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 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.
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: 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).
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.
2. Experimental setup
3. Cellular loading with VF2.1Cl
NOTE: All steps involving this dye must be carried out in low light conditions.
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.
5. Optical action potential acquisition
NOTE: This protocol uses a commercial software for acquisition and analysis.
6. Data analysis
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.
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.
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.
The authors have nothing to disclose.
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).
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 |