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Bioengineering

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

doi: 10.3791/61890 Published: December 22, 2020
Fitzwilliam Seibertz1,2, Martyn Reynolds3, Niels Voigt1,2,4

Abstract

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.

Introduction

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). “Gold standard” 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.

Figure 1
Figure 1: Chemical, spectral and mechanistic parameters of VF2.1Cl dye. (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 µm pinhole pattern; LDI-7 illuminator; Prime95B camera and a PlanApo Lambda 100x objective. Scale bar: 20 µm. Please click here to view a larger version of this figure.

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. These experimental protocols have been approved by the ethics committee of the University Medical Center Göttingen (No. 10/9/15).

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Protocol

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.

  1. Prepare a culture medium of basal medium (RPMI 1640) and 2% supplement (B27). Store at 4 °C. Use at room temperature (RT).
  2. Prepare a plating medium of basal medium (RPMI 1640), 2% supplement (B27) and 1:2000 ROCK inhibitor. Store at 4 °C. Use at RT.
  3. Coat sterilized 10 mm round glass #0 coverslips with 150 µL of 1:60 factor free basement membrane matrix and incubate at 4 °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 µL is recommended for 10 mm round coverslips. Batch size, coverslip type, coverslip volume and culture plate type can be suited to the experimenters needs.
  4. 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 µL pipette.
  5. Transfer the cellular suspension into a 15 mL tube and add double volume plating medium. Centrifuge for 10 min at 100 x g.
  6. Resuspend the pellet with a desired volume (resuspension volume) of plating medium. Count cells manually or electronically.
  7. Select optimal density per coverslip (15,000) which will allow for the isolated cellular analysis later.
  8. Calculate the volume of ‘active’ cellular suspension (A) needed to plate all coverslips at this desired density. Apply the following formula and withdraw into a separate tube:
    Equation 1
  9. 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:
    Equation 2
  10. Remove the matrix from coverslips and apply the ‘coverslip volume’ of cell suspension (A+B) to each coverslip. Regularly resuspend in the tube to ensure even cellular distribution.
  11. Incubate at 37 °C for 1 h. Gently fill the well with plating medium.
  12. After 24 h, exchange media with normal culture medium and maintain every 2-3 days.

2. Experimental setup

  1. Equip an inverted epifluorescence microscope with a 40x magnification, high numerical aperture lens (N.A: > 0.75) to conduct experiments.
  2. 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.
  3. 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.
  4. Insert a microscope cube containing a 495 nm long pass beam splitter in the mirror unit carousel within the microscope.
  5. Fit a detection arm containing an adjustable field diaphragm to the microscope C-mount port to allow region of interest selection.
  6. Separately couple a photomultiplier detector (PMT) and a USB camera to the microscope. This will form the basis of the emission detection system.
  7. 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.
  8. Couple the PMT to a power supply and a PMT amplifier. Connect the PMT amplifier output to an analogue input pin of a data acquisition system.
  9. Filter analogue data from the PMT at 1 kHz or higher.
  10. 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.

3. Cellular loading with VF2.1Cl

NOTE: All steps involving this dye must be carried out in low light conditions.

  1. Prepare a Tyrode’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 °C.
  2. Prepare an aliquot of loading solution in a microcentrifuge tube by mixing 5 µL of 1,000x VF2.1Cl and 50 µL of 20% solubilizing poloxamer solution.
  3. Apply 5 µL of the loading solution to 5 mL of warmed Tyrode’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.
  4. Add a single iPSC-CM cover slip to the dish and incubate at 37 °C for 20 min.
  5. Assemble a heated live cell imaging chamber. Mount onto the microscope stage and fill with 500 µL of fresh Tyrode’s solution.
  6. Wash the coverslip with fresh Tyrode’s solution at 37 °C.
  7. 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’s solution.

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.

  1. Attach a stimulation insert with two platinum electrodes spaced 5 mm apart into the recording chamber.
  2. Connect an external stimulator to the stimulation insert. Set to 5 ms bipolar field pulses at 0.5 Hz.
  3. 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.
  4. Fix stimulation frequency with the external stimulator or trigger it with acquisition software.

5. Optical action potential acquisition

NOTE: This protocol uses a commercial software for acquisition and analysis.

  1. Visualize the myocytes in brightfield view using the transmitted light path and the USB camera.
  2. Select an isolated cell and tightly crop its optical path with the field diaphragm ensuring only light from the cell of interest is monitored.
  3. Activate the PMT amplifier and set the PMT supply to 750 V.
  4. 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).
  5. 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.
  6. Record 10 sweeps ensuring stable action potentials are detected.
  7. 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.
  8. If desired, locally perfuse reference drugs such as 1 µM nifedipine to identify cellular responses to pharmacological manipulation.
  9. In a sequential fashion, repeat steps 5.2–5.7, 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.

6. Data analysis

  1. Open a saved recording with the analysis software and average 10 sweeps containing stimulated action potentials from a single cell.
  2. Take a mean of the baseline signal representing Foffset and subtract this from the averaged trace.
  3. Calculate ∆F/F0 with the following formula (where F is measured fluorescence and F0 is diastolic fluorescence):
    Equation 3
  4. 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.
  5. Export the data from this single cell to a spreadsheet software.
  6. Repeat steps 6.1 – 6.5 for all recordings. Assess results with appropriate unpaired tests or analysis of variance.

Figure 2
Figure 2: Loading and image acquisition protocols. (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. Please click here to view a larger version of this figure.

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Representative Results

Figure 3
Figure 3: Optical action potential (AP) profiles of isolated native cardiomyocytes and human induced pluripotent stem cell derived cardiomyocytes (iPSC-CM). (A) Representative optical AP of a single murine cardiomyocyte (center) with Mean ± SEM of APD50 and APD90 (n = 7, right). (B) Representative optical AP of a single human iPSC derived cardiomyocyte (center) with Mean ± SEM of APD50 and APD90 (n = 48, right). (C) Pharmacological manipulation of iPSC-CM AP (center) with 1 µM nifedipine. Mean ± SEM of APD90 alteration after nifedipine application (n=5). **p < 0.01. Please click here to view a larger version of this figure.

A high signal to noise ratio was regularly observed in our samples, despite the reduced field of view when focusing on a single cell. More noise was observed in smaller iPSC-CMs, but that did not hinder analysis subsequent to ensemble averaging. AP morphology is clearly defined, giving a thorough overview of cellular electrophysiological function and repolarization mechanics across cardiomyocyte constructs. Features of note are the previously described15 sharp upstroke velocity and pronounced phase 1 characteristics of murine cardiomyocytes (APD50: 58.34±17.98 ms, APD90: 160.9±30.15 ms, n=7; Figure 3A) which are morphologically distinct from human iPSC-CM optical signals (APD50: 170.1±11.18 ms, APD90: 317.5±15.56 ms, n=48; Figure 3B). We observed a higher rate of photobleaching and cellular toxicity after optical investigation in native cardiomyocytes compared with cultured human iPSC-CMs. Protocols for preparation and dye loading in native cardiomyocytes are included in the Supplementary Material.

Human iPSC-CM are responsive to pharmacological manipulation with nifedipine (1 µM). A known L-type Ca2+ channel (CaV1.2) antagonist, nifedipine is expected to decrease the AP duration in cells with physiological function. During continuous drug application, a 41.5% decrease in APD90 was observed (n=5), suggesting both the physiological expression of CaV1.2 channels in these cells and the functionality of VF2.1Cl-based imaging as a platform for prospective high throughput cardiac drug screening studies (Figure 3C).

Supplementary Material: Native murine cardiomyocyte preparation for voltage imaging. Please click here to download this file.

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Discussion

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 their function and morphology16. This can also hinder the effectiveness of cardiotoxicity studies. We report generally robust responses to extended experimental investigation with minimal indicator-induced cytotoxicity at low excitation light intensities. Standardization of basic dissociation and coverslip seeding is critical to ensure consistent quality of iPSC-CM preparations. Loose coverslips, which can be easily inserted and replaced inside the imaging platform, are incredibly useful for rapid data acquisition and characterization of single cardiomyocytes. It should be noted however, that we do observe a slight decline in cell viability after extended periods (i.e., more than 4 weeks) of sparse culture on glass coverslips.

The modular construction of a high-speed photometry system outlined in step 2 is of critical importance to this protocol. Many optics-based setups are commercially available and can be optimized for a wide range of signal recording requirements. These range from quantitative imaging using high resolution, large area cameras through to photometry measurements of total signal from a defined area. Using the latter, we quantify fluorescence at high speed from a single masked area using a photomultiplier (PMT). Combined with fast switching illumination components, this allows for thorough dissection of fast action potential components with extremely high temporal resolution (analog signal up to 1 kHz). High sampling rates are required to ensure reliable measurements of action potential upstroke in cardiomyocytes and can be critical in other excitable constructs (i.e. neurons) with extremely fast excitation kinetics. Further, this flexible system is beneficial because 1) it allows for thorough investigation of single cell electrophysiology with field diaphragm selection, 2) digitization of the analogue signal is not integrated or pre-processed and is therefore directly under experimenter control, 3) the modular nature of the photometry instrumentation allows simple reconfiguration to enable simultaneous optical measurement with other indicators or microelectrodes. Addition of a second photomultiplier channel, with appropriate filter sets to avoid spectral overlap, will enable true simultaneous measurement of the fluorescent intracellular calcium indicator CAL-590 alongside VF2.1Cl. It is also important to note that this multipurpose system can be used alternatively for deep assessment of intracellular calcium handling as previously described17,18,19,20.

Whilst the temporal resolution of our system brings many advantages, it should be noted that parameters requiring analysis of the spatial distribution of fluorescence signals, such as excitation patterns or conduction velocity, are not suitable for investigation by the photometry techniques that we use here and are in the realms of optical mapping. The hardware described here can be readily converted to an optical mapping configuration by exchange of the photomultiplier for a suitable specialist camera with high frame rates and large well capacity. Our described configuration can offer extremely detailed temporal information at a fraction (up to 1/10th) of the commercial price of advanced large area cameras with equivalent capabilities. Confocal line scan techniques are also more spatially detailed than our method, however limitations in z restrict fluorescence acquisition from multiple transverse planes at once. This is not ideal for imaging membrane bound reporters held by iPSC-CMs with typically heterogeneous morphologies. Photometry measurements using a standard epifluorescence microscope avoid this issue by accessing the entire cellular surface instantaneously, again at a much lower cost per data point.

We highly recommend VF2.1Cl for conducting voltage assays with excitable cells. Currently, many VSDs are available that operate under a multitude of different mechanisms, each with their own inherent limitations. Common electrochromic styryl dyes like di-8-ANEPPS or di-4-ANBDQBS that display fast responses to transmembrane voltage are however hindered by low sensitivity and high capacitive load21,22. Genetically encoded voltage indicators (GEVIs) utilize the fusion of fluorescent proteins to molecular voltage sensing domains23 or microbial opsins24 and provide highly sensitive dissections of cellular voltage dynamics, but are hindered by reduced kinetics and nonlinear responses. A list of common VSDs and their basic dynamic properties are included in Table 1. PeT probes like VF2.1Cl offer a good compromise, displaying fast kinetics and decent sensitivity with minimal cellular disruption. However, this VSD is limited because, unlike ratiometric styryl dyes21, it cannot be used to resolve absolute membrane potential. This inherent disadvantage highlights the unfortunate consequence of maintaining data accuracy at higher throughput when assaying cellular electrophysiology.

Sensitivity
(% ∆F/F per 100mV)
Speed (ms)
PeT-based dyes
VF2.1Cl7 27 <1
BeRST125 24 <1
RhoVR126 47 <1
Hemicyanine dyes
Di-8-ANEPPS21 10 <1
Di-4-ANEPPS27 8 <1
Di-4-ANBDQBS22 10–20 <1
RH23728 11 <1
PGH129 17.5 <1
GEVIs
ArcLight30 32 12324
Arch D95N31 40 4124
Mermaid32 40 17.4
VSFP 2.333 13.3 2523
QuasAr224 90 11
FRET
Dio/DPA34 56 2

Table 1: A brief list of common VSDs and their major fluorescent properties.

We note that excitation-contraction uncouplers such as blebbistatin or 2,3-butane-dione monoxime (BDM) are often used in optical voltage measurements to reduce motion artefact by suppressing cardiac contraction35. However, previous reports have shown that both compounds significantly prolong AP duration and flatten AP restitution in whole perfused hearts36,37. In addition, blebbistatin has fluorescent properties which overlap with the spectra of VF2.1Cl and therefore may not be appropriate for use with this dye38.

Our versatile protocol requires minimal adjustments for optical investigation into a variety of two-dimensional and three-dimensional excitable constructs. This method allows for rapid and precise quantification of repolarization mechanics, which can provide valuable insights into cellular ionic abnormalities. In our hands, this protocol delivers clear optical signals with good signal to noise ratios even in smaller cells. Our simple and effective platform can be applied for non-invasive validation of cardiac drug safety and high throughput screening studies using patient specific iPSC-CMs.

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Disclosures

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

Acknowledgments

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’ 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’s Excellence Strategy - EXC 2067/1- 390729940) and the Else-Kröner-Fresenius Stiftung (EKFS 2016_A20).

Materials

Name Company Catalog Number Comments
Reagents
0.25 Trypsin EDTA  Gibco  25200056
B27 Supplement  Gibco  17504044
CaCl2 Carl Roth  HN04.2
D(+)-Glucose anhydrous BioChemica ITW Reagents A1422
Fetal Bovine Serum  Gibco  10270-106
FluoVolt Membrane Potential Kit  Invitrogen  F10488
HEPES Carl Roth  HN77.4
KCl Sigma-Aldrich 6781.1
Lamanin Sigma-Aldrich 114956-81-9
Matrigel  BD 354230
NaCl  Sigma-Aldrich 9265.2
Nifedipine  Sigma-Aldrich 21829-25-4
Penicillin/Streptomycin Invitrogen  15140
ROCK Inhibitor Y27632 Stemolecule 04-0012-10
RPMI 1640 Medium  Gibco  61870010
Versene EDTA  Gibco  15040033
Equipment
495LP Dichroic Beamsplitter  Chroma Technology
Axopatch 200B Amplifier  Molecular Devices
Circle Coverslips, Thickness 0 Thermo Scientific CB00100RA020MNT0
Digidata 1550B Molecular Devices
Dual OptoLED Power Supply  Cairn Research 
ET470/40x Excitation Filter  Chroma Technology
ET535/50m Chroma Technology
Etched Neubauer Hemacytometer Hausser Scientific
Filter Cubes  Cairn Research 
IX73 Inverted Microscope  Olympus 
MonoLED Cairn Research 
Multiport Adaptors  Cairn Research 
Myopacer Cell Stimulator  IonOptix
Optomask Shutter  Cairn Research 
Optoscan System Controller Cairn Research 
PH-1 Temperature Controlled Platform   Warner Instruments 
Photomultiplier Detector  Cairn Research 
PMT Amplifier Insert Cairn Research 
PMT Supply Insert Cairn Research 
RC-26G Open Bath Chamber  Warner Instruments 
SA-OLY/2AL Stage Adaptor  Olympus 
T565lpxr Dichroic Beamsplitter  Chroma Technology
T660lpxr Dichroic Beamsplitter Chroma Technology
TC-20 Dual Channel Temperature Controller  npi Electronic
UPLFLN 40X Objective Olympus 
USB 3.0 Colour Camera  Imaging Source
Software
Clampex 11.1 Molecular Devices 
Clampfit 11.1 Molecular Devices 
IC Capture 2.4  Imaging Source 
Prism 8 Graphpad

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References

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Seibertz, F., Reynolds, M., Voigt, N. Single-Cell Optical Action Potential Measurement in Human Induced Pluripotent Stem Cell-Derived Cardiomyocytes. J. Vis. Exp. (166), e61890, doi:10.3791/61890 (2020).More

Seibertz, F., Reynolds, M., Voigt, N. Single-Cell Optical Action Potential Measurement in Human Induced Pluripotent Stem Cell-Derived Cardiomyocytes. J. Vis. Exp. (166), e61890, doi:10.3791/61890 (2020).

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