Evaluation of Optogenetic Electrophysiology Tools in Human Stem Cell-Derived Cardiomyocytes

Abstract

Current cardiac drug safety assessments focus on hERG channel block and QT prolongation for evaluating arrhythmic risks, whereas the optogenetic approach focuses on the action potential (AP) waveform generated by a monolayer of human cardiomyocytes beating synchronously, thus assessing the contribution of several ion channels on the overall drug effect. This novel tool provides arrhythmogenic sensitizing by light-induced pacing in combination with non-invasive, all-optical measurements of cardiomyocyte APs and will improve assessment of drug-induced electrophysiological aberrancies. With the help of patch clamp electrophysiology measurements, we aimed to investigate whether the optogenetic modifications alter human cardiomyocytes' electrophysiology and how well the optogenetic analyses perform against this gold standard. Patch clamp electrophysiology measurements of non-transduced stem cell-derived cardiomyocytes compared to cells expressing the commercially available optogenetic constructs Optopatch and CaViar revealed no significant changes in action potential duration (APD) parameters. Thus, inserting the optogenetic constructs into cardiomyocytes does not significantly affect the cardiomyocyte's electrophysiological properties. When comparing the two methods against each other (patch clamp vs. optogenetic imaging) we found no significant differences in APD parameters for the Optopatch transduced cells, whereas the CaViar transduced cells exhibited modest increases in APD-values measured with optogenetic imaging. Thus, to broaden the screen, we combined optogenetic measurements of membrane potential and calcium transients with contractile motion measured by video motion tracking. Furthermore, to assess how optogenetic measurements can predict changes in membrane potential, or early afterdepolarizations (EADs), cells were exposed to cumulating doses of E-4031, a hERG potassium channel blocker, and drug effects were measured at both spontaneous and paced beating rates (1, 2 Hz). Cumulating doses of E-4031 produced prolonged APDs, followed by EADs and drug-induced quiescence. These observations were corroborated by patch clamp and contractility measurements. Similar responses, although more modest were seen with the IKs potassium channel blocker JNJ-303. In conclusion, optogenetic measurements of AP waveforms combined with optical pacing compare well with the patch clamp gold standard. Combined with video motion contractile measurements, optogenetic imaging provides an appealing alternative for electrophysiological screening of human cardiomyocyte responses in pharmacological efficacy and safety testings.

Full text

ORIGINAL RESEARCH
published: 02 November 2017
doi: 10.3389/fphys.2017.00884
Frontiers in Physiology | www.frontiersin.org 1November 2017 | Volume 8 | Article 884
Edited by:
Stefano Morotti,
University of California, Davis,
United States
Reviewed by:
Daniël Antonie Pijnappels,
Leiden University, Netherlands
Joel Kralj,
University of Colorado Boulder,
United States
*Correspondence:
Eero Mervaala
eero.mervaala@helsinki.fi
†These authors have contributed
equally to this work.
Specialty section:
This article was submitted to
Integrative Physiology,
a section of the journal
Frontiers in Physiology
Received: 30 June 2017
Accepted: 18 October 2017
Published: 02 November 2017
Citation:
Björk S, Ojala EA, Nordström T,
Ahola A, Liljeström M, Hyttinen J,
Kankuri E and Mervaala E (2017)
Evaluation of Optogenetic
Electrophysiology Tools in Human
Stem Cell-Derived Cardiomyocytes.
Front. Physiol. 8:884.
doi: 10.3389/fphys.2017.00884
Evaluation of Optogenetic
Electrophysiology Tools in Human
Stem Cell-Derived Cardiomyocytes
Susann Björk 1†, Elina A. Ojala 1† , Tommy Nordström 2, Antti Ahola 3, Mikko Liljeström 4,
Jari Hyttinen 3, Esko Kankuri 1and Eero Mervaala 1
*
1Department of Pharmacology, Faculty of Medicine, University of Helsinki, Helsinki, Finland, 2Department of Physiology,
Faculty of Medicine, University of Helsinki, Helsinki, Finland, 3BioMediTech Institute and Faculty of Biomedical Sciences and
Engineering, Tampere University of Technology, Tampere, Finland, 4Department of Anatomy, Faculty of Medicine and HiLIFE,
University of Helsinki, Helsinki, Finland
Current cardiac drug safety assessments focus on hERG channel block and QT
prolongation for evaluating arrhythmic risks, whereas the optogenetic approach
focuses on the action potential (AP) waveform generated by a monolayer of human
cardiomyocytes beating synchronously, thus assessing the contribution of several ion
channels on the overall drug effect. This novel tool provides arrhythmogenic sensitizing
by light-induced pacing in combination with non-invasive, all-optical measurements of
cardiomyocyte APs and will improve assessment of drug-induced electrophysiological
aberrancies. With the help of patch clamp electrophysiology measurements, we aimed
to investigate whether the optogenetic modifications alter human cardiomyocytes’
electrophysiology and how well the optogenetic analyses perform against this gold
standard. Patch clamp electrophysiology measurements of non-transduced stem
cell-derived cardiomyocytes compared to cells expressing the commercially available
optogenetic constructs Optopatch and CaViar revealed no significant changes in action
potential duration (APD) parameters. Thus, inserting the optogenetic constructs into
cardiomyocytes does not significantly affect the cardiomyocyte’s electrophysiological
properties. When comparing the two methods against each other (patch clamp vs.
optogenetic imaging) we found no significant differences in APD parameters for the
Optopatch transduced cells, whereas the CaViar transduced cells exhibited modest
increases in APD-values measured with optogenetic imaging. Thus, to broaden the
screen, we combined optogenetic measurements of membrane potential and calcium
transients with contractile motion measured by video motion tracking. Furthermore, to
assess how optogenetic measurements can predict changes in membrane potential, or
early afterdepolarizations (EADs), cells were exposed to cumulating doses of E-4031, a
hERG potassium channel blocker, and drug effects were measured at both spontaneous
and paced beating rates (1, 2 Hz). Cumulating doses of E-4031 produced prolonged
APDs, followed by EADs and drug-induced quiescence. These observations were
corroborated by patch clamp and contractility measurements. Similar responses,
although more modest were seen with the IKs potassium channel blocker JNJ-303.
In conclusion, optogenetic measurements of AP waveforms combined with optical

Björk et al. Evaluation of Optogenetic Tools in hiPSC-CMs
pacing compare well with the patch clamp gold standard. Combined with video motion
contractile measurements, optogenetic imaging provides an appealing alternative for
electrophysiological screening of human cardiomyocyte responses in pharmacological
efficacy and safety testings.
Keywords: optogenetics, human iPSC-derived cardiomyocytes, optical action potential, contractile motion, hERG,
cardiac electrophysiology, arrhythmia, safety pharmacology
INTRODUCTION
Present cardiac safety assessments focus on the in vitro block
of the human rapid component of the delayed inward rectifier
IKr (hERG) channel, combined with in vivo QT prolongation
for evaluating the arrhythmic risks of novel drug candidates
in preclinical development. Block of the hERG channel delays
cardiac repolarization, prolonging the action potential duration
(APD) and the QT interval on ECG, and potentially increases
the risk for the development of the cardiac arrhythmia Torsades
de Pointes (TdP) (Sanguinetti et al., 1995; Redfern et al., 2003;
Gintant et al., 2016). However, although these assays have been
effective in preventing drugs that induce TdP proarrhythmia
from entering the market, it has been recognized that a drug’s
proarrhythmic effect often is shaped by its action on multiple
ion channels (Mirams et al., 2011). The lack of specificity of
the hERG assay therefore often leads to unwarranted attrition
of drugs, which is costly for the pharmaceutical industry. A
more focused approach to address and eliminate cardiovascular
toxicity early in development has thus been proposed by the
Comprehensive in vitro Proarrhythmia Assay (CIPA) initiative
(Gintant et al., 2016). CIPA proposes a multimodal approach
of cardiac safety screening based on the integrated effects of
drugs on the multiple cardiac ion channels that define cardiac
excitability and repolarization and that play a role in delayed
ventricular repolarization. Reconstructions of the drug effects are
evaluated in silico on a computationally reconstructed human
ventricular cardiomyocyte action potential (AP) (Cavero and
Holzgrefe, 2014; Fermini et al., 2016; Gintant et al., 2016;
Page et al., 2016). Finally, predicted effects are verified with
electrophysiological experiments in human induced pluripotent
stem cell-derived cardiomyocytes (hiPSC-CM).
Numerous methodology development studies have appeared
which have tried to assess the criteria set up by CIPA. Optical,
non-invasive measurements of AP parameters performed in
hiPSC-CM has been associated with great potential over the
current gold standard, patch clamping, since it focuses on the
AP waveform from multiple cells beating synchronously and
thus assesses the contribution of several ion channels on the
overall drug effect (Entcheva, 2013; Ambrosi and Entcheva, 2014;
Chang Liao et al., 2015; Dempsey et al., 2016; Klimas et al.,
2016). Optogenetics utilizes light sensitive proteins (microbial
opsins), that are genetically encoded and expressed on the
cardiomyocyte plasma membrane, where they function as optical
actuators or sensors, which enables all-optical shaping of the
AP. Hochbaum and colleagues developed several optogenetic
constructs, of which Optopatch2 utilizes a modified version of
the channelrhodopsin cation channel (CheRiff) that in response
to blue light at 488 nm depolarizes the cardiomyocyte, enabling
pacing of cardiomyocytes at elevated beating rates. By combining
this optogenetic actuator with the genetically encoded voltage
indicator QuasAr2, a modified, non-pumping version of the
protein pump Archaerhodopsin3, which in response to red light
at 640 nm generates an optical signal that is proportional to the
membrane potential, all optical electrophysiological experiments
were demonstrated in neuronal cells (Hochbaum et al., 2014),
and later in hiPSC-CMs (Dempsey et al., 2016). Another
construct CaViar, based on the genetically encoded voltage sensor
Arch(D95N) combined with the genetically encoded calcium
sensor, GCaMP5f, was developed to allow for simultaneous
AP dynamics and intracellular calcium transient determinations
(Hou et al., 2014).
Traditional electrophysiological methods, such as calcium
imaging, measures the ionic functions which regulate the
contractile movement of the cells. However, these measurements
do not directly quantify the biomechanics of the cell. Different
video-based block matching methods have been developed
to non-invasively measure the contractile movement in
cardiomyocytes and the results on cellular biomechanics have
been linked to clinical findings (Kiviaho et al., 2015; Laurila et al.,
2016). Since contractile cardiotoxicity also is a safety concern,
combining optogenetic electrophysiology experiments with
contractile measurements would therefore bring added value to
safety screens. Furthermore, the arrhythmogenic sensitizing by
light-driven pacing in combination with optical measurements
of cardiomyocyte APs is essential to detect toxic drug effects
evident only under elevated beat rates. Due to the stringent
and meticulous requirements of cardiac safety testing it is of
utmost importance that these novel tools are studied in detail
to understand how the optogenetic modifications possibly alter
human cardiomyocytes’ electrophysiology. The purpose of this
study was to characterize the optogenetic constructs (Optopatch
and CaViar) against non-transduced cardiomyocytes, and more
importantly, to compare how well-optogenetic analyses perform
against the gold standard, patch clamp electrophysiology.
MATERIALS AND METHODS
Cell Culture
Human induced pluripotent stem cell-derived cardiomyocytes
(hiPSC-CMs), Cor.4U R
, were acquired from Axiogenesis Inc.
(Germany). These spontaneously beating cells represent a
mixture of atrial, nodal and ventricular cardiomyocytes, with
60% being of the ventricular type. Cor.4U R
hiPSC-CMs were
delivered as fresh cells in T25-flasks (Nunc©, Thermo Fisher
Scientific) and kept in an incubator (5% CO2, 37◦C) and fed
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Björk et al. Evaluation of Optogenetic Tools in hiPSC-CMs
daily with Cor.4U R
complete culture medium (Axiogenesis Inc.)
supplemented with 1X Antibiotic-Antimycotic (Thermo Fisher
Scientific) to prevent bacterial and fungal infection. For further
passaging T25 flasks were pre-coated with 10 µg/ml fibronectin
(Sigma) in PBS (with Ca2+and Mg2+, Gibco R
, Thermo
Fisher Scientific) for 3 h at 37 or at 4◦C o/n and the solution
was removed shortly before plating the cells. For passaging,
cells were detached using Accumax (Millipore) according to
the manufacturer’s instructions. The cells were collected by
centrifugation (200 g, 3 min), the supernatant was removed and
the cell pellet was gently re-suspended in the culture medium.
Viable cells were counted using the trypan blue exclusion method
and the cell density calculated according to viable cells. After
plating, the cells were kept in the cell culture hood for 15 min
to ensure that the cells settled evenly.
Lentiviral Transduction of Cardiomyocytes
To express light-gated voltage sensors and actuators on the
plasma membrane of the hiPSC-CMs, the cells were transduced
with lentiviral vectors bearing the constructs of interest.
CaViar, pJMK019 (Addgene plasmid # 42168) and CMV-
Optopatch2_FCK, pMOS001 (Addgene plasmid # 62984) were
gifts from Adam Cohen, acquired through the non-profit plasmid
repository Addgene. The production of lentiviral particles,
lentiviral titer determinations and replication competent virus
(RCV) tests were purchased from the Biomedicum Virus Core
Unit in the Faculty of Medicine, University of Helsinki. For
the toxicological testing of lentiviral particles, four different
amounts of lentivirus stock were tested for both optogenetic
constructs (0.25, 0.50, 0.75, and 1.00 pg/cell for CaViar; 0.50, 0.75,
1.00, and 1.25 pg/cell for Optopatch, as determined by the p24
capsid protein concentration), where final concentrations used
are underlined. For the transduction procedure, the lentiviral
stock was diluted 1:2 in serum-free BMCC medium (Axiogenesis)
supplemented with polybrene (4 µg/ml final concentration) to
assist the penetration of the viruses through the cell membrane.
The cells were incubated for 6–7 h with the lentivirus mix and
then washed with PBS to remove excess virus. The lentiviral
transduction was confirmed by CheRiff-tagged GFP or GCaMP5f
introduced in the transduced cells, imaged by an EVOS R
FL
Imaging System. After the lentiviral transduction, the cells were
washed with 3x PBS and supplemented with fresh culture
medium daily. 24 h after the transduction procedure the cells
were screened for cytotoxicity and cytolysis using an absorbance-
based lactate dehydrogenase (LDH) release assay (Pierce, Thermo
Fisher Scientific). In order to get rid of the replicative virus
prior to patch clamp measurements and optogenetic imaging, the
transduced cells were passaged for two times during a time period
of 2–3 weeks in a BSL2 safety level laboratory. Passaging was done
as described above and cells were re-plated in fibronectin-coated
T25 flasks.
Patch Clamp Electrophysiological
Measurements
Whole-cell recordings were performed using an EPC 9/2
double patch clamp amplifier and pulse v 8.80 software
(HEKA Elektronik, Lambrecht, Germany). For current clamp
recordings, non-transduced control hiPSC-CMs and hiPSC-
CMs expressing the optogenetic constructs were plated as sub-
confluent monolayer in fibronectin-coated petri dishes (30 mm,
Nunc), which were placed on an inverted microscope (Olympus
IX71) and visualized using an AxioCam HRM digital camera
(AxioVision 4.6 software). For the recordings cells were perfused
with a bathing solution composed of 143 mM NaCl, 4 mM KCl,
1.2 mM MgCl2, 1.8 mM CaCl2, 5 mM D-glucose, and 10 mM
HEPES (pH 7.4 NaOH). The internal pipette solution contained
122 mM K+-Gluconate, 30 mM KCl, 1 mM MgCl2, 5 mM
HEPES (pH 7.2, KOH). The microelectrodes were pulled from
borosilicate glass (outer diameter 1.5 mm) on a two-stage pipette
puller (PC-10, Narishige) and heat polished with a Micro Forge
MF-90 heater (Narishige). The resistance of the pipettes used in
the experiments were 2.5–3.5 M. Membrane capacitance and
series resistance were compensated electronically. The HEKA
amplifier was set to current clamp at zero applied current, and
spontaneous APs were recorded for 20 s in each data sweep.
The cells were superfused with the bathing solution at a rate
of 1.0 ml/min. All experiments were done at 37◦C by using a
TC-344B Dual automatic temperature controller (Warner). To
minimize the volume in the petri dish, a petri dish insert was used
(Bioscience Tools). Action potentials were digitized at 10 kHz and
low-pass filtered at 3 kHz.
Preparation of Cells for Optogenetic
Measurements
For the optogenetic imaging, transduced hiPSC-CMs were plated
as confluent monolayer on Geltrex (Gibco R
, Thermo Fisher
Scientific)-coated glass-bottom dishes (10 mm Ø, P35G-1.5-10-
C, MatTek), by seeding 90,000 cells per dish. Geltrex was
pre-incubated on the glass-bottom dishes at 37◦C for 1 h and
removed shortly before plating the cells. After plating, cells
were left in the cell culture hood for 15 min to ensure an even
monolayer of the cells. Cells were cultured on glass-bottom
dishes for 1 week to ensure full integration of the beating
monolayer. Just before the optogenetic imaging the culture
medium was exchanged for imaging buffer, which was identical
to the patch clamp bathing solution. Separate dishes were utilized
for spontaneous beating, 1 and 2 Hz pacing (three dishes for each
condition).
Drug Dilutions
Dried powders of E-4031 and JNJ-303 (Tocris) were dissolved
in DMSO to make a stock concentration of 10 mM. Compounds
were solubilized by vortexing the stock solution at RT and stock
solutions were stored at −20◦C until use. The drug dilutions
were prepared fresh at the day of the experiment from stocks
in imaging buffer and kept at 37◦C in 5% CO2. For optogenetic
imaging, the addition of drugs started from a blank (fresh
imaging buffer) to check proper beating of the monolayer,
followed by vehicle and drug doses. The entire volume (2 ml) in
the dish was exchanged at each drug dose, as we noted that the
cells needed fresh buffer at regular intervals for proper beating. A
delay of ∼1 min before imaging was allowed in order for the drug
to take effect. The vehicle DMSO concentrations were 0.001%
(v/v) for E-4031 series and 0.03% (v/v) for JNJ-303 series. The
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Björk et al. Evaluation of Optogenetic Tools in hiPSC-CMs
final concentrations for E-4031 were 3, 10, 30, and 100 nM and
the final concentrations for JNJ-303 were 0.03, 0.1, 0.3, 1, 3,
and 10 µM. For E-4031 patch clamp measurements drugs were
diluted in bathing solution and administered through perfusion.
Optogenetic Measurement Setup
The optogenetic imaging platform was designed for fast
photo manipulation and analysis of live cells. The platform
included an environmental chamber (5% CO2, 37◦C, EMBL), a
fully motorized inverted wide field epifluorescence microscope
(Nikon Eclipse Ti-E equipped with a Nikon IR-based Perfect
Focus System, PFS). Pulses (10 ms) of blue laser illumination
(Argon, λ=488 nm, 17 mW/mm2) were used to pace CheRiff
at 1 or 2 Hz frequencies and a red laser λ=647 nm, Ef=550
mW/mm2excited fluorescence of QuasAr2. For CaViar imaging,
the laser lines were used in combination with a beam splitter
(Hamamatsu Gemini) to allow simultaneous use of the two
separate illumination sources (λ=647 nm, Ef=550 mW/mm2
to excite fluorescence of ArchD95N and λ=488 nm, Ef=
3 mW/mm2for GCaMP5f). Fluorescence was collected via a
60×oil immersion objective (PlanApo VC) with a numerical
aperture (NA) of 1.4. Illumination was limited to the ocular field
of view (FOV, 22mm) of the Nikon Ti-E inverted microscope by
adjusting the field stop. The illuminated area was calculated from
the FOV and the objective magnification (giving a surface area
of 0.106 mm2). Optical power was measured on the microscope
sample plane with an EXFO X-Cite XR2100 power meter. Laser
illumination at 488 nm was measured with the acousto-optic
tunable filter (AOTF) set to 100% transmission (giving a power
reading of 1.8 mW). At 647 nm the laser output was set to 100
mW and the AOTF transmission was limited to 50% to avoid
saturation (giving a power reading of 9.7 mW). The actual laser
power used for imaging at 647 nm (laser output 300 mW, AOTF
100%) was calculated assuming a linear response (resulting in a
power of 58.2 mW). The software for the platform operation was
NIS-Elements advanced research v. 4.2 with 6D image acquisition
module. Signals were recorded with an Andor iXon3 897 back-
illuminated EMCCD camera (512 ×512 px) or Andor iXon+
885 EMCCD camera (1,004 ×1,002 px) for CaViar. Imaging
was conducted at a framerate of 50 frames per second. The raw
imaging data from optogenetic imaging was recorded as image
sequences, from which the total intensity signal was exported
to MS Excel in numerical format. The raw data trace was
acquired as an average signal from the cells in the whole FOV.
To calculate averages for each condition or drug concentration,
image sequences from six FOVs were recorded.
Automated Data Processing and Curve
Analysis
For the automated processing of optogenetic raw data traces
and the analysis of key features of cardiac electrophysiology,
we developed the cPot Cardiac Action Potential Calculator
software, written in MATLAB. With cPot, all raw data traces from
optogenetic imaging were normalized by fitting the acquired
signal to an exponential function. Then, peaks with larger than a
selected threshold (10% of maximum amplitude) were detected
in the normalized signal. The detected peak time points and
their respective signal values were then used to determine the
AP parameters and other key features. The key features analyzed
and reported in this study were APD at 90, 50, and 30%
repolarization (APD90, APD50, and APD30, s, respectively),
beat to beat interval (s), frequency (Hz), maximum signal level
of the peak, i.e., amplitude (1F/F for optogenetics, mV for
patch clamp) and minimum signal level between peaks (MDP).
Respective percentage levels for APDs were determined so that
100% was the overall change in signal from Peak Height to the
following MDP. Patch clamp data was analyzed with cPot in
the same way, but without normalization since the baseline in
patch clamp measurements is steady. Optogenetic calcium traces
for contraction analysis were normalized by fitting the acquired
signal to an exponential function.
Simultaneous Contraction Analysis of
Video Microscopy
The contractile movement of the cardiomyocytes was analyzed
from Optopatch and CaViar video microscopy sets using a semi-
automatic CellVisus tool (Ahola et al., 2014). It uses particle
image velocimetry based on minimum quadratic difference
to determine velocity vector fields between consecutive video
frames. Directional motion velocity signals are calculated from
AP video data by using an estimated beating focus point as a
reference. Contraction signals are generated from these motion
velocity signals by integrating with respect to time and fitting the
signal on a spline for baseline correction. Contraction amplitude
was normalized to comply with the AP and the calcium
transient for illustration. Here, we analyzed the motion from AP
measurement in both Optopatch and CaViar microscopy from 10
image sequences each.
For signal characterization, calcium transient duration (CTD)
parameters at different amplitude levels were calculated. Calcium
transient durations (CTD) parameters CTD90, CTD50, and
CTD30 were calculated by determining percentage values so that
100% was the overall change in signal from Peak Height to the
following MDP. For contraction movement, we calculated the
contraction time and relaxation time, as well as total contraction
duration (CD) parameters CD90, CD50, CD30 defined by
the beginning of the contraction and the end of relaxation
movement. Further, we measured the time difference between
the AP, calcium transient and contraction signal peaks from the
same region of interest. The effect of E-4031 to contraction was
measured by analyzing in total 92 image sequences for vehicle
and 3, 10, 30 and 100 nM drug concentrations. In addition to
the CD parameters listed above, average motion magnitude was
measured.
Statistical Testing
The APD-values were beat rate adjusted, so that beating
intervals were corrected to 60 bpm by Fridericia’s correction
formula (based on the cube-root of beating interval). Statistical
comparisons were done either with a Student’s two-sample t-test,
or for cumulating drug responses with a one-way ANOVA with
Dunnett’s test for statistical significance. Significant p-values were
∗p<0.05, ∗∗p<0.01, ∗∗∗ p<0.001.
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Björk et al. Evaluation of Optogenetic Tools in hiPSC-CMs
RESULTS
Lentiviral Transduction of hiPSC-CMs with
Optogenetic Constructs Exhibit No
Significant Side Effect on the
Electrophysiological Properties of
Cardiomyocytes
To validate the effect of lentiviral transduction of hiPSC-
CMs with the optogenetic construct Optopatch (Hochbaum
et al., 2014), we measured AP parameters with patch clamp
electrophysiology and optogenetic imaging and compared
the results against non-transduced control cells (Figure 1).
There were no significant changes in the APD at 90%
repolarization (Figure 1A), at 50% repolarization (Figure 1B), at
30% repolarization (Figure 1C), nor in beating rates (Figure 1D)
between non-transduced cells and Optopatch-transduced cells.
Nor were there any significant changes between the two methods
of measurement: patch clamp electrophysiology and optogenetic
imaging. The same stemmed for peak amplitude (Figure 1E),
although this could only be compared using patch clamp
electrophysiology, as the measured parameter for patch clamp
(mV) is not measured by optogenetic imaging. A summary
for the numerical AP parameters measured on non-transduced
and Optopatch-transduced cells in Figures 1A–E is outlined
in Figure 1F. Pearson correlations for the measured APD
parameters were 0.99 (p<0.001) both for non-transduced
against Optopatch-transduced cells, as well as for correlations
between APD parameters measured by patch clamp and
optogenetic imaging. Thus, we propose that inserting the
Optopatch construct into cardiomyocytes does not affect the
cardiomyocyte’s electrophysiological properties. To determine
the amount of viral particles for delivery of the constructs to
the cells, initial optimizations were performed with four different
lentiviral concentrations of Optopatch, in which the lowest
concentration yielded a monolayer of cells that did not pace at
elevated frequencies and the highest concentration resulted in
some cell death, therefore the second highest concentration of
virus was used. However, screened by an absorbance-based LDH
cytotoxicity method, this toxicological measurement revealed
no significant cytotoxicity or cytolysis in any of the used
lentivirus concentrations compared to non-transduced Cor.4U R

cells (Supplementary Figure 1).
The optogenetic construct CaViar (Hou et al., 2014) holds
the potential to measure changes in intracellular calcium
in addition to AP dynamics and we therefore additionally
validated the effect of transducing hiPSC-CMs with CaViar,
and measured AP parameters (Figure 2) similarly as for the
Optopatch construct. There were no significant changes in
APD90 (Figure 2A), APD50 (Figure 2B), APD30 (Figure 2C),
beating rates (Figure 2D) nor in amplitude (Figure 2E) between
non-transduced cells and CaViar-transduced cells, measured
by patch clamp electrophysiology. Pearson correlation values
for the measured APD parameters were accordingly 0.99 (p
<0.001). This indicated that inserting the CaViar construct
into hiPSC-CMs does not affect the electrophysiology of the
cardiomyocyte. Neither was there any toxicity in any of the
used concentrations of CaViar in toxicological measurement.
However, when comparing APD parameters for the CaViar
construct acquired with optogenetic imaging against those
acquired with patch clamp electrophysiology, modest, but still
statistically significant changes for APD90 and APD30 were seen.
Thus, e.g., APD90 measured by patch clamp (300 ±16 ms)
and optogenetic imaging (350 ±4 ms) exhibited a statistical
difference, whereas APD50 did not. However, correlations values
for all measured APD-values were still 0.98 (p<0.001). A
summary for the numerical AP parameters measured on non-
transduced and CaViar-transduced cells in Figures 2A–E is
outlined in Figure 2F.
Simultaneous Measurement of Action
Potential, Calcium Transients, and
Contractile Motion: Signal
Characterization and Timings
Action potentials, calcium transients, and contractions
were measured from CaViar and Optopatch recordings.
Representative signals from a CaViar recording are shown in
Figure 3A, and from an Optopatch recording in Figure 3B.
The CaViar measurements displays the AP (red) preceding
the calcium transient (green), which is then followed by a
contraction (blue). Contraction ended rapidly after reaching a
peak (slope coefficient −0.0108). The calcium transient curve
showed very similar kinetics (slope coefficient −0.0114). The
timing of peaks (Table 1) for both constructs well adheres
to cellular physiology. There was ∼30 ms interval between
the AP and calcium peaks, and a 10 ms interval between the
calcium and contraction peaks, thus a total of 40 ms between
the AP and contraction peaks in CaViar measurements. For
Optopatch measurements, the same value was 50 ms, albeit
with a 30 ms variance indicating a close similarity to the AP-
contraction dynamics of the two constructs. The Optopatch
construct does not allow for calcium measurements and
therefore the AP-Calcium interval could not be calculated.
When measuring the directional velocities, contraction time
was measured to be 180 ms in CaViar measurements and
180 ms in Optopatch measurements. Relaxation times were 270
and 230 ms, respectively. The difference was not statistically
significant in a two-sample t-test.
The measured signals were further characterized by peak
width parameters at 90, 50, and 30 signal amplitude levels.
The results are shown in Table 2. None of the differences were
statistically significant indicating very similar characteristics of
the two constructs, CaViar and Optopatch. Linear correlations
were calculated for the characterization parameters. The results
were −0.14 for CD90/CTD90, 0.18 for CD50/CTD50, and 0.48
for CD30/CTD30.
Optogenetic and Patch Clamp
Measurements Show Dose-Dependent
APD Prolongation and Early
Afterdepolarizations upon Exposure to the
hERG Potassium Channel Blocker E-4031
Many compounds have failed early on in drug development
due to block of the hERG potassium channel, and we
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Björk et al. Evaluation of Optogenetic Tools in hiPSC-CMs
FIGURE 1 | Validation of action potential parameters of virally non-transduced (control) and Optopatch-transduced hiPSC-derived cardiomyocytes with patch clamp
electrophysiology and optogenetic imaging. Action potential parameters of Optopatch-transduced cells were measured with patch clamp electrophysiology and
optogenetic (opto) imaging and the results compared against non-transduced control cells measured with patch clamp. (A) Action potential duration (APD) at 90%
repolarization, (B) at 50% repolarization, and (C) at 30% repolarization, as wells as (D) beating intervals and (E) peak amplitude. (F) Summary of parameters
measured in (A–E), averages and S.E.M, control; n=16, Optopatch measured with patch clamp; n=11, Optopatch measured with optogenetic imaging; n=26.
therefore assessed whether optogenetic measurements can show
proarrhythmia events or early afterdepolarizations (EADs),
which are known effects of hERG channel block. We utilized
the potent and selective hERG blocker E-4031, as it commonly
has been used as a positive control. Cumulating doses of E-4031
(3–100 nM) were applied to Optopatch-transduced hiPSC-CMs,
and drug effects were measured by optogenetic imaging and by
patch clamp. In optogenetic measurements, both spontaneous
and optically paced beating rates (1 and 2 Hz) were screened,
whereas patch clamp measurements only enabled recordings
at spontaneous beating rates. Cumulating doses of E-4031
produced prolonged APDs dose-dependently in both methods
(Figure 4).
In optogenetic measurements, averaged APDs from three
dishes showed that APD90 at 30 nM E-4031 concentration
was prolonged to 176% over vehicle APD90 (spontaneous
beating), 167% under 1 Hz pacing and 143% at 2 Hz, with
EADs evident at 30 nM under spontaneous and 1 Hz beat rates.
APD90 was further increased to 219% over vehicle at 100 nM
for 1 Hz and to 158% at 2 Hz. At 100 nM E-4031, APD90
decreased to 136% over vehicle under spontaneous beating which
represented an average of the different behaviors seen; either very
prolonged APDs with EADs or a decrease in peak amplitude
with an increase in frequency, finally followed by drug-induced
quiescence until beating stopped (Figures 4A,E). Similarly, in
patch clamp measurements, the prolongation of APD90 (140%)
was accompanied by EADs at 30 nM E-4031. APD90 further
increased to 247% at 100 nM. In optogenetic measurements,
a significant (p<0.001) dose-dependent decrease in peak
amplitude was evident at both spontaneous (22% over vehicle),
1 Hz (37%) and 2 Hz (46%) at 100 nM of E-4031 (Figure 4D).
Representative traces for each drug concentration are shown in
Figure 4E.
Contractile measurements revealed a dose-dependent (E-
4031), non-significant prolongation of CD90 up to 10 nM
(Table 3). EADs were detected as small twitches from 30 nM
onwards after contraction and initial relaxation had occurred.
E-4031 dose-dependently increased total relaxation time, being
significant at 30 nM (p<0.05). Motion magnitude decreased
dose-dependently, reaching significance at 30 nM (p<0.05). At
30–100 nM level, some image sequences could not be analyzed
due to the motion reaching levels undetectable by the method.
The IKs Blocker JNJ-303 Prolongs APD
Slightly and Decreases Peak Amplitude
Dose-Dependently, under Both
Spontaneous and Elevated Beating Rates
Cumulating doses of JNJ-303 (0.03–10 µM) were applied to
hiPSC-CMs, and drug effects were measured at both spontaneous
and 1–2 Hz paced beating rates (Figure 5). Prolongation of
APD90 was seen at 100–300 nM JNJ-303 under spontaneous
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Björk et al. Evaluation of Optogenetic Tools in hiPSC-CMs
FIGURE 2 | Validation of action potential parameters of virally non-transduced (control) and CaViar-transduced hiPSC-derived cardiomyocytes with patch clamp
electrophysiology and optogenetic imaging. Action potential parameters of CaViar-transduced cells were measured with patch clamp electrophysiology and
optogenetic (opto) imaging and the results compared against non-transduced control cells measured with patch clamp. (A) Action potential duration (APD) at 90%
repolarization, (B) at 50% repolarization, and (C) at 30% repolarization, as well as (D) beating intervals and (E) peak amplitude. (F) Summary of parameters measured
in (A–E), averages and S.E.M, control; n=16, CaViar measured with patch clamp; n=9, CaViar measured with optogenetic imaging; n=24. *p<0.05, **p<0.01,
***p<0.001.
beat rates, and at 30–100 nM JNJ-303 under paced frequencies
(Figure 5A). The prolongations were however statistically non-
significant. A significant dose-dependent decrease in peak
amplitude was evident already at 1 µM (58% of vehicle),
decreasing to 45% at 10 µM under spontaneous beating. A
similar decrease in the peak amplitude was also seen under 1 Hz
pacing, whereas only 10 µM exhibited a significant decrease in
peak amplitude (49% of vehicle) under 2 Hz pacing (Figure 5B).
The cells stopped responding to pacing frequencies already at
100 nM of JNJ-303, but AP recordings were still continued under
light stimulation at indicated pacing rates.
DISCUSSION
The present study was conducted to evaluate the optogenetic
electrophysiology tools against the gold standard, patch clamp
electrophysiology. Moreover, we evaluated how well-optogenetic
tools and contractile motion measurements can predict
proarrhythmia events and delayed repolarization (EADs) in
cardiac drug safety screens.
Block of the potassium channel hERG plays a critical role
in defining ventricular repolarization, however mechanistic and
translational studies demonstrate that block of IKr alone is not
highly specific for predicting either delayed repolarization or
clinical proarrhythmia events (Gintant et al., 2016). Indeed,
several drugs such as verapamil and ranolazine are potent hERG
blockers, but are not associated with either QT prolongation
or risk of TdP (Chouabe et al., 1998; Schram et al., 2004).
It has been estimated that 60% of new molecular entities
developed as potential therapeutic targets test positive in
hERG blocking assays and are thus abandoned early on in
the development pipeline (Gintant et al., 2016). However,
these hERG-expressing immortalized cell-based assays do not
represent the highly differentiated human cardiac myocyte.
The technology of generating hiPSC-cardiomyocytes holds great
potential for preclinical cardiac efficacy and safety screens
(Grskovic et al., 2011; Matsa et al., 2014). These cells are
somewhat immature, and phenotypically more close to an
embryonic myocyte than adult, reflected in their less-negative
resting potential, reduced upstroke velocities and spontaneous
automaticity. In spite of this, hiPSC-CMs have been shown to
respond in a highly predictable manner to over 40 compounds
that have a known pharmacological effect on the human
heart (Fermini et al., 2016). In addition, hiPSC technology
enables the generation of cardiomyocytes from patients with
e.g., congenital long-QT syndrome. Drug responses and toxicity
measurements adapted on these cells could thus bring toxicity
screens to a deeper level, stratifying patient responses and
reducing late-stage clinical failures (Grskovic et al., 2011; Matsa
et al., 2014). To overcome the shortcomings in current drug
safety screens, the CIPA initiative proposes the investigation
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Björk et al. Evaluation of Optogenetic Tools in hiPSC-CMs
FIGURE 3 | Simultaneous measurements of APD, calcium transients and cell contractility. Representative signals from (A) a CaViar-transduced cardiomyocyte
recording showing the action potential (red), calcium transient (green), and contraction (blue) simultaneously, and from (B) an Optopatch-transduced cardiomyocyte
recording showing the action potential (red) and contraction (blue), measured over time (s). Amplitudes describe changes in calcium (A) and action potential (B). The
insets are close-ups of a single peak.
TABLE 1 | Simultaneous measurement of CaViar and Optopatch peak intervals
and contraction time parameters.
AP-calcium
peak
interval (ms)
AP-contraction
peak interval
(ms)
Contractile
time (ms)
Relaxation
time (ms)
CaViar 30 ±20 40 ±20 180 ±30 270 ±70
Optopatch n.a. 50 ±30 180 ±30 230 ±50
The measures illustrate mean ±standard deviation (ms), (n =10), n.a., not applicable;
AP, action potential.
of drug effects on multiple human cardiac currents, tested in
hiPSC-CMs.
Although hiPSC-CM transmembrane potential measured
with patch clamp electrophysiology provide the most detailed
characterization of electrophysiological effects on cellular
repolarization, this technique is slow, technically demanding and
very low-throughput (Gintant et al., 2016). Extracellular field
potential recordings obtained through multielectrode arrays
provide a means of adapting the assay to a high-throughput
format, and communicates the rate of electrical activity and
the timing of repolarization, but lack information regarding
the morphological changes in the configuration of the AP
and the actual end of repolarization. Voltage sensing optical
platforms however offers significant advantage over these
platforms (Chang Liao et al., 2015; Dempsey et al., 2016;
Hortigon-Vinagre et al., 2016; Klimas et al., 2016), as they
provide information on the whole AP waveform, which
represent a readout of the integrated activity of multiple cardiac
ion channels. Additionally, since the cardiotoxic effect of
some drugs is evident only at elevated beat rates, insertion of
genetically encoded actuators enables light-induced pacing of
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Björk et al. Evaluation of Optogenetic Tools in hiPSC-CMs
cells to higher beat frequencies. Combination of optogenetic
pacing and voltage sensitive dyes has been reported (Park
et al., 2014; Klimas et al., 2016), however the suitability of
fluorescence dyes for long term incubations is questioned since
fluorescent dyes can cause photodynamic damage to isolated
cardiomyocytes under prolonged incubation (Schaffer et al.,
1994) or potentially alter the electrophysiological properties of
the cells (Novakova et al., 2008). HiPSC-CMs can be maintained
in culture for a long time, and thus these cells per se (in contrast to
primary animal myocytes) permit long-term experiments. Non-
invasive optogenetic measurements on hiPSC-CMs transduced
with genetically encoded optical actuators and sensors therefore
enable in vitro evaluation of chronic drug effects and delayed
cardiotoxicity, as reported in (Dempsey et al., 2016). Optogenetic
electrophysiology assessment tools thus have the potential to
improve sensitivity and specificity in the early detection of
genuine cardiotoxicity risks, thereby reducing the likelihood of
mistakenly discarding viable drug candidates and speeding the
progression of worthy drugs into clinical trials (Dempsey et al.,
2016; Gintant et al., 2016; Klimas et al., 2016).
We have evaluated hiPSC-CMs transduced with the
commercially available constructs Optopatch and CaViar
(Hochbaum et al., 2014; Hou et al., 2014) against non-transduced
cells with patch clamp electrophysiology and shown that
none of the measured AP parameters were affected. There
were no significant differences in APD at 90, 50, or 30%
repolarization, nor in peak amplitude or beat rates. The APD90
for Optopatch was 300 ±12 ms measured with patch clamp
and 320 ±8 ms measured with optogenetic imaging, both of
which compare well to the computational human ventricle
APD90-value of 300 ms (O’Hara et al., 2011), as well as to
reported APD90-values for ventricular-like hiPSC-CMs (312–
320 ms) (Zhang et al., 2009; Lahti et al., 2012). The measured
APD parameters were selected from recommendations by
the CIPA initiative, and were calculated by a MATLAB-based
software that was generated by us. When looking at how
well the optogenetic imaging experiments compared to patch
clamp measurements, we found no significant differences
between the two techniques for the Optopatch construct
(Figure 1), whereas for the CaViar construct the APD90- and
APD30-values exhibited modest, but significant differences
for APD parameters between the two techniques (Figure 2).
We utilized the CaViar construct published by Hou et al.
(2014) that contains the Arch(D95N) voltage indicator and not
the QuasAr2 voltage indicator published by Dempsey et al.
(2016), as this newer version of CaViar was not available from
Addgene at the time of our experiments. Compared to QuasAr2,
Arch(D95N) has been shown to exhibit slower responses to
voltage transients (Gong et al., 2013), which might provide a
possible explanation for the prolongation of the APD in the
Caviar expressing cells measured with optogenetic imaging.
The largest difference was seen at APD90, measured close to
the base of the AP, where small changes in the AP waveform
due to the slower kinetics of the Arch(D95N) can results in
a broader AP waveform. The newer CaViar version based on
the QuasAr2 voltage indicator might possibly alleviate this
shortcoming.
In order to rule out most methodological restrictions related
to the cellular material, we used only the well-documented,
standardized and validated commercial Axiogenesis Cor.4U R

hiPSC-CMs in our experimentations (typical confluent cell
monolayer is illustrated in Supplementary Figure 2). Future
experiments should be focused on ruling out the contribution
of atrial hiPSC-CMs, as well as on comparing the results from
Cor.4U R
cells used in this study to ventricular-enriched hiPSC-
CMs such as vCor.4U R
cells. In the future, continued evaluation
of the most optimal cell platform will provide great value in
further validating the cellular system best compatible with the
optogenetic electrophysiology tools. The experimental procedure
for lentiviral delivery of the optogenetic constructs was quite
time-consuming, as cells had to be kept at the viral core facility
until RCV negative, which took 2–3 weeks, including washing
and media exchange every day, as well as one passage of the
cells which resulted in loss of cardiomyocytes. One of the
bigger drawbacks of optogenetic imaging is the lack of ability to
determine absolute values for the resting potential. However, the
cardiac AP waveform is sensitive to resting membrane potential,
and changes in APD are expected if compounds induce a shift
in resting voltage (Dempsey et al., 2016). We can calculate
percentile differences in amplitude peak height but not compare
these to amplitudes measured by patch clamp, which yields
accurate numerical values. The red voltage fluorescence trace
photobleaches in the first few seconds of imaging, yielding a
fluorescence trace with a steep descend. To solve this problem
and to accurately measure AP parameters from curves, a
normalization step was inserted in the automated processing
of raw data by the cPot software. Dempsey et al. (2016) tested
for photobleaching or phototoxicity arising from imaging of
QuasAr2 in cardiomyocytes by measuring fluorescence during
500 s of continuous red laser illumination (500 mW/mm2) and
showed a modest signal amplitude decrease of 12% during the
acquisition with a small variability in the AP width, which was
within the natural variation in spontaneously beating hiPSC-
CMs.
Contractile and structural cardiotoxicity, seen with e.g., some
kinase-targeted cancer drugs, represent another safety concern
(Cheng and Force, 2010). Cellular electric impedance assays have
been implemented with multielectrode assays for contractility
measurements (Obergrussberger et al., 2016), but do not provide
the spatial resolution to detect movements within the cell in
detail, in contrast to the minimum quadratic difference method
based on video motion tracking used here (Ahola et al., 2014,
2017). We aimed to evaluate whether optogenetic measurements
of cardiomyocyte electrophysiology can be combined with an
assay measuring the end point of the cardiomyocyte electrical
activity, i.e., contraction and relaxation. To our knowledge this is
the first study to combine video motion tracking with optogenetic
measurement of APD and calcium transients (Figure 3,Table 1).
The measurement setup provided a detailed view on the key
components related to cardiomyocyte function, and revealed a
physiological order of AP, calcium and contraction peaks (Bers,
2002). It widens the scope of studying drug effects as changes
in any of the three signals can be quantified simultaneously.
Motion analysis can also reveal information that is beyond the
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Björk et al. Evaluation of Optogenetic Tools in hiPSC-CMs
FIGURE 4 | Optogenetic and patch clamp measurements of APD parameters under increasing doses of the hERG channel inhibitor E-4031. Quantification of the
electrophysiological parameters (A) APD90, (B) APD50, (C) APD30, and (D) AP amplitude as a function of E-4031 concentrations, at spontaneous beating (black
trace) rate and under 1 Hz (blue) and 2 Hz (red) pacing rates in the optogenetic imaging. Patch clamp measurements are represented as a green trace in (A–C). Dots
on y axis represent vehicle. Results are means ±S.E.M. from n=14–20 (opto measurements) and n=3–5 (patch clamp measurements). The table summarizes the
percent change of the drug-induced response in comparison to vehicle and indicates the statistical significance, where *p<0.05, **p<0.01, ***p<0.001.
(E) Representative traces of alterations in the optical AP waveform induced by the indicated concentrations of the hERG channel inhibitor, E-4031 at spontaneous
beating and under 1 and 2 Hz pacing rates, as well as action potential traces measured with patch clamp.
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Björk et al. Evaluation of Optogenetic Tools in hiPSC-CMs
TABLE 2 | Characterization of calcium transient and contraction signals measured using CaViar and Optopatch.
CD90 (ms) CD50 (ms) CD30 (ms) CTD90 (ms) CTD50 (ms) CTD30 (ms)
CaViar 290 ±60 170 ±30 130 ±30 470 ±60 210 ±20 140 ±20
Optopatch 250 ±40 170 ±30 140 ±30 n.a. n.a. n.a.
CTD, calcium transient duration; CD, contraction duration. The measures illustrate mean ±standard deviation (ms), (n =10), n.a., not applicable.
TABLE 3 | The effect of cumulating doses of E-4031 on CD90, relaxation time and
motion magnitude in spontaneously beating Optopatch-transduced hiPSC-CMs.
E-4031 (nM) CD90 (ms) Relaxation time (ms) Change in motion
magnitude (%)
0 (vehicle) 220 ±50 180 ±50 100
3 230 ±50 200 ±50 99
10 250 ±70 220 ±80 71
30 180 ±60* 270 ±170* 59*
100 190 ±90 280 ±170* 22*
The measures illustrate mean ±standard deviation (ms) in CD90 and relaxation time and
percentage change in motion magnitude, (n =18, 17, 20, 21 and 16, for vehicle, 3, 10,
30 and 100 nM, respectively).
*p<0.05.
electrical properties and ion fluxes in cardiomyocytes, such as
actual biomechanical timing and possible intracellular motion
defects (Ahola et al., 2014). From the contractile motion point
of view, Optopatch and CaViar appear to function similarly, as
can be seen as the relatively similar values describing contraction-
relaxation parameters between the two constructs (Tables 1,2).
None of the differences found were statistically significant (p
<0.05). The correlations between CD and CTD parameters
indicated that a connection between the two can be found, but
one cannot be directly deduced from the other. A very low
correlation value was found in CD90, indicating high variance
near baseline. This is not unexpected, as a prolongation in
calcium transient near baseline does not equate to longer cell
relaxation. The measured correlation values are lower than
previously reported by Ahola et al. (2017) where the correlations
were in the range of 0.6–0.7. However, the differences may be
explained by a different calculation method of the measurement
parameters in these two studies.
To assess how optogenetic measurement can reveal changes
in cardiomyocyte repolarization we exposed the hiPSC-CMs
to the hERG potassium channel blocker E-4031 (Figure 4).
IKr inhibition by E-4031 prolonged APD in the late phase of
repolarization consistent with the role of IKr in phase 3 of
repolarization in the adult ventricular myocyte (Gintant, 2000).
Cardiomyocytes showed cellular arrhythmias in response to
cumulating doses of E-4031. 100 nM E-4031 has been shown to
induce EADs in stem-cell derived cardiomyocytes (Peng et al.,
2010). We, similarly to Obergrussberger et al. (2016) detected
significant APD90 prolongation already at 30 nM under both
spontaneous and paced beat rates in optogenetic measurements
as well as under spontaneous beating measured by patch clamp.
In both methods, the prolongation of APD under spontaneous
beat rate was followed by EADs at 30–100 nM concentration of
E-4031. In optogenetic imaging, however, exposure to 100 nM
E-4031 more often resulted in a decrease in signal amplitude with
an increase in frequency, followed by drug-induced quiescence.
In patch clamp, the somewhat large variations in APD S.E.M.s
were due to a small sample size in the labor-intensive patch
clamp method. However, the two methods showed significant
prolongation of APD90, as well as EADs detected at the same
concentration (30 nM E-4031). Thus, we show that E-4031, which
in clinical settings has been shown to cause the proarrhythmic
event TdP, also when measured with optogenetic tools caused
an increase in the APD and induced EADs. In contraction
signals, E-4031 increased relaxation duration, decreased motion
magnitude and caused EADs at 30 nM levels. The results suggest
that contraction analysis can be a feasible tool in detecting
drug responses in high-throughput applications. However, large
sample sizes are required for definite conclusions as high
concentrations applied to individual cells or small clusters may
terminate the beating altogether, causing variances in measured
parameters.
Repolarization of the cardiac AP is not only dependent on
the hERG channel but on several ion channels including the
IKs, the second main potassium channel involved in ventricular
repolarization, and thus the length of the QT interval. To
assess whether optogenetics could detect activities of this
channel, we applied JNJ-303, a potent blocker of IKs, to hiPSC-
CMs. Previous studies using this drug revealed no peculiar
activity in standard hERG screens, but subsequently evoked
unprovoked TdP in vivo in an anesthetized dog model (Towart
et al., 2009). Our results (Figure 5) revealed a statistically non-
significant, yet visible prolongation of the APD90 already at
low concentrations of JNJ-303 under both spontaneous and
paced rates, accompanied by possible delayed afterpolarizations
(DADs). A significant decrease in signal amplitude starting
from 1 µM JNJ-303, accompanied by an increase in frequency,
which was followed by drug-induced quiescence and finally
beating arrest, thus indicating blocking activity of cardiomyocyte
repolarization. Additional experiments with a sodium channel
blocker could have shed light on how inhibited depolarization
could be measured by optogenetic imaging, though this has been
already reported by Dempsey et al. (2016).
Finally, we have shown that optogenetics reliably can detect
changes in the AP waveform, including APD prolongation,
EADs, and drug-induced quiescence. Yet, we do not propose that
optogenetic electrophysiology experiments completely replace
comprehensive patch clamp electrophysiological assessments,
but it allows for a faster prediction of successful and safe
drug candidates in a high-throughput screening (HTS) format.
Optogenetics could be utilized in the early stages of preclinical
drug development and could thus extensively reduce cost for the
pharmaceutical industry. Selected candidates taken further for
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Björk et al. Evaluation of Optogenetic Tools in hiPSC-CMs
FIGURE 5 | Optogenetic measurements of APD parameters under increasing doses of the IKs blocker JNJ-303. (A) Representative traces of alterations in the AP
waveform induced by the indicated concentrations of the IKs blocker JNJ-303 at spontaneous beating (black) and under 1 (blue) and 2 Hz (red) pacing rates. (B) AP
amplitude as a function of JNJ-303 concentrations, at the spontaneous beating rate and under 1 and 2 Hz pacing rates. Dots on y axis represent vehicle. Results are
means ±S.E.M. from n=14–25. The table summarizes the percent change in amplitude of the drug-induced response in comparison to vehicle and indicates the
statistical significance, where **p<0.01, ***p<0.001.
clinical trials could then be studied in detail on a single cell level
with patch clamp.
In conclusion, we have shown that optogenetic imaging
allows for AP waveform recordings from a cardiomyocyte
monolayer. Due to the non-invasiveness and non-toxicity of
genetically encoded voltage sensors and actuators, chronic drug
exposures are enabled. Furthermore, light-induced pacing of cells
to elevated beat rates allows for arrhythmogenic sensitization.
With the high throughput screening compatibility of hiPSC-CMs
and the optogenetic technique, broader high content screens
can be established with integrated contractility studies. Thus,
optogenetic measurements provide an appealing alternative
to electrophysiological screening of human cardiomyocyte
responses for pharmacological efficacy and safety testing.
AUTHOR CONTRIBUTIONS
SB, EO, TN, AA, ML, JH, EK, and EM: designed the research; SB,
EO, and TN: collected the data; SB, EO, TN, and AA: analyzed
the data; and SB, EO, and AA: wrote the manuscript. All authors
reviewed the manuscript.
ACKNOWLEDGMENTS
We thank Prof. Adam Cohen and Prof. Emilia Entcheva for
all their valuable advice when getting started in the field
of optogenetics and Prof. Cohen for sharing his optogenetic
constructs, and for his advice on curve processing. We thank
COB Seppo Orsila and Dr. Petteri Uusimaa (Modulight) as well
as Drs. Ari-Pekka Koivisto and Hugh Chapman (Orion Pharma)
for meaningful insights on the research project and light sources.
The fluorescence microscope instrumentation for optogenetic
imaging was set up in collaboration with the Biomedicum
Imaging Unit in the Faculty of Medicine and HiLIFE, University
of Helsinki and we are grateful for their support. The cPot
Cardiac Action Potential Calculator software was developed in
collaboration with the Systems Biology Laboratory, University
of Helsinki and we are grateful to Ville Rantanen for his
support and expertise in MATLAB. This research was supported
by the Academy of Finland, the Finnish Foundation for
Cardiovascular Research, the Sigrid Jusélius Foundation, the
Swedish Cultural Foundation in Finland, the Instrumentarium
Science Foundation and Tekes, the Finnish Funding Agency for
Innovation.
SUPPLEMENTARY MATERIAL
The Supplementary Material for this article can be found
online at: https://www.frontiersin.org/articles/10.3389/fphys.
2017.00884/full#supplementary-material
Supplementary Figure 1 | Virus-induced cytotoxicity (LDH release)
measurements. (A) Optopatch-transduced hiPSC-CMs revealed no significant
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Björk et al. Evaluation of Optogenetic Tools in hiPSC-CMs
increase in cytotoxicity or cytolysis compared to control (Cor.4U®
non-transduced cells, dotted line), measured at 24 h after lentiviral
transduction. (B) Neither CaViar-transduced hiPSCs revealed any significant
cytotoxicity over control cells. Results are means ±S.E.M. from n=3.
OD, optical density.
Supplementary Figure 2 | Confluent monolayer of hiPSC-derived
cardiomyocytes. Phase contrast microscopy image (20X) showing typical
structural characteristics and cell density of confluent monolayer of Cor.4U®
hiPSC-CMs cultured on Geltrex-coated glass-bottom dishes for optogenetic
imaging. Scale bar =100 µm.
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Conflict of Interest Statement: The authors declare that the research was
conducted in the absence of any commercial or financial relationships that could
be construed as a potential conflict of interest.
Copyright © 2017 Björk, Ojala, Nordström, Ahola, Liljeström, Hyttinen, Kankuri
and Mervaala. This is an open-access article distributed under the terms of
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