development.

Generation of human cardiomyocytes (CMs), cardiac fibroblasts (CFs), and endothelial cells (ECs) from induced pluripotent stem cells (iPSCs) has provided ...

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 ...

Fibroblast Derived Human Engineered Connective Tissue for Screening Applications

Fibroblasts are phenotypically highly dynamic cells, which quickly transdifferentiate into myofibroblasts in response to biochemical and biomechanical stimuli. The current understanding of fibrotic processes, including cardiac fibrosis, remains poor, which hampers the development of new anti-fibrotic therapies. Controllable and reliable human model systems are crucial for a better understanding of fibrosis pathology. This is a highly reproducible and scalable protocol to generate engineered connective tissues (ECT) in a 48-well casting plate to facilitate studies of fibroblasts and the pathophysiology of fibrotic tissue in a 3-dimensional (3D) environment. ECT are generated around the poles with tunable stiffness, allowing for studies under a defined biomechanical load. Under the defined loading conditions, phenotypic adaptations controlled by cell-matrix interactions can be studied. Parallel testing is feasible in the 48-well format with the opportunity for the time-course analysis of multiple parameters, such as tissue compaction and contraction against the load. From these parameters, biomechanical properties such as tissue stiffness and elasticity can be studied.

Presented here is a protocol to generate engineered connective tissues for a parallel culture of 48 tissues in a multi-well plate with double poles, suitable for mechanistic studies, disease modeling, and screening applications. The protocol is compatible with fibroblasts from different organs and species and is exemplified here with human primary cardiac fibroblasts.

Fibroblasts are phenotypically highly dynamic cells, which quickly transdifferentiate into myofibroblasts in response to biochemical and biomechanical ...