We performed whole-cell patch-clamp recordings in transiently transfected HEK 293T cells to characterize in detail the GtACR1-mediated photocurrents and their ion dependency. Upon green-light application, GtACR1 activation results in large photocurrents whose direction and amplitude depends on the Cl⁻ concentration gradient (Figures 1A–C). E_rev for GtACR1-mediated currents are −15.0 ± 0.3, −1.7 ± 0.2, and +35.6 ± 1.1 mV for extracellular Cl⁻ concentrations ([Cl⁻]_ex) of 151.4, 71.4, and 11.4 mM, respectively, (with an intracellular Cl⁻ concentration [Cl⁻]_in of 54.0 mM) (Figure 1G). The experimentally observed reversal potentials deviate from calculated reversal potentials for a pure chloride channel at 25 °C in a non-linear manner (theoretical reversal potentials according to the Nernst equation are −26.5, −7.2, and +40.0 mV, respectively). Decreasing Cl⁻ levels of the internal (pipette) solution shifts reversal potentials to more negative values (−32.1 ± 1.0 and −45.8 ± 2.8 mV at 15 and 4 mM [Cl⁻]_in, respectively; Supplementary Figure 1A), again with major deviations from theoretically predicted potentials for a pure Cl⁻ conductance (−59.4 and −93.4 mV, respectively). For all Cl⁻ concentration gradients tested, we observe inward currents (in line with a net outward movement of anions) at negative membrane potentials. To test for a potential additional cation conductance that would explain differences between theoretical and experimentally observed reversal potentials, we also varied external pH, K⁺ and Na⁺ concentrations. Photocurrent amplitudes and reversal potentials are not altered when reducing external proton concentration to pH 9.0 (Figures 1D,H) or varying external K⁺ concentration (Supplementary Figure 1B). Decreasing the extracellular Na⁺ concentration shifts the observed whole-cell current reversal potential to slightly more positive values (−9.4 ± 0.3 and −8.9 ± 0.3 mV at 12 and 1 mM [Na⁺]_ex, respectively; [Cl⁻]_in of 54.0 mM), while average current amplitudes at negative membrane potentials appear unaffected (Figures 1E,F,I). Similar changes in reversal potential by variation of the internal Cl⁻ concentration, and the absence of a sensitivity to pH were confirmed in cardiomyocytes expressing GtACR1 (Supplementary Figures 1C,D).

Using adenoviral transduction, GtACR1-eGFP was expressed in cultured primary ventricular cardiomyocytes from rabbit hearts (Figure 2A insert). We performed whole-cell patch clamp recordings to test the functionality of the Cl⁻ channel in cardiomyocytes. At a holding potential of −60 mV, green light pulses elicit large inward currents that can be graded by applied light intensity and/or pulse duration (Figures 2A–G). Half-maximal peak amplitudes are reached at 3.08 and 0.93 mW·mm⁻² for short light pulses of 1 and 10 ms, respectively. Using longer light pulses (≥100 ms) half-maximal amplitudes are observed at approximately 0.58 mW·mm⁻². Channel closure follows a bi-exponential decline after illumination terminates (see Table 1 for τ values for selected exposure times and light intensities). When activated by longer light exposure, a proportion of channels inactivates (or transitions to an open state of lower conductance) resulting in stationary currents of reduced amplitude (Figure 2D). Repetitive channel activation results in transient channel inactivation reflected by reduction of the peak current (I_p) by 4%, which recovers to initial values with τ_recovery = (2.2 ± 0.2) s (Figures 2H,I). Our data provides evidence that GtACR1 is functional in cardiomyocytes without addition of external retinal and that current characteristics can be titrated using light pulses of different intensity and duration.

Based on previous reports using GtACR1 for neuronal and cardiac silencing, we assessed the utility of using GtACR1 to inhibit electrically evoked AP in ventricular cardiomyocytes. In current-clamp recordings, AP are triggered by 10-ms current ramps at 0.25 Hz. Prolonged green light application (64 s at 4 mW·mm⁻²) results in membrane depolarization (to −21.0 ± 1.7 mV), thereby preventing further AP generation (Figure 3A). Light-induced inhibition is reversible, and AP before and after optical silencing show comparable durations at 90% repolarization (APD₉₀; Figures 3B–D). Using an illumination protocol with varying pulse duration and light intensity for repeated depolarization of cardiomyocytes, we found that the average membrane potential during repeated light exposure increases to more positive values over time (starting membrane potential: −59.6 mV, end membrane potential: −57.4 mV). This may be due to changes in intracellular ion concentrations (Figure 3E, for exemplary current traces see Supplementary Figure 2). In contrast, green light application does not affect resting membrane potential and APD₉₀ in non-transduced control cells (Supplementary Figures 1G,H). Light-induced silencing of cardiomyocytes was also assessed in intact cells by measuring sarcomere length changes during electrical field stimulation. Similar to electrophysiological recordings, sustained light inhibits cardiomyocyte activity (Figure 3F). Upon onset of sustained illumination, we observed a short and low-amplitude contraction. While resting sarcomere length is slightly reduced in GtACR1-expressing cells compared to non-transduced control cells, no differences were observed in fractional sarcomere shortening or contraction kinetics before and after light-induced silencing compared to control cells (Figures 3G,H, Supplementary Figures 1I–N).

Since GtACR1 activation leads to cardiomyocyte membrane depolarization at negative membrane potentials, we tested whether GtACR1-expressing cardiomyocytes can be optically paced. Short low-intensity light pulses (10 ms, 0.1 mW·mm⁻²) are sufficient to reliably trigger AP in cultured GtACR1-positive cells (Figures 4D,E), but not in wild-type controls. When compared to electrically induced AP (Figures 4A–G), optically stimulated AP show prolonged late repolarization, reflected by a significantly longer APD₉₀, while APD at 20% repolarization (APD₂₀) is not different between optical (OP) and electrical (EP) pacing. Also, optically triggered AP show a slower onset and a late depolarized phase, as seen in the voltage difference plot between optically and electrically stimulated AP shown in Figure 4F. Diastolic membrane potential was slightly increased (more positive) with EP compared to OP (Supplementary Figure 1E). Resting membrane potential and APD₉₀ with OP are not altered when light intensity is increased from threshold to 2 × threshold (Figure 4H, Supplementary Figure 1F). Finally, in experiments tracking sarcomere length changes in intact cells, we observe successful OP, leading to cardiomyocyte contractions similar to electrically triggered beats (Figures 4I,J).

Zebrafish served as a model system to study effects of GtACR1 activation in native myocardium (Rafferty and Quinn, 2018; Ravens, 2018). GtACR1-eGFP was expressed specifically in cardiac myocytes in the hearts of zebrafish using Tol2 transposon transgenesis by microinjection of a cmlc2:GtACR1-eGFP plasmid at the one-cell stage. Fish with successful gene insertion, indicated by cardiac eGFP expression, were raised to 3 months. Hearts were isolated and intracellular microelectrode recordings of ventricular myocyte membrane potential were performed.

GtACR1-eGFP expression in the ventricle was largely epicardial and heterogeneous at the cellular level (Supplementary Figure 3), with regions of high eGFP expression next to eGFP-free areas at the whole-heart level (Figures 5, 6). Light for activation of GtACR1 was focused on regions of high or no eGFP expression (spot size of 0.16 mm²). When sustained light (5 mW·mm⁻²) is applied to regions of high eGFP expression, there is an immediate depolarization of resting membrane potential (E_R) and a decrease in the maximum rate of membrane potential change (dE/dt_max) during the AP upstroke, as well as in AP amplitude (AP_Amp), APD₅₀ and APD₉₀ (Figure 5, n = 34 cells, N = 7 zebrafish). Additionally, in regions of high eGFP-expression in some hearts (N = 3/7), sustained light locally inhibits AP generation by depolarization of membrane potential (Figure 6). Ventricular contractions could also be inhibited in vivo by exposing the entire ventricle (spot size of 0.05 mm²) of intact eGFP-positive zebrafish larvae (21 days post fertilization, N = 3) to sustained light (1.7 mW·mm⁻²; Supplementary Movie 1).

When pulsed light (10 ms, 2 × sinus heart rate) of supra-threshold intensity (1.2 ± 0.2 mW·mm⁻²; determined in each heart by increasing light intensity until AP stimulation occurred) is applied to regions of high eGFP expression, AP are elicited, resulting in OP of the heart (Figure 5). Comparison of AP, stimulated by OP and EP at the same rate, shows no difference in AP characteristics (Supplementary Figure 4). As with sustained light, pulsed light also had an effect in vivo in intact eGFP-positive zebrafish larvae, eliciting ventricular contractions resulting from OP. In contrast to GtACR1-expressing regions, neither sustained nor pulsed light application to eGFP-negative myocardial tissue in the same animals, or to ventricular myocardium from wild-type zebrafish (N = 3), induces any changes in membrane potential.

pUC57-GtACR1 was a kind gift from Dr. Jonas Wietek, Humboldt-Universität zu Berlin. The codon-optimized (mouse, human) sequence encoding for GtACR1 (GenBank accession AKN63094.1) was subcloned into the peGFP-N1 vector (Clontech, Mountain View, CA, United States) using FastDigest NheI and AgeI (Thermo Fisher Scientific, Waltham, MA, USA).

For expression in cardiomyocytes, GtACR1-eGFP was cloned into pAdeno-CMV following the manufacturer's instructions (Adeno-X Adenoviral System, Clontech). Adenovirus was produced by the Viral Core Facility of the Charité—Universitätsmedizin Berlin, Germany.

HEK 293T cells were seeded onto coverslips at a final density of 25,000 cells/mL. After 24h, jetPEI Transfection Reagent (Polyplus transfection, Illkirch, France) was used for transfecting GtACR1-eGFP into HEK 293T cells. HEK cells were measured 24–48 h post-transfection.

All rabbit experiments were carried out according to the guidelines stated in Directive 2010/63/EU of the European Parliament on the protection of animals used for scientific purposes and approved by the local authorities in Baden-Württemberg (X-16/10R). Culturing of isolated ventricular cells from rabbit hearts was performed as previously described (Bernal Sierra et al., 2018). Rabbits (9-10 weeks old of both sex [3 male, 4 female], total N = 7) were anesthetized via intramuscular injection of esketamine hydrochloride (Ketanest®S 25 mg/mL, Pfizer Pharma PFE GmbH, Berlin, Germany; 12.5 mg/kg body weight) and xylazine hydrochloride (Rompun® 2%, Bayer Vital GmbH, Leverkusen, Germany; 0.2 mL/kg body weight). During anesthesia 1,000 units heparin (Heparin-Sodium 5,000 I.E./mL, B. Braun Melsungen AG, Melsungen, Germany) and 5 mg esketamine hydrochloride were given intravenously. Thiopental (Thiopental Inresa 0.5 g, Inresa Arzneimittel GmbH, Freiburg, Germany; 12.5 mg/mL) was injected intravenously until apnea. The heart was excised and rinsed in normal Tyrode solution (Table 2). The aorta was cannulated, the heart was transferred to a Langendorff perfusion setup and perfused with normal Tyrode solution to wash out the blood. The heart was then perfused with Ca²⁺-free cardioplegic solution (Table 2) for 3 more minutes after the heart had stopped beating, and it was then digested with enzyme solution (Table 2) until the tissue appeared soft (45–60 min). Left ventricle and septum were separated and cells were released by mechanical dissociation in blocking solution (Table 2).

After digestion cell suspensions were filtrated through a mesh (pore size [1 mm²]) and centrifuged for 2 min at 22 × g (gravitational acceleration). Fibroblasts remaining in the supernatant were removed and cardiomyocytes were resuspended in blocking solution. After cardiomyocytes settled, the supernatant was removed and cells were resuspended in plating medium (in mM: 5 creatine, 2 L-carnitine hydrochloride, 5 taurine, 1 sodium pyruvate, plus 0.25 U/L insulin, 10 μM cytosine-β-D-arabinofuranoside, 5% FCS, 0.05 mg/mL gentamycin [gibco®, Grand Island, NY, United States] in medium 199). Cardiomyocytes (17,500 cells/mL) were cultured on laminin (100 μg/mL) coated coverslips. Medium was exchanged 3–4 h after seeding cells, and cells were either measured after 48 h (control cells) or treated with adenovirus. Adenovirus (type 5) coding for GtACR1 was added immediately after the 3–4 h medium exchange (MOI75 for 48–72 h). Medium was renewed after 48–72 h of transduction immediately before cells were used for functional experiments.

Whole-cell patch-clamp measurements on single isolated cardiomyocytes or HEK 293T cells were performed at room temperature using an inverted DMI 4000B microscope (Leica Microsystems, Wetzlar, Germany), an Axopatch 200B amplifier and an Axon Digidata 1550A (Molecular Devices, San José, CA, United States). Activation light was delivered by a 525-nm LED (Luminus Devices PT-120-G, Sunnyvale, CA, United States) using a 530/20x filter and controlled via custom-built hardware (Essel Research and Development, Toronto, Canada). Light intensity in the object plane was determined with an optical power meter (PM100D, Thorlabs; Newton, NJ, United States) and the illuminated area was determined using a stage micrometer (A = 0.8 mm²). LED-input power was controlled via the custom-built LED control unit. If not specified otherwise, external (bath) and internal (pipette) standard solutions were used (Tables 3, 4).

Cardiomyocyte contractions were followed by detecting changes in sarcomere length as previously described (Peyronnet et al., 2017). Cardiomyocytes were imaged with a MyoCam-S camera (IonOptix, Dublin, Ireland) on an inverted microscope (DM IRBE, Leica Microsystems, Wetzlar, Germany). Sarcomere length was calculated in real time by a Fast Fourier Transform of the power spectrum of the striation pattern (IonWizard, IonOptix). Cells were field stimulated at 14–15 V/0.25 Hz with a Myopacer (IonOptix). For the contraction inhibition protocol, a light pulse of 525 nm was applied for 64 s. For OP, light pulses of 10 ms were applied at 0.25 Hz. Resting sarcomere length, time to peak, time to 90% relaxation, fractional sarcomere shortening, maximum velocity of contraction and relaxation were analyzed as described before (Peyronnet et al., 2017).

Transduced cardiomyocytes were washed two times with Dulbecco's Phosphate Buffered Saline (PBS). Ice-cold acetone was added, the samples were kept for 5 min at −20°C and washed again with PBS. Cells on coverslips were fixed with Mountant, Perma Fluor (Thermo Fisher Scientific) and flipped onto an object slide. The mounted samples were imaged with a confocal laser-scanning microscope (LSM 880; Carl Zeiss Microscopy, Oberkochen, Germany).

All experimental procedures in zebrafish were approved by the Dalhousie University Committee for Laboratory Animals and followed the guidelines of the Canadian Council on Animal Care. Details of experimental protocols are reported following the Minimum Information about a Cardiac Electrophysiology Experiment (MICEE) reporting standard (Quinn et al., 2011).

A cmlc2:GtACR1-eGFP plasmid was generated using standard cloning and microbiology techniques combined with the multisite Gateway system for Tol2 transposon transgenesis (Rafferty and Quinn, 2018). The 5′ entry plasmid p5E-cmlc2 and destination plasmid pDestTol2pA2 were obtained from Dr. Ian Scott (University of Toronto, Toronto, Canada). The donor plasmid pME-TA and 3′ entry plasmid p3E-polyA were obtained from Dr. Jason Berman (Dalhousie University, Halifax, Canada). To generate the middle entry plasmid, the sequence of the L13_CMV_GtACR1-eGFP plasmid was confirmed (using primers CMV_Forward: 5′-CGCAAATGGGCGGTAGGCGTG-3′ and EGFP-C-REV: 5′-GTTCAGGGGGAGGTGTG-3′) and then GtACR1-eGFP was amplified out of the plasmid using polymerase chain reaction (PCR) with primers GtACR1_for: 5′-GCCACCATGAGCAGCATTAC-3′ and EGFP-stop_rev: 5′-TTTACTTGTACAGCTCGTCCAT-3′. The GtACR1-eGFP PCR product, purified using a Gel Extraction Kit (QIAEX II, Qiagen; Hilden, Germany) and with the addition of a polyA (pA) tail was ligated into pME-TA to generate the middle entry plasmid pME-GtACR1-eGFP-pA, which was then subjected to restriction digest and sequencing (using primers M13F(-21): 5′-TGTAAAACGACGGCCAGT-3′, GtACR1, and EGFP_stop_rev). An expression plasmid was then generated by ligating p5E-cmlc2, pME-GtACR1-eGFP-pA, and p3E-polyA into pDestTol2pA2. Restriction digest and sequencing (using primers T7: 5′-TAATACGACTCACTATAGGG-3′ and EGFP-stop_rev) confirmed the sequence of isolated expression plasmid DNA. The cmlc2-GtACR1-eGFP-pA-pDestTol2pA2 DNA was purified with a PCR Purification Kit (QIAquick, Qiagen) before microinjection of 22.7 ng/μL DNA and 16 ng/μL Tol2 transposase mRNA into one-cell stage Casper zebrafish embryos. Injected fish were screened for successful gene transduction by eGFP expression using a fluorescent microscope (MZ16F, Leica; Wetzlar, Germany).

Hearts were isolated from 3-month post fertilization (mpf) zebrafish expressing GtACR1-eGFP as previously described (Stoyek et al., 2016, 2018; MacDonald et al., 2017). Fish were anesthetized in Tris-buffered (pH 7.4, BP152, Fisher Scientific; Hampton, United States) tricaine (1.5 mM MS-222) in room temperature tank water until opercular movements ceased and the animals lacked response to fin pinch with forceps. A midline incision was made through the ventral body wall and tissues of the ventral aorta, ventricle, atrium, and venous sinus were removed, pinned into a 15 mL dish lined with Sylgard (DC 170, Dow Corning; Midland, United States) containing Tyrode's solution (in mM: 142 NaCl, 4.7 KCl, 1 MgCl₂, 1.8 CaCl₂, 10 Glucose, 10 HEPES) of an osmolality of 300 ± 5 mOsm checked with an osmometer (Model 5004 μOsmette, Precision Systems; Natick, United States) and with pH titrated to 7.4 with NaOH. In addition, 10 μM (±)-blebbistatin (B592490, Toronto Research Chemicals; Toronto, Canada), a myosin inhibitor used for excitation-contraction uncoupling was added to the bath to eliminate contraction for stability of intracellular microelectrode recordings.

Intracellular microelectrode recordings were performed as previously described (Smith et al., 2016). Microelectrodes made from borosilicate glass tubing (0.5 mm inner diameter, 1.0 mm outer diameter, with internal filament; type BF/100/50/10, Sutter Instruments; Novato, United States) were pulled on a Brown/Flaming micropipette puller (Model P97, Sutter Instruments) to tip diameters resulting in a 40–60 MΩ resistance when filled with 3 M KCl. Electrodes were coupled to the headstage of an amplifier (Model 1600 Neuroprobe Amplifier, A/M Systems; Everett, United States) operated in current clamp mode with an electrode holder (ESW-M10N, Warner Instruments; Hamden, United States). Electrodes were advanced with a mechanical manipulator (MX/4, Narishige; Tokyo, Japan) into the ventricular epicardium. Before cell penetration, the tip potential of the electrode was nulled using the bridge controls of the intracellular amplifier with the electrode tip in the bath. At the end of a recording, the microelectrode was withdrawn from the cell, the null potential was checked, and the previous data adjusted if necessary. Transmembrane potential was taken as the difference between the potential measured in the bath with a silver/silver-chloride lead and the intracellular potential. Successful impalement was signaled by a sudden step of the electrode potential to a negative value. Criteria for accepting a cell were AP with a stable resting membrane potential (E_R) < −60 mV and an AP peak >0 mV that were maintained throughout the recording. Transmembrane potential was recorded at 10 kHz using a software-controlled data acquisition system (LabChart and PowerLab, ADInstruments; Sydney, Australia).

Experiments were performed in zebrafish isolated hearts and intact zebrafish larvae at room temperature using an upright microscope (BX51WI, Olympus; Shinjuku, Japan). Light for activation of GtACR1 was delivered by a white LED (CFT-90-W, Luminus Devices; Sunnyvale, USA) through a 531/22 nm filter (FF02-531/22, Semrock; Rochester, USA) focused on the ventricle with a 10X (for isolated hearts) or 20X (for zebrafish larvae) water-immersion objective (UMPLFLN 10XW or XLUMPLFLN 20XW, Olympus). The microscope field stop was set at the smallest aperture to produce a 0.16 mm² spot with an intensity of 1–5 mW/mm². Light application was either sustained for 15 s or pulsed for 10 ms at a rate 2 × sinus heart rate (HR) in regions displaying high or no eGFP expression, checked with 466/40 nm excitation (FF01-466/40, Semrock), 495 nm dichroic (FF495-Di03, Semrock), and 525/50 nm (ET525/50m, Chroma Technology; Bellows Falls, United States) filters. OP was compared to EP at the same rate applied extracellularly by a pair of silver wire electrodes on either side of the heart coupled to a constant current stimulus isolation unit (PSIU6D, Grass Technologies; West Warwick, USA) driven by 10 ms rectangular pulses from a waveform generator (S44, Grass). Measurements of HR, E_R, upstroke velocity (dE/dt_max), AP amplitude (AP_Amp), APD₅₀ and APD₉₀ were averaged over three consecutive heart beats before and during light stimulation. Light intensity was measured at the end of the experiment using a USB power meter (PM16-120, Thorlabs; Newton, United States).

GtACR1-eGFP expression in the ventricle was investigated in two subsets of zebrafish isolated hearts by confocal microscopy. The first subset included specimens from activation of GtACR1 in zebrafish isolated heart experiments. In this group immediately following the activation experiment isolated hearts were immersed in 2% methyl-cellulose on a depression slide, put under a cover slip, and GFP expression imaged. In the second subset, whole hearts were labeled with an antibody against GFP (1:100, PA1-980A, Thermo Fisher Scientific; Waltham, United States) to enhance the signal associated with the eGFP tagged to ACR1. Tissues were fixed overnight in 2% paraformaldehyde (RT-15710, Electron Microscopy Sciences; Hatfield, United States) with 1% dimethyl sulfoxide in phosphate-buffered saline (PBS, in mM): 140 NaCl, 50 Na₂HPO₄, with pH titrated to 7.2 with NaOH. Fixed tissues were rinsed in PBS, and then incubated with primary GFP-antibody diluted in a solution containing 0.1% Triton X-100 in PBS (PBS-T) for 36 h at room temperature with gentle agitation. Tissues were then rinsed and transferred to a solution of PBS-T containing anti-rabbit AlexaFluor 488 (1:200, A-21206, Life Technologies; Carlsbad, United States) for 24 h at room temperature with gentle agitation. Final rinsing was done with PBS and specimens were placed in Scale CUBIC-1 clearing solution (Susaki et al., 2014) overnight at room temperature with gentle agitation. Tissues were mounted on glass slides in CUBIC-1 for confocal microscopy.

In both subsets, processed specimens were examined as whole-mounts using an LSM 510 confocal microscope (Carl Zeiss Microscopy) with a 10 × , 0.45 NA objective (Plan-Apochromat SF25, Carl Zeiss) or a 25 × , 0.80 NA objective (LCI Plan-Neofluar, Carl Zeiss). Preparations were epi-illuminated with a 488 nm argon laser reflected by a 488 nm dichroic mirror (HFT 488; Carl Zeiss). Emitted fluorescence was collected using a 500–530 nm band-pass filter (Carl Zeiss AG). Confocal image Z-stacks ranging from 20 to 50 μm in depth, including a region of 2–5 μm above and below the region of interest (to ensure that all structures were captured, while limiting issues of light scattering) were acquired and processed using Zen2009 software (Carl Zeiss) from regions surrounding immunoreactive tissues.

Data was analyzed with pClamp 11, IonWizard, OriginPro, and with custom routines in Matlab. For unpaired datasets the non-parametric Mann-Whitney-test was applied. The Wilcoxon-test and two-tailed paired Student's t-test were used to pair-wise compare two datasets. For more than two datasets ANOVA one-way repeated (post-hoc test: Tukey or Dunnett) was used. A p < 0.05 was taken to indicate a significant difference between means.

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. The handling editor and reviewer CH declared their involvement as co-editors in the Research Topic, and confirm the absence of any other collaboration.