The authors would like to acknowledge Cairn Research Ltd. for their kind financial contribution which covered production costs of this publication. In addition, we thank Ms. Ines Mueller and Ms. Stefanie Kestel for their excellent technical support.
The authors&#8217; research is supported by the German Center for Cardiovascular Research (DZHK), the Deutsche Forschungsgemeinschaft (DFG, German Research Foundation, VO 1568/3-1, IRTG1816 RP12, SFB1002 TPA13 and under Germany&#8217;s Excellence Strategy - EXC 2067/1- 390729940) and the Else-Kr&#246;ner-Fresenius Stiftung (EKFS 2016_A20).

1. Cellular preparations
NOTE: Human iPSCs used in this protocol were derived from healthy donors and differentiated in monolayers using fully defined small molecule modulation of WNT signaling and lactate purification techniques as previously described12,13,14. iPSC-CMs were maintained every 2-3 days with a culture medium outlined below.
<ol>
	<li>Prepare a culture medium of basal medium (RPMI 1640) and 2% supplement (B27). Store at 4 &#176;C. Use at room temperature (RT).</li>
	<li>Prepare a plating medium of basal medium (RPMI 1640), 2% supplement (B27) and 1:2000 ROCK inhibitor. Store at 4 &#176;C. Use at RT.</li>
	<li>Coat sterilized 10 mm round glass #0 coverslips with 150 &#181;L of 1:60 factor free basement membrane matrix and incubate at 4 &#176;C for 4 h. 
	NOTE: Optimization of coverslip volume is necessary to ensure the entire glass is covered while maintaining adequate surface tension to prevent spillage. 150 &#181;L is recommended for 10 mm round coverslips. Batch size, coverslip type, coverslip volume and culture plate type can be suited to the experimenters needs.</li>
	<li>Begin iPSC-CM dissociation with an EDTA-based cell dissociation reagent. Ensure that the monolayer is completely detached by gently flushing with a 1,000 &#181;L pipette.</li>
	<li>Transfer the cellular suspension into a 15 mL tube and add double volume plating medium. Centrifuge for 10 min at 100 x g.</li>
	<li>Resuspend the pellet with a desired volume (resuspension volume) of plating medium. Count cells manually or electronically.</li>
	<li>Select optimal density per coverslip (15,000) which will allow for the isolated cellular analysis later.</li>
	<li>Calculate the volume of &#8216;active&#8217; cellular suspension (A) needed to plate all coverslips at this desired density. Apply the following formula and withdraw into a separate tube: 
	<img alt="Equation 1" src="/files/ftp_upload/61890/61890equ01v2.jpg" /></li>
	<li>Calculate the volume of extra plating medium (B) needed to accommodate each coverslip at a desired volume. Apply the following formula and add the resulting volume to the tube of active suspension: 
	<img alt="Equation 2" src="/files/ftp_upload/61890/61890equ02v2.jpg" /></li>
	<li>Remove the matrix from coverslips and apply the &#8216;coverslip volume&#8217; of cell suspension (A+B) to each coverslip. Regularly resuspend in the tube to ensure even cellular distribution.</li>
	<li>Incubate at 37 &#176;C for 1 h. Gently fill the well with plating medium.</li>
	<li>After 24 h, exchange media with normal culture medium and maintain every 2-3 days.</li>
</ol>
2. Experimental setup
<ol>
	<li>Equip an inverted epifluorescence microscope with a 40x magnification, high numerical aperture lens (N.A: &#62; 0.75) to conduct experiments.</li>
	<li>Couple a fast switching warm white LED to the transmitted illumination port of the microscope. Insert a simple red 660 nm filter into the transmitted light path. 
	NOTE: This light can be activated throughout photometry experiments to observe the sample without contaminating the green fluorescent signal.</li>
	<li>Mount a fast switching 470 nm LED head for photometry recording. Insert a 470/40 excitation filter at the epifluorescent port of the microscope to clean-up the light generated by the LED. 
	NOTE: For optimum signal quantification an illumination system with high speed feedback control of optical output is recommended.</li>
	<li>Insert a microscope cube containing a 495 nm long pass beam splitter in the mirror unit carousel within the microscope.</li>
	<li>Fit a detection arm containing an adjustable field diaphragm to the microscope C-mount port to allow region of interest selection.</li>
	<li>Separately couple a photomultiplier detector (PMT) and a USB camera to the microscope. This will form the basis of the emission detection system.</li>
	<li>Insert a filter cube containing a 565 nm long pass beam splitter and a 535/50 emission filter into the PMT port. This splits emission light between the two detectors. 
	NOTE: A camera attached to the transmitted port of the emission detection system can detect transmitted light under brightfield throughout all experiments.</li>
	<li>Couple the PMT to a power supply and a PMT amplifier. Connect the PMT amplifier output to an analogue&#160;input&#160;pin&#160;of&#160;a&#160;data acquisition system.</li>
	<li>Filter analogue data from the PMT at 1 kHz or higher.</li>
	<li>Digitize data at a frequency that is at least double that of the highest frequency present in analogue signal (2 kHz or higher) to fulfill Nyquist criteria and prevent aliasing.</li>
</ol>
3. Cellular loading with VF2.1Cl
NOTE: All steps involving this dye must be carried out in low light conditions.
<ol>
	<li>Prepare a Tyrode&#8217;s bath solution of (in mM): 140 NaCl, 10 HEPES, 10 Glucose, 4 KCl, 1 MgCl2, 2 CaCl2, pH = 7.35 and warm to 37 &#176;C.</li>
	<li>Prepare an aliquot of loading solution in a microcentrifuge tube by mixing 5 &#181;L of 1,000x VF2.1Cl and 50 &#181;L of 20% solubilizing poloxamer solution.</li>
	<li>Apply 5 &#181;L of the loading solution to 5 mL of warmed Tyrode&#8217;s solution (total 0.1x dye concentration) in a 20 mm Petri dish. 
	NOTE: Final dye concentration is 0.1x. This is 1/10th that suggested by the manufacturer. This conserves resources, ensures negligible cytotoxicity, and importantly, still retains clear optical signals from loaded cells with high signal to noise ratios.</li>
	<li>Add a single iPSC-CM cover slip to the dish and incubate at 37 &#176;C for 20 min.</li>
	<li>Assemble a heated live cell imaging chamber. Mount onto the microscope stage and fill with 500 &#181;L of fresh Tyrode&#8217;s solution.</li>
	<li>Wash the coverslip with fresh Tyrode&#8217;s solution at 37 &#176;C.</li>
	<li>Carefully apply the iPSC-CM coverslip to the pre-warmed bath chamber using fine point forceps. 
	NOTE: Ensure the chamber and its contents are always heated at physiological temperatures. If desired, commence with continuous perfusion of warmed Tyrode&#8217;s solution.</li>
</ol>
4. Electrical field stimulation
NOTE: External triggering of iPSC-CM is optional but useful for standardization of cellular dynamics and experimental parameters. It increases the ease of analysis and allows for the investigation of frequency-dependent effects.
<ol>
	<li>Attach a stimulation insert with two platinum electrodes spaced 5 mm apart into the recording chamber.</li>
	<li>Connect an external stimulator to the stimulation insert. Set to 5 ms bipolar field pulses at 0.5 Hz.</li>
	<li>Determine optimal stimulation voltage by increasing the stimulus from 1 V upwards. Threshold stimulus is defined as the lowest voltage at which cells begin to contract. Apply voltages roughly 25% above this threshold. Normal range is between 1 V and 30 V.</li>
	<li>Fix stimulation frequency with the external stimulator or trigger it with acquisition software.</li>
</ol>
5. Optical action potential acquisition
NOTE: This protocol uses a commercial software for acquisition and analysis.
<ol>
	<li>Visualize the myocytes in brightfield view using the transmitted light path and the USB camera.</li>
	<li>Select an isolated cell and tightly crop its optical path with the field diaphragm ensuring only light from the cell of interest is monitored.</li>
	<li>Activate the PMT amplifier and set the PMT supply to 750 V.</li>
	<li>Run the stimulation protocol (see step 4) along with the acquisition software and simultaneously activate the 470 nm excitation light. The latter can be done via a remote panel or automated at a fixed intensity (TTL signal).</li>
	<li>Adjust gain and offset of the PMT amplifier to make sure the signal does not saturate and is optimized for the detection range of the recording system.</li>
	<li>Record 10 sweeps ensuring stable action potentials are detected.</li>
	<li>Continue recording and immediately move the microscope stage to briefly acquire background signal from a region devoid of cells. Turn off the excitation light. 
	NOTE: This background value (Foffset ) will be used to account for any background fluorescence.</li>
	<li>If desired, locally perfuse reference drugs such as 1 &#181;M nifedipine to identify cellular responses to pharmacological manipulation.</li>
	<li>In a sequential fashion, repeat steps 5.2&#8211;5.7, each time selecting a new cell. Swap out coverslips if desired to ensure high experimental turnover in a single sitting. 
	NOTE: Loading and image acquisition protocols are described in Figure 2.</li>
</ol>
6. Data analysis
<ol>
	<li>Open a saved recording with the analysis software and average 10 sweeps containing stimulated action potentials from a single cell.</li>
	<li>Take a mean of the baseline signal representing Foffset and subtract this from the averaged trace.</li>
	<li>Calculate &#8710;F/F0 with the following formula (where F is measured fluorescence and F0 is diastolic fluorescence): 
	<img alt="Equation 3" src="/files/ftp_upload/61890/61890equ03v2.jpg" /></li>
	<li>Identify the trace baseline (diastolic) and area of interest (AP) and measure the desired cardiac action potential parameters. This includes but is not limited to the decay time for 50% (APD50) and 90% (APD90) repolarization.</li>
	<li>Export the data from this single cell to a spreadsheet software.</li>
	<li>Repeat steps 6.1 &#8211; 6.5 for all recordings. Assess results with appropriate unpaired tests or analysis of variance.</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61890/61890fig2v2.jpg" /> 
Figure 2: Loading and image acquisition protocols.&#160;(A) Flow chart of complete VF2.1Cl loading protocol for iPSC-CMs and native cardiomyocytes. (B) A simplified schematic of beam splitter (BS) and filter configurations used in this protocol for excitation and detection of VF2.1Cl emission in response to changes in transmembrane voltage. <a href="https://www.jove.com/files/ftp_upload/61890/61890fig2v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+electrophysiological+and+mechanical+effects+of+2,3-butane-dione+monoxime+and+cytochalasin-D+in+the+Langendorff+perfused+rabbit+heart.">The electrophysiological and mechanical effects of 2,3-butane-dione monoxime and cytochalasin-D in the Langendorff perfused rabbit heart.</a> Experimental Physiology. 89 (2), 163-172 (2004).</li><li>K&#233;pir&#243;, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=para-Nitroblebbistatin,+the+non-cytotoxic+and+photostable+Myosin+inhibitor.">para-Nitroblebbistatin, the non-cytotoxic and photostable Myosin inhibitor.</a> Angewandte Chemie International Edition. 53 (31), 8211-8215 (2014).</li></ol>

Cairn Research Ltd supported this publication by covering production costs of the video file.

Here we describe a basic protocol to easily acquire detailed AP profiles from isolated iPSC-CMs suitable for electrophysiological modelling and cardiac drug screening. We detect regular, robust APs from our sparsely seeded iPSC-CMs which suggests both indicator functionality and methodological fidelity.
Due to the wide spectrum of commercial methodologies for iPSC reprogramming and lack of standardization for cardiac differentiation protocols, iPSC based models can show immense variability in ...

Electrophysiological modeling of cardiomyocytes and the construction of efficient platforms for cardiac drug screening is essential for the development of therapeutic strategies for a variety of arrhythmic disorders. Rapid expansion of induced pluripotent stem cell (iPSC) technology has produced promising inroads into human disease modelling and pharmacological investigation using isolated patient derived cardiomyocytes (iPSC-CM). &#8220;Gold standard&#8221; techniques for electrophysiological characterization of these cells through patch-clamp (current-clamp) can quantify action potential (AP) morphology and duration, however, this method is incredibly complex and slow, and not well suited for high throughput data acquisition1. iPSC-CMs are regularly reported to have an increased diastolic membrane potential and increased leak current when compared to adult native cardiomyocytes2. It is suggested that smaller cell size and reduced membrane capacitance observed in iPSC-CMs may produce some systematic error when using the current-clamp technique, perhaps explaining these deviations3. In order to maximize the usefulness of an iPSC-CM platform, an additional method is valuable to increase throughput and ensure data accuracy when characterizing transmembrane voltage changes at a single cell level in iPSC-CMs.
Voltage sensitive dyes (VSD) have long been a proposed method to provide faster, non-invasive and equivalent analysis of cardiac AP kinetics comparative to those of traditional techniques4. A recent study has demonstrated the suitability of ratiometric voltage sensitive probe photometry to accurately quantify the cardiac AP5. Furthermore, the ability to readily scale up optical photometry approaches lends this technique to large scale cardiotoxicity screens critical in therapeutic drug development (e.g., CiPA). Development of standardized cardiotoxicity protocols in a blinded multi-site study using microelectrode array and voltage-sensing optical techniques has demonstrated the key value of this approach6.
Many potentiometric dyes are commercially available, and ongoing synthetic development of new probes show exciting potential for streamlining their effectiveness across a variety of cardiac and neural constructs. The ideal VSD will have augmented kinetics and sensitivity, while displaying decreased capacitive load, photobleaching and cytotoxicity. The recently synthesized VF2.1Cl (FluoVolt) expresses many of these beneficial properties largely due to its novel wire-based molecular structure, shared by other members of the new VoltageFluor (VF) family7. In contrast to common electrochromic VSDs in which simple probes molecularly and electrically conjugate to the plasma membrane, this dye consists of a passively inserted, membrane-spanning synthetic wire which pairs an electron-rich donor with a modified fluorescein fluorophore (FITC). Mechanistic details are provided in Figure 1. This dye demonstrates excellent sensitivity to membrane voltage fluctuations, displaying a 27% change in emission intensity per 100 mV as opposed to ~10% seen in other common probes at comparable speeds7. In addition, wire-based PeT systems do not directly interact with the cellular electric field which produces minimal electrical interference and negligible changes in cellular capacitive load.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61890/61890fig01.jpg" /> 
Figure 1: Chemical, spectral and mechanistic parameters of VF2.1Cl dye.&#160;(A) Chemical structure of VF2.1Cl. Molecular features to note include multiple alkyl groups within the phenylene vinylene molecular wire which facilitate insertion into the plasma membrane. A negatively charged sulfonic acid group conjugated to the FITC probe ensures fluorophore stabilization on the extracellular surface and aids near perpendicular insertion relative to the electrical field of the lipid bilayer. (B) A simplified schematic of perpendicular VF2.1Cl embedding into the plasma membrane of a target cell. (C) Absorption and emission spectra of VF2.1Cl dye. Spectra is identical to that of standard FITC and GFP probes. (D) Depiction of the mechanistic mode of action of VF2.1Cl. In resting conditions (hyperpolarized), negative intracellular voltages drive free electrons towards the rostral fluorophore. Electron abundance ensures photo-induced electron transfer (PeT) is favored as a pathway out of the excited state after the optical excitation, effectively quenching fluorescence. In contrast, a depolarized membrane potential influences downward electron movement favoring fluorescence upon optical excitation. The resulting fluorescent response is linearly related to membrane voltage and can be precisely utilized to gather detailed temporal information on cellular electrophysiological kinetics. (E) Representative brightfield (upper) and fluorescence at 470 nm (lower) images of leporine cardiomyocytes loaded with VF2.1Cl. (F) Z stack of a single loaded cardiomyocyte. Arrows indicate areas of clear localization of VF2.1Cl to the cellular membrane. Images were acquired with a spinning disk confocal system consisting of a X-lightv3 spinning disk confocal head with a 50 &#181;m pinhole pattern; LDI-7 illuminator; Prime95B camera and a PlanApo Lambda 100x objective. Scale bar: 20 &#181;m. <a href="https://www.jove.com/files/ftp_upload/61890/61890fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
The FITC probe conjugated to VF2.1Cl ensures that it can be used effectively under standard and GFP filter configurations, and it only requires a single channel acquisition system, both of which are common features of fluorescent imaging platforms. Analysis of dense human iPSC-CM monolayers with this dye has been recently reported8,9,10,11. Our protocol differs to these studies due to our investigation of single, isolated iPSC-CMs, unperturbed by the electrical and paracrine influences of dense syncytial monolayers, and our use of an affordable and customizable photometry system as opposed to complex confocal or wide-field imaging arrangements.
Here, we describe our protocol for the rapid acquisition and analysis of robust optical APs from isolated human iPSC-derived cardiomyocytes and native cardiomyocytes (see Supplementary File). We use VF2.1Cl coupled with a customizable state of the art platform for single cell photometry measurements.&#160;These experimental protocols have been approved by the ethics committee of the University Medical Center G&#246;ttingen (No. 10/9/15).

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			<td>HN77.4</td>
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			<td>6781.1</td>
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			<td>Invitrogen&#160;</td>
			<td>15140</td>
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			<td>Cairn Research&#160;</td>
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			<td>Chroma Technology</td>
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			<td>Hausser Scientific</td>
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			<td>Filter Cubes&#160;</td>
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			<td>Olympus&#160;</td>
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			<td>Cairn Research&#160;</td>
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			<td>Myopacer Cell Stimulator&#160;</td>
			<td>IonOptix</td>
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			<td>Optomask Shutter&#160;</td>
			<td>Cairn Research&#160;</td>
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			<td>Cairn Research&#160;</td>
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			<td>Warner Instruments&#160;</td>
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			<td>Cairn Research&#160;</td>
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			<td>Cairn Research&#160;</td>
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			<td>Cairn Research&#160;</td>
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			<td>Olympus&#160;</td>
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			<td>T565lpxr Dichroic Beamsplitter&#160;</td>
			<td>Chroma Technology</td>
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			<td>T660lpxr Dichroic Beamsplitter</td>
			<td>Chroma Technology</td>
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			<td>TC-20 Dual Channel Temperature Controller&#160;</td>
			<td>npi Electronic</td>
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			<td>UPLFLN 40X Objective</td>
			<td>Olympus&#160;</td>
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			<td>USB 3.0 Colour Camera&#160;</td>
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			<td></td>
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			<td>Software</td>
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			<td>Clampex 11.1</td>
			<td>Molecular Devices&#160;</td>
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			<td>Clampfit 11.1</td>
			<td>Molecular Devices&#160;</td>
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			<td>IC Capture 2.4&#160;</td>
			<td>Imaging Source&#160;</td>
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			<td>Prism 8</td>
			<td>Graphpad</td>
			<td></td>
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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.

<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61890/61890fig03.jpg" /> 
Figure 3: Optical action potential (AP) profiles of isolated native cardiomyocytes and human induced pluripotent stem cell derived cardiomyocytes (iPSC-CM).&#160;(A) Representative optical AP of a single murine cardiomyocyte (center) with Mean &#177; SEM of APD50 and APD90 (n = 7, right). (B) Representative optical AP of a singl...

Watch this Scientific Journal Video about Single-Cell Optical Action Potential Measurement in Human Induced Pluripotent Stem Cell-Derived Cardiomyocytes at JoVE.com

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

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 ...

single-cell-optical-action-potential-measurement-human-induced

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4.0K Views.  University Medical Center Goettingen, Goettingen, Germany. Here we describe optical acquisition and characterization of action potentials from induced pluripotent stem cell derived cardiomyocytes using a high-speed modular photometry system.

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

Optical Mapping of Action Potentials and Calcium Transients in the Mouse Heart

The mouse heart is a popular model for cardiovascular studies due to the existence of low cost technology for genetic engineering in this species. Cardiovascular physiological phenotyping of the mouse heart can be easily done using fluorescence imaging employing various probes for transmembrane potential (Vm), calcium transients (CaT), and other parameters. Excitation-contraction coupling is characterized by action potential and intracellular calcium dynamics; therefore, it is critically important to map both Vm and CaT simultaneously from the same location on the heart1-4. Simultaneous optical mapping from Langendorff perfused mouse hearts has the potential to elucidate mechanisms underlying heart failure, arrhythmias, metabolic disease, and other heart diseases. Visualization of activation, conduction velocity, action potential duration, and other parameters at a myriad of sites cannot be achieved from cellular level investigation but is well solved by optical mapping1,5,6. In this paper we present the instrumentation setup and experimental conditions for simultaneous optical mapping of Vm and CaT in mouse hearts with high spatio-temporal resolution using state-of-the-art CMOS imaging technology. Consistent optical recordings obtained with this method illustrate that simultaneous optical mapping of Langendorff perfused mouse hearts is both feasible and reliable.

This paper details the dissection procedure, instrumental setup, and experimental conditions during optical mapping of transmembrane potential (Vm) and intracellular calcium transient (CaT) in intact isolated Langendorff perfused mouse hearts.

The mouse heart is a popular model for cardiovascular studies due to the existence of low cost technology for genetic engineering in this species. ...

Development, Expansion, and In vivo Monitoring of Human NK Cells from Human Embryonic Stem Cells (hESCs) and Induced Pluripotent Stem Cells (iPSCs)

We present a method for deriving natural killer (NK) cells from undifferentiated hESCs and iPSCs using a feeder-free approach. This method gives rise to high levels of NK cells after 4 weeks culture and can undergo further 2-log expansion with artificial antigen presenting cells. hESC- and iPSC-derived NK cells developed in this system have a mature phenotype and function. The production of large numbers of genetically modifiable NK cells is applicable for both basic mechanistic as well as anti-tumor studies. Expression of firefly luciferase in hESC-derived NK cells allows a non-invasive approach to follow NK cell engraftment, distribution, and function. We also describe a dual-imaging scheme that allows separate monitoring of two different cell populations to more distinctly characterize their interactions in vivo. This method of derivation, expansion, and dual in vivo imaging provides a reliable approach for producing NK cells and their evaluation which is necessary to improve current NK cell adoptive therapies.

Development, Expansion, and In vivo Monitoring of Human NK Cells from Human Embryonic Stem Cells hESCs and Induced Pluripotent Stem Cells iPSCs

This protocol describes the development, expansion, and in vivo imaging of NK cells derived from hESCs and iPSCs.

We present a method for deriving natural killer (NK) cells from undifferentiated hESCs and iPSCs using a feeder-free approach. This method gives rise to ...

Transplantation of Induced Pluripotent Stem Cell-derived Mesoangioblast-like Myogenic Progenitors in Mouse Models of Muscle Regeneration

Patient-derived iPSCs could be an invaluable source of cells for future autologous cell therapy protocols. iPSC-derived myogenic stem/progenitor cells similar to pericyte-derived mesoangioblasts (iPSC-derived mesoangioblast-like stem/progenitor cells: IDEMs) can be established from iPSCs generated from patients affected by different forms of muscular dystrophy. Patient-specific IDEMs can be genetically corrected with different strategies (e.g. lentiviral vectors, human artificial chromosomes) and enhanced in their myogenic differentiation potential upon overexpression of the myogenesis regulator MyoD. This myogenic potential is then assessed in vitro with specific differentiation assays and analyzed by immunofluorescence. The regenerative potential of IDEMs is further evaluated in vivo, upon intramuscular and intra-arterial transplantation in two representative mouse models displaying acute and chronic muscle regeneration. The contribution of IDEMs to the host skeletal muscle is then confirmed by different functional tests in transplanted mice. In particular, the amelioration of the motor capacity of the animals is studied with treadmill tests. Cell engraftment and differentiation are then assessed by a number of histological and immunofluorescence assays on transplanted muscles. Overall, this paper describes the assays and tools currently utilized to evaluate the differentiation capacity of IDEMs, focusing on the transplantation methods and subsequent outcome measures to analyze the efficacy of cell transplantation.

Induced pluripotent stem cell (iPSC)-derived myogenic progenitors are promising candidates for cell therapy strategies to treat muscular dystrophies. This protocol describes transplantation and functional measurements required to evaluate the engraftment and differentiation of iPSC-derived mesoangioblasts (a type of muscle progenitors) in mouse models of acute and chronic muscle regeneration.

Patient-derived iPSCs could be an invaluable source of cells for future autologous cell therapy protocols. iPSC-derived myogenic stem/progenitor cells ...

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 ...

Isolation of Adult Human Dermal Fibroblasts from Abdominal Skin and Generation of Induced Pluripotent Stem Cells Using a Non-Integrating Method

Induced pluripotent stem cells (iPSCs) could be considered, to date, a promising source of pluripotent cells for the management of currently untreatable diseases, for the reconstitution and regeneration of injured tissues and for the development of new drugs. Despite all the advantages related to the use of iPSCs, such as the low risk of rejection, the lessened ethical issues, and the possibility to obtain them from both young and old patients without any difference in their reprogramming potential, problems to overcome are still numerous. In fact, cell reprogramming conducted with viral and integrating viruses can cause infections and the introduction of required genes can induce a genomic instability of the recipient cell, impairing their use in clinic. In particular, there are many concerns about the use of c-Myc gene, well-known from several studies for its mutation-inducing activity. Fibroblasts have emerged as the suitable cell population for cellular reprogramming as they are easy to isolate and culture and are harvested by a minimally invasive skin punch biopsy. The protocol described here provides a detailed step-by-step description of the whole procedure, from sample processing to obtain cell cultures, choice of reagents and supplies, cleaning and preparation, to cell reprogramming by the means of a commercial non-modified RNAs (NM-RNAs)-based reprogramming kit. The chosen reprogramming kit allows an effective reprogramming of human dermal fibroblast to iPSCs and small colonies can be seen as early as 24 h after the first transfection, even with modifications with the respect to the standard datasheet. The reprogramming procedure used in this protocol offers the advantage of a safe reprogramming, without the risk of infections caused by viral vector-based methods, reduces the cellular defense mechanisms, and allows the generation of xeno-free iPSCs, all critical features that are mandatory for further clinical applications.

Cell reprogramming requires the introduction of key genes, which regulate and maintain the pluripotent cell state. The protocol described enables the formation of induced pluripotent stem cells (iPSCs) colonies from human dermal fibroblasts without viral/integrating methods but using non-modified RNAs (NM-RNAs) combined with immune evasion factors reducing cellular defense mechanisms.

Induced pluripotent stem cells (iPSCs) could be considered, to date, a promising source of pluripotent cells for the management of currently untreatable ...

Guided Differentiation of Mature Kidney Podocytes from Human Induced Pluripotent Stem Cells Under Chemically Defined Conditions

Kidney disease affects more than 10% of the global population and costs billions of dollars in federal expenditures. The most severe forms of kidney disease and eventual end-stage renal failure are often caused by the damage to the glomerular podocytes, which are the highly specialized epithelial cells that function together with endothelial cells and the glomerular basement membrane to form the kidney&#8217;s filtration barrier. Advances in renal medicine have been hindered by the limited availability of primary tissues and the lack of robust methods for the derivation of functional human kidney cells, such as podocytes. The ability to derive podocytes from renewable sources, such as stem cells, could help advance current understanding of the mechanisms of human kidney development and disease, as well as provide new tools for therapeutic discovery. The goal of this protocol was to develop a method to derive mature, post-mitotic podocytes from human induced pluripotent stem (hiPS) cells with high efficiency and specificity, and under chemically defined conditions. The hiPS cell-derived podocytes produced by this method express lineage-specific markers (including nephrin, podocin, and Wilm&#8217;s Tumor 1) and exhibit the specialized morphological characteristics (including primary and secondary foot processes) associated with mature and functional podocytes. Intriguingly, these specialized features are notably absent in the immortalized podocyte cell line widely used in the field, which suggests that the protocol described herein produces human kidney podocytes that have a developmentally more mature phenotype than the existing podocyte cell lines typically used to study human kidney biology.

Presented here is a chemically defined protocol for the derivation of human kidney podocytes from induced pluripotent stem cells with high efficiency (&#62;90%) and independent of genetic manipulations or subpopulation selection. This protocol produces the desired cell type within 26 days and could be useful for nephrotoxicity testing and disease modeling.

Kidney disease affects more than 10% of the global population and costs billions of dollars in federal expenditures. The most severe forms of kidney ...

Isogenic Kidney Glomerulus Chip Engineered from Human Induced Pluripotent Stem Cells

Chronic kidney disease (CKD) affects 15% of the U.S. adult population, but the establishment of targeted therapies has been limited by the lack of functional models that can accurately predict human biological responses and nephrotoxicity. Advancements in kidney precision medicine could help overcome these limitations. However, previously established in vitro models of the human kidney glomerulus-the primary site for blood filtration and a key target of many diseases and drug toxicities-typically employ heterogeneous cell populations with limited functional characteristics and unmatched genetic backgrounds. These characteristics significantly limit their application for patient-specific disease modeling and therapeutic discovery. 
This paper presents a protocol that integrates human induced pluripotent stem (iPS) cell-derived glomerular epithelium (podocytes) and vascular endothelium from a single patient to engineer an isogenic and vascularized microfluidic kidney glomerulus chip. The resulting glomerulus chip is comprised of stem cell-derived endothelial and epithelial cell layers that express lineage-specific markers, produce basement membrane proteins, and form a tissue-tissue interface resembling the kidney's glomerular filtration barrier. The engineered glomerulus chip selectively filters molecules and recapitulates drug-induced kidney injury. The ability to reconstitute the structure and function of the kidney glomerulus using isogenic cell types creates the opportunity to model kidney disease with patient specificity and advance the utility of organs-on-chips for kidney precision medicine and related applications.

Presented here is a protocol to engineer a personalized organ-on-a-chip system that recapitulates the structure and function of the kidney glomerular filtration barrier by integrating genetically matched epithelial and vascular endothelial cells differentiated from human induced pluripotent stem cells. This bioengineered system can advance kidney precision medicine and related applications.

Chronic kidney disease (CKD) affects 15% of the U.S. adult population, but the establishment of targeted therapies has been limited by the lack of ...

Evaluation of Cardiac Contractility Modulation Therapy in 2D Human Stem Cell-Derived Cardiomyocytes

Human induced pluripotent stem cell-derived cardiomyocytes (hiPSC-CMs) are currently being explored for multiple in vitro applications and have been used in regulatory submissions. Here, we extend their use to cardiac medical device safety or performance assessments. We developed a novel method to evaluate cardiac medical device contractile properties in robustly contracting 2D hiPSC-CMs monolayers plated on a flexible extracellular matrix (ECM)-based hydrogel substrate. This tool enables the quantification of the effects of cardiac electrophysiology device signals on human cardiac function (e.g., contractile properties) with standard laboratory equipment. The 2D hiPSC-CM monolayers were cultured for 2-4 days on a flexible hydrogel substrate in a 48-well format.
The hiPSC-CMs were exposed to standard cardiac contractility modulation (CCM) medical device electrical signals and compared to control (i.e., pacing only) hiPSC-CMs. The baseline contractile properties of the 2D hiPSC-CMs were quantified by video-based detection analysis based on pixel displacement. The CCM-stimulated 2D hiPSC-CMs plated on the flexible hydrogel substrate displayed significantly enhanced contractile properties relative to baseline (i.e., before CCM stimulation), including an increased peak contraction amplitude and accelerated contraction and relaxation kinetics. Furthermore, the utilization of the flexible hydrogel substrate enables the multiplexing of the video-based cardiac-excitation contraction coupling readouts (i.e., electrophysiology, calcium handling, and contraction) in healthy and diseased hiPSC-CMs. The accurate detection and quantification of the effects of cardiac electrophysiological signals on human cardiac contraction is vital for cardiac medical device development, optimization, and de-risking. This method enables the robust visualization and quantification of the contractile properties of the cardiac syncytium, which should be valuable for nonclinical cardiac medical device safety or effectiveness testing. This paper describes, in detail, the methodology to generate 2D hiPSC-CM hydrogel substrate monolayers.

Here, we demonstrate a non-invasive cardiac medical device contractility evaluation method using 2D human induced pluripotent stem cell-derived cardiomyocyte (hiPSC-CM) monolayers, plated on a flexible substrate, coupled with video-based microscopy. This tool will be useful for the in vitro evaluation of the contractile properties of cardiac electrophysiology devices.

Human induced pluripotent stem cell-derived cardiomyocytes (hiPSC-CMs) are currently being explored for multiple in vitro applications and have been used ...

Generation of Human Blood Vessel Organoids from Pluripotent Stem Cells

An organoid is defined as an engineered multicellular in vitro tissue that mimics its corresponding in vivo organ such that it can be used to study defined aspects of that organ in a tissue culture dish. The breadth and application of human pluripotent stem cell (hPSC)-derived organoid research have advanced significantly to include the brain, retina, tear duct, heart, lung, intestine, pancreas, kidney, and blood vessels, among several other tissues. The development of methods for the generation of human microvessels, specifically, has opened the way for modeling human blood vessel development and disease in vitro and for the testing and analysis of new drugs or tissue tropism in virus infections, including SARS-CoV-2. Complex and lengthy protocols lacking visual guidance hamper the reproducibility of many stem cell-derived organoids. Additionally, the inherent stochasticity of organoid formation processes and self-organization necessitates the generation of optical protocols to advance the understanding of cell fate acquisition and programming. Here, a visually guided protocol is presented for the generation of 3D human blood vessel organoids (BVOs) engineered from hPSCs. Presenting a continuous basement membrane, vascular endothelial cells, and organized articulation with mural cells, BVOs exhibit the functional, morphological, and molecular features of human microvasculature. BVO formation is initiated through aggregate formation, followed by mesoderm and vascular induction. Vascular maturation and network formation are initiated and supported by embedding aggregates in a 3D collagen and solubilized basement membrane matrix. Human vessel networks form within 2-3 weeks and can be further grown in scalable culture systems. Importantly, BVOs transplanted into immunocompromised mice anastomose with the endogenous mouse circulation and specify into functional arteries, veins, and arterioles. The present visually guided protocol will advance human organoid research, particularly in relation to blood vessels in normal development, tissue vascularization, and disease.

This protocol describes the generation of self-organizing blood vessels from human pluripotent and induced pluripotent stem cells. These blood vessel networks exhibit an extensive and connected endothelial network surrounded by pericytes, smooth muscle actin, and a continuous basement membrane.

An organoid is defined as an engineered multicellular in vitro tissue that mimics its corresponding in vivo organ such that it can be used to study ...

Generation, High-Throughput Screening, and Biobanking of Human-Induced Pluripotent Stem Cell-Derived Cardiac Spheroids

Human-induced pluripotent stem cell-derived cardiomyocytes (hiPSC-CMs) are of paramount importance for human cardiac disease modeling and therapeutics. We recently published a cost-effective strategy for the massive expansion of hiPSC-CMs in two dimensions (2D). Two major limitations are cell immaturity and a lack of three-dimensional (3D) arrangement and scalability in high-throughput screening (HTS) platforms. To overcome these limitations, the expanded cardiomyocytes form an ideal cell source for the generation of 3D cardiac cell culture and tissue engineering techniques. The latter holds great potential in the cardiovascular field, providing more advanced and physiologically relevant HTS. Here, we describe an HTS-compatible workflow with easy scalability for the generation, maintenance, and optical analysis of cardiac spheroids (CSs) in a 96-well-format. These small CSs are essential to fill the gap present in current in vitro disease models and/or generation for 3D tissue engineering platforms. The CSs present a highly structured morphology, size, and cellular composition. Furthermore, hiPSC-CMs cultured as CSs display increased maturation and several functional features of the human heart, such as spontaneous calcium handling and contractile activity. By automatization of the complete workflow, from the generation of CSs to functional analysis, we increase intra- and inter-batch reproducibility as demonstrated by high-throughput (HT) imaging and calcium handling analysis. The described protocol allows modeling of cardiac diseases and assessing drug/therapeutic effects at the single-cell level within a complex 3D cell environment in a fully automated HTS workflow. In addition, the study describes a straightforward procedure for long-term preservation and biobanking of whole-spheroids, thereby providing researchers the opportunity to create next-generation functional tissue storage. HTS combined with long-term storage will substantially contribute to translational research in a wide range of areas, including drug discovery and testing, regenerative medicine, and the development of personalized therapies.

Presented here is a set of protocols for the generation and cryopreservation of cardiac spheroids (CSs) from human-induced pluripotent stem cell-derived cardiomyocytes cultured in a high-throughput, multidimensional format. This three-dimensional model functions as a robust platform for disease modeling, high-throughput screenings, and maintains its functionality after cryopreservation.

Human-induced pluripotent stem cell-derived cardiomyocytes (hiPSC-CMs) are of paramount importance for human cardiac disease modeling and therapeutics. We ...