We thank Stefanie Perez-Feliz for excellent technical assistance, Dr. Jonas Wietek (Humboldt-University, Berlin, Germany) for providing the pUC57-GtACR1 plasmid, Prof. Dr. Michael Schupp (Charit&#233;- Universit&#228;tsmedizin Berlin, Institut f&#252;r Pharmakologie, Berlin) for the adenovirus production and Dr. Anastasia Khokhlova (Ural Federal University) for sharing her expertise to improve the cell isolation protocol and to re-design the pipette bending setup. The project was funded by the German Research Foundation (SPP1926: SCHN 1486/1-1; Emmy-Noether fellowship: SCHN1486/2-1) and the ERC Advanced Grant CardioNECT.

<ol>
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The authors have nothing to disclose.

Whereas optogenetic tools enable modulation of excitable cell electrophysiology in a non-invasive manner, they need thorough characterization in different cell types (e.g., CM) to allow one to choose the best available tool for a specific experimental design. The patch-clamp technique is a standard method for assessing cellular electrophysiology. In the whole-cell configuration, it allows one to record photo-activated currents across the plasma membrane or temporal changes in membrane voltage following light stimulation/inhibition. Optogenetic manipulation of electrical excitation also affects CM contractions. We use sarcomere tracking and carbon fiber-assisted force measurements to quantify the effects of optical interrogation on the mechanical activity of myocytes.
We describe a protocol to characterize the basic effects of a light-gated chloride channel, GtACR1, in CM. As model system, we chose rabbit CM, as their electrophysiological characteristics (e.g., AP shape and refractory period) resemble those observed in human CM more closely than rodent CM. Moreover, rabbit CM can be cultured for several days, long enough for adenoviral delivery and expression of GtACR1-eGFP. Notably, isolated CM change their structural properties in culture over time, including rounding of cell endings and gradual loss of cross-striation, T-tubular system and caveolae23,24. In line with this, functional alterations have been reported in cultured CM: depolarization of the resting membrane potential, prolongation of the AP and changes in cellular Ca2+ handling. For review of cellular adaptations in culture, please see Louch et al.25. Supplemental Figure 2 shows exemplary AP and contraction measurements of freshly isolated CM for comparison with those observed in cultured CM (Figure 6, Figure 7) using the here presented protocol.
Whole-cell patch-clamp recordings enable direct measurements of photocurrent properties (e.g., amplitudes and kinetics) and light-induced changes in membrane potential or AP characteristics at high temporal resolution. However, such recordings have several limitations: Firstly, the cytosol is replaced by the pipette solution in whole-cell recordings, which is advantageous to control ionic electrochemical gradients, but has the intrinsic disadvantage of washing-out cellular organelles, proteins and other compounds, thus potentially affecting cellular electrical responses. Secondly, side effects like activation of additional ion channels resulting from non-physiologically long depolarization (e.g., slow time constants of light-gated ion channels) are difficult to assess as our method only allows one to detect changes in APD, but not to conduct direct measurements of ionic concentrations in electrophysiologically relevant cell compartments. This could be done with fluorescent indicators (e.g., Ca2+ sensors) or ion-selective electrodes. Further characterization may include light intensity titrations, determination of pH-dependency, photocurrent kinetics at different membrane potentials, and recovery kinetics during repetitive light stimulation.
In contrast to patch-clamp recordings, single-cell force measurements enable analysis of cellular contractions of intact myocytes without affecting their intracellular milieu. Secondary effects on ion concentrations (e.g., Ca2+) can be indirectly assessed by determining generated force amplitude and dynamics (e.g., maximum velocity of contraction and relaxation; here not analyzed). Force measurements with the carbon fiber technique have an advantage over freely contracting cells as they provide direct information on passive and active forces in pre-loaded cells (i.e., in conditions that are more similar to the in situ or in vivo settings). Mechanical preloading is especially important when analyzing cellular contractility, as stretch affects force production and relaxation26,27.
Optogenetic approaches allow for spatiotemporally precise manipulation of the cellular membrane potential, both in single CM and intact cardiac tissue. Classically, ChR2, a light-gated cation non-selective channel, has been used for depolarization of the membrane potential, whereas light-driven proton and/or chloride pumps were used for membrane hyperpolarization. Both groups of optogenetic actuators require high expression levels, as ChR2 is characterized by an intrinsically low single-channel conductance28 and light-driven pumps maximally transport one ion per absorbed photon. Furthermore, prolonged activation of ChR2 in CM may lead to Na+ and/or Ca2+ overload, and light-driven pumps may change trans-sarcolemmal H+ or Cl- gradients29,30. In search for alternative tools for optogenetic control of CM activity, we recently tested the natural anion channelrhodopsin GtACR1, characterized by a superior single-channel conductance and higher light sensitivity compared to cation ChR such as ChR2. We found that GtACR1 activation depolarizes CM and can be used for optical pacing and inhibition, depending on the light pulse timing and duration. An additional advantage of using ACR instead of cation ChR might be the more negative reversal potential of Cl- compared to Na+, reducing artificially introduced ion currents. As we have previously shown, optical pacing with GtACR1 may lead to AP prolongation as a result of the slow component of GtACR1 channel closure, which could be overcome by using faster GtACR1 mutants19. However, AP prolongation is much less pronounced when using a lower, more physiological intracellular Cl- concentration (see Figure 6). Moreover, GtACR1-mediated inhibition by prolonged illumination results in profound membrane depolarization, which again could activate secondary Na+ and Ca2+ influx, thereby altering the activity of voltage-gated channels. In our measurements, we find that AP and contraction parameters recover to baseline within 40 s after a light-induced inhibition for 1 min (see Kopton et al. 2018, Figure 6, Figure 7). Light-gated K+ channels offer a potent alternative for silencing CM without affecting the CM resting membrane potential31.
In future we would like to quantitatively compare different optogenetic tools for their potential to inhibit cardiac activity. To this end, we test a variety of light-gated ion channels including ACR, ChR2 and red-shifted ChR variants32, as well as hyperpolarizing actuators such as halorhodopsin or the light gated adenylyl cyclase bPAC in combination with the potassium channel SthK (PAC-K)31.
The here presented protocol can be used for in-depth characterization of the electromechanical properties of CM. It is principally applicable also to CM from other species, and to CM isolated from diseased myocardium. Optical stimulation allows one to pace CM at different frequencies, and different preloads can be tested during carbon fiber contraction experiments. An interesting experiment would be to use low-intensity illumination for subthreshold depolarization, to mimic gradual increase in the resting membrane potential, as can be observed during the development of cardiac tissue remodeling during disease progression. Finally, functional measurements could be combined with Ca2+ imaging for further insight into excitation-contraction coupling, or with pharmacological interventions to evaluate the effects of different drugs on CM activity.

ChR-mediated photocurrents were first recorded in the eyespot of unicellular green algae1,2. Soon after genetic cloning and heterologous expression of Chlamydomonas reinhardtii ChR1 and ChR2, ChR were used as tools to alter the membrane potential in Xenopus oocytes and mammalian cells by light3,4. Cation non-selective ChR depolarize the membrane of cells with a resting membrane potential that is negative to the reversal potential of ChR. They can thus be used to elicit AP in excitable cells, including neurons and CM, allowing optical pacing5,6.
Complementary to cation ChR, light-driven proton, chloride and sodium pumps7,8,9 have been used to inhibit neuronal activity10,11,12. However, the latter have limitations, requiring high light intensities and sustained illumination, as one ion is transported per absorbed photon. In 2014, two independent studies by Wietek et al. and Berndt et al. described the conversion of cation-conducting ChR into ACR via mutations in the channel pore13,14. One year later, natural ACR were discovered in the cryptophyte Guillardia theta (GtACR)15. As engineered ACR showed residual cation conductance, they were replaced by natural ACR, characterized by a large single-channel conductance and high light sensitivity15. GtACR were used to silence neuronal activity by polarizing the membrane potential towards the reversal potential of chloride16,17. Govorunova et al. applied GtACR1 to cultured rat ventricular CM and showed efficient photoinhibition at low light intensity levels that were not sufficient to activate previously available inhibition tools, such as the proton pump Arch18. Our group recently reported that GtACR1-mediated photoinhibition of CM is based on depolarization and that GtACR1 can also be used, therefore, for optical pacing of CM19.
Here, we present a protocol for studying the electrophysiological and mechanical effects of GtACR1 photoactivation on cultured rabbit ventricular CM. We first describe cell isolation, culturing and transduction. Electrophysiological effects are measured using whole-cell patch-clamp recordings. Light-mediated currents at a given membrane voltage are assessed in V-clamp mode. Membrane potential dynamics are measured while electrically or optically pacing CM (I-clamp mode). Optical inhibition of electrically triggered AP is tested using sustained light application. Mechanical effects are measured using carbon fibers in combination with imaging-based tracking of sarcomere length. To do so, optically paceable cells are mechanically preloaded by attaching two carbon fibers to the plasma membrane near opposite cell ends. Sarcomere length changes are recorded during optical or electrical pacing. Finally, photoinhibition is measured during electrical field stimulation of the cells, and generated forces are analyzed.
The protocol includes the following steps shown in the flowchart in Figure 1: rabbit deep anesthesia, thiopental overdose injection, heart excision, Langendorff-perfusion and tissue digestion, mechanical dissociation of the tissue to release cells, microscopic analysis of CM yield, culturing of CM, transduction with adenovirus type 5, followed by incubation and functional experiments.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60490/60490fig1a.jpg" /> 
Figure 1: Flowchart of the protocol used to obtain electrically and optically paceable CM. Hearts are excised from rabbits 9-10 weeks old, and cardiac tissue is digested while being perfused using a Langendorff setup. Cells are released by mechanical agitation. The CM yield is counted under a microscope. CM are cultured, transduced with adenovirus type 5 and functional experiments are performed 48-72 hours post-transduction. <a href="https://www.jove.com/files/ftp_upload/60490/60490fig1alarge.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table><tbody><tr><td>&lt;strong&gt;Equipment - Cell isolation/Culturing/Transduction&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Adeno-X Adenoviral System 3 CMV</td><td>TaKaRa, Clontech Laboratories, Inc., Mountain View, California, USA</td><td></td><td></td></tr><tr><td>Aortic cannula</td><td>Radnoti</td><td></td><td>4.8 OD x 3.6 ID x 8-9 L mm</td></tr><tr><td>Coverslips &amp;oslash; 16 mm, Thickness No. 0</td><td>VWR International GmbH, Leuven, Belgium</td><td>631-0151</td><td>Borosilicate Glass</td></tr><tr><td>Griffin Silk, Black, 2 m Length, Size 3, 0.5 mm</td><td>Samuel Findings, London, UK</td><td>TSGBL3</td><td></td></tr><tr><td>Incubator</td><td>New Brunswick, Eppendorf, Sch&amp;ouml;nenbuch, Switzerland</td><td>Galaxy 170S</td><td></td></tr><tr><td>Langendorff-perfusion set-up</td><td>Zitt-Thoma Laborbedarf Glasbl&amp;auml;serei, Freiburg, Germany</td><td></td><td>Custom-made</td></tr><tr><td>Langendorff-pump</td><td>Ismatec, Labortechnik-Analytik, Glattbrugg-Z&amp;uuml;rich, Switzerland</td><td>ISM444</td><td></td></tr><tr><td>Mesh: Nylon Monodur filter cloth</td><td>Cadisch Precision Meshes Ltd</td><td></td><td>800 &amp;micro;m holes, 1 m wide</td></tr><tr><td>Neubauer chamber</td><td>VWR International GmbH, Leuven, Belgium</td><td>717806</td><td></td></tr><tr><td>Rabbit, New Zealand White</td><td>Charles River</td><td>Strain Code: 052</td><td></td></tr><tr><td>Scissors</td><td>Aesculap AG, Tuttlingen, Germany</td><td>BC774R</td><td>Bauchdeckenschere ger. 18cm</td></tr><tr><td>Sterile filter, 0.22 &amp;micro;m</td><td>Merck, Darmstadt, Germany</td><td>SLGP033RB</td><td></td></tr><tr><td>&lt;strong&gt;Equipment - Patch-clamp&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Amplfier</td><td>AxonInstruments, Union City, CA, United States</td><td>Axopatch 200B</td><td></td></tr><tr><td>Coverslip &amp;oslash; 50 mm, Thickness No. 1</td><td>VWR International GmbH, Leuven, Belgium</td><td>631-0178</td><td>Borosilicate Glass</td></tr><tr><td>Digitizer Axon Digidata</td><td>Molecular Devices, San Jos&amp;eacute;, CA, United States</td><td>1550A</td><td></td></tr><tr><td>Filter (530/20)</td><td>Leica Microsystems, Wetzlar, Germany</td><td>11513878</td><td>BZ:00</td></tr><tr><td>Filter (630/20)</td><td>Chroma Technology, Bellows Falls, Vermont, United States</td><td>227155</td><td></td></tr><tr><td>Headstage</td><td>AxonInstruments, Union City, CA, United States</td><td>CV203BU</td><td></td></tr><tr><td>Interface</td><td>Scientifica, Uckfield, UK</td><td>1U Rack, 352036</td><td></td></tr><tr><td>LED 525 nm</td><td>Luminus Devices, Sunnyvale, CA, United States</td><td>PT-120-G</td><td></td></tr><tr><td>LED control software</td><td>Essel Research and Development, Toronto, Canada</td><td></td><td></td></tr><tr><td>LED control system</td><td>custom-made</td><td></td><td></td></tr><tr><td>Micropipette Puller</td><td>Narishige Co., Tokyo, Japan</td><td>PP-830</td><td></td></tr><tr><td>Microscope inverted</td><td>Leica Microsystems, Wetzlar, Germany</td><td>DMI4000B</td><td></td></tr><tr><td>Motorised Micromanipulator</td><td>Scientifica, Uckfield, UK</td><td>PatchStar</td><td></td></tr><tr><td>Optical power meter</td><td>Thorlabs, Newton, NJ, United States</td><td>PM100D</td><td></td></tr><tr><td>Silicone Grease</td><td>RS Components, Corby, UK</td><td>494-124</td><td></td></tr><tr><td>Silver wire</td><td>A-M Systems, Sequim, WA, United States</td><td>787500</td><td>Silver, Bare 0.015&#039;&#039;, Coated 0.0190&#039;&#039;, Length 25 Feet</td></tr><tr><td>Soda lime glass capillaries</td><td>Vitrex Medical A/S, Vasekaer, Denmark</td><td>160213 BRIS, ISO12772</td><td>1.55 OD x 1.15 ID x 75 L mm</td></tr><tr><td>Software Axon pClamp</td><td>Molecular Devices, San Jos&amp;eacute;, CA, United States</td><td>Version 10.5</td><td></td></tr><tr><td>Software MatLab2017</td><td>The MathWorks, Inc.</td><td></td><td></td></tr><tr><td>Stage micrometer</td><td>Graticules Optics LTD, Tonbridge, UK</td><td>1 mm</td><td></td></tr><tr><td>&lt;strong&gt;Equipment - Carbon fiber&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Carbon fibers</td><td>provided from Prof. Jean-Yves Le Guennec</td><td></td><td>BZ:00</td></tr><tr><td>Digitizer Axon Digidata</td><td>Molecular Devices, Sunnyvale, CA, United States</td><td>1550B</td><td></td></tr><tr><td>Filter (530/20)</td><td>Leica Microsystems, Wetzlar, Germany</td><td>11513878</td><td></td></tr><tr><td>Filter (630/20)</td><td>Chroma Technology, Bellows Falls, Vermont, United States</td><td>227155</td><td></td></tr><tr><td>Fluorescence System Interface</td><td>IonOptix, Milton, MA United States</td><td>FSI-800</td><td>2.0 OD x 1.16 ID x 100 L mm</td></tr><tr><td>Force Transducer System</td><td>Aurora Scientific Inc., Ontario, Canada</td><td>406A</td><td></td></tr><tr><td>Glass capillaries for force measurements</td><td>Harvard Apparatus, Holliston, Massachusetts, United States</td><td>GC200F-10</td><td></td></tr><tr><td>Interface National Instruments</td><td>National Instruments, Budapest, Hungary</td><td>BNC-2110</td><td></td></tr><tr><td>LED 525 nm</td><td>Luminus Devices, Sunnyvale, CA, United States</td><td>PT-120-G</td><td></td></tr><tr><td>LED control box</td><td>Essel Research and Development, Toronto, Canada</td><td></td><td></td></tr><tr><td>LED control system</td><td>custom-made</td><td></td><td></td></tr><tr><td>Microcontroller</td><td>Parallax Inc., Rocklin, California, United States</td><td>Propeller</td><td></td></tr><tr><td>Micropipette Puller</td><td>Narishige Co., Tokyo, Japan</td><td>PC-10</td><td></td></tr><tr><td>Microscope inverted</td><td>Leica Microsystems, Wetzlar, Germany</td><td>DMI4000B</td><td></td></tr><tr><td>MyoCam-S camera</td><td>IonOptix, Dublin, Ireland</td><td></td><td></td></tr><tr><td>MyoCam-S camera Power</td><td>IonOptix, Milton, MA, United States</td><td>MCS-100</td><td></td></tr><tr><td>MyoPacer Field Stimulator</td><td>IonOptix Cooperation, Milton, MA, United States</td><td>MYP100</td><td></td></tr><tr><td>Piezo Motor</td><td>Physik Instrumente (PI) GmbH &amp;amp; Co. KG, Karlsruhe, Germany</td><td>E-501.00</td><td></td></tr><tr><td>Silicone Grease</td><td>RS Components, Corby, UK</td><td>494-124</td><td></td></tr><tr><td>Software Axon pClamp</td><td>Molecular Devices, San Jos&amp;eacute;, CA, United States</td><td>Version 10.5</td><td></td></tr><tr><td>Software IonWizard</td><td>IonOptix, Dublin, Ireland</td><td>Version 6.6.10.125</td><td></td></tr><tr><td>Software MatLab2017</td><td>The MathWorks, Inc.</td><td></td><td></td></tr><tr><td>Stage micrometer</td><td>Graticules Optics LTD, Tonbridge, UK</td><td>1 mm</td><td></td></tr><tr><td>&lt;strong&gt;Chemicals&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Adenosine</td><td>Sigma-Aldrich, St. Louis, Missouri, United States</td><td>A9251-100G</td><td></td></tr><tr><td>Bovine serum albumin</td><td>Sigma-Aldrich, St. Louis, Missouri, United States</td><td>A7030-50G</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Honeywell Fluka, Muskegon, MI, USA</td><td>21114-1L</td><td></td></tr><tr><td>L-Carnitine hydrochloride</td><td>Sigma-Aldrich, St. Louis, Missouri, United States</td><td>C9500-25G</td><td></td></tr><tr><td>Collagenase type 2, 315 U/mg</td><td>Worthington, Lakewood, NJ, USA</td><td>LS004177</td><td></td></tr><tr><td>Creatine</td><td>Sigma-Aldrich, St. Louis, Missouri, United States</td><td>C0780-50G</td><td></td></tr><tr><td>Cytosine-&amp;beta;-D-arabinofuranoside</td><td>Sigma-Aldrich, St. Louis, Missouri, United States</td><td>C1768-100MG</td><td></td></tr><tr><td>EGTA</td><td>Carl Roth GmbH + Co. KG, Karlsruhe, Germany</td><td>3054.3</td><td></td></tr><tr><td>Esketamine hydrochloride, Ketanest S 25 mg/mL</td><td>Pfizer Pharma PFE GmbH, Berlin, Germany</td><td>PZN-07829486</td><td></td></tr><tr><td>Fetal Bovine Serum</td><td>Sigma-Aldrich, St. Louis, Missouri, United States</td><td>F9665</td><td></td></tr><tr><td>Gentamycin 50 mg/mL</td><td>Gibco, Life Technologies, Waltham, MA, USA</td><td>15750-037</td><td></td></tr><tr><td>Glucose</td><td>Sigma-Aldrich, St. Louis, Missouri, United States</td><td>G7021-1KG</td><td></td></tr><tr><td>Heparin-Sodium, 5,000 IU/mL</td><td>Braun Melsungen AG, Melsungen, Germany</td><td>PZN-03029843</td><td></td></tr><tr><td>HEPES</td><td>Sigma-Aldrich, St. Louis, Missouri, United States</td><td>H3375-1KG</td><td></td></tr><tr><td>Insulin (bovine pancreas)</td><td>Sigma-Aldrich, St. Louis, Missouri, United States</td><td>I6634-50MG</td><td></td></tr><tr><td>K-aspartate</td><td>Sigma-Aldrich, St. Louis, Missouri, United States</td><td>A6558-25G</td><td></td></tr><tr><td>KCl</td><td>VWR International GmbH, Leuven, Belgium</td><td>26764.260</td><td>1 mg/mL</td></tr><tr><td>KOH</td><td>Honeywell Fluka, Muskegon, MI, USA</td><td>35113-1L</td><td></td></tr><tr><td>Laminin from Engelbreth-Holm-Swarm murine sarcoma basement membrane</td><td>Sigma-Aldrich, St. Louis, Missouri, United States</td><td>L2020-1MG</td><td></td></tr><tr><td>M199-Medium</td><td>Sigma-Aldirch, St. Louis, Missouri, United States</td><td>M4530</td><td></td></tr><tr><td>Mg-ATP</td><td>Sigma-Aldrich, St. Louis, Missouri, United States</td><td>A9187-1G</td><td></td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma-Aldrich, St. Louis, Missouri, United States</td><td>63069-500ML</td><td></td></tr><tr><td>NaCl</td><td>Fisher Scientific, Loughborough, Leics., UK</td><td>10428420</td><td></td></tr><tr><td>NaCl-Solution 0.9%, Isotone Kochsalz-L&amp;ouml;sung 0.9%</td><td>Braun Melsungen AG, Melsungen, Germany</td><td>3200950</td><td></td></tr><tr><td>NaOH</td><td>AppliChem GmbH, Darmstadt, Germany</td><td>A6579</td><td>without Ca&lt;sup&gt;2+&lt;/sup&gt;/Mg&lt;sup&gt;2+&lt;/sup&gt;</td></tr><tr><td>Na-pyruvat</td><td>Sigma-Aldrich, St. Louis, Missouri, United States</td><td>P2256-100MG</td><td></td></tr><tr><td>Phosphate Buffered Saline</td><td>Sigma-Aldrich, St. Louis, Missouri, United States</td><td>D1408-500ML</td><td></td></tr><tr><td>Poly(2-hydroxyethyl methacrylate)</td><td>Sigma, Poole, UK</td><td>192066</td><td></td></tr><tr><td>Protease XIV from Streptomyces griseus</td><td>Sigma-Aldrich, St. Louis, Missouri, United States</td><td>P5147-1G</td><td></td></tr><tr><td>Taurine</td><td>Sigma-Aldrich, St. Louis, Missouri, United States</td><td>T0625-500G</td><td></td></tr><tr><td>Thiopental Inresa 0.5 g</td><td>Inresa Arzneimittel GmbH, Freiburg, Germany</td><td>PZN-11852249</td><td></td></tr><tr><td>Xylazine hydrochloride, Rompun 2%</td><td>Bayer Vital GmbH, Leverkusen, Germany</td><td>PZN-01320422</td><td></td></tr></tbody></table>

electromechanical assessment of optogenetically modulated cardiomyocyte activity

Over the past two decades, optogenetic tools have been established as potent means to modulate cell-type specific activity in excitable tissues, including the heart. While Channelrhodopsin-2 (ChR2) is a common tool to depolarize the membrane potential in cardiomyocytes (CM), potentially eliciting action potentials (AP), an effective tool for reliable silencing of CM activity has been missing. It has been suggested to use anion channelrhodopsins (ACR) for optogenetic inhibition. Here, we describe a protocol to assess the effects of activating the natural ACR GtACR1 from Guillardia theta in cultured rabbit CM. Primary readouts are electrophysiological patch-clamp recordings and optical tracking of CM contractions, both performed while applying different patterns of light stimulation. The protocol includes CM isolation from rabbit heart, seeding and culturing of the cells for up to 4 days, transduction via adenovirus coding for the light-gated chloride channel, preparation of patch-clamp and carbon fiber setups, data collection and analysis. Using the patch-clamp technique in whole-cell configuration allows one to record light-activated currents (in voltage-clamp mode, V-clamp) and AP (current-clamp mode, I-clamp) in real time. In addition to patch-clamp experiments, we conduct contractility measurements for functional assessment of CM activity without disturbing the intracellular milieu. To do so, cells are mechanically preloaded using carbon fibers and contractions are recorded by tracking changes in sarcomere length and carbon fiber distance. Data analysis includes assessment of AP duration from I-clamp recordings, peak currents from V-clamp recordings and force calculation from carbon fiber measurements. The described protocol can be applied to the testing of biophysical effects of different optogenetic actuators on CM activity, a prerequisite for the development of a mechanistic understanding of optogenetic experiments in cardiac tissue and whole hearts.

GtACR1-eGFP was expressed in cultured rabbit CM (Figure 6 insert) and photocurrents were measured with the patch-clamp technique. Photoactivation of GtACR1 shows large inward directed currents at -74 mV. In Figure 6A peak current (IP) at 4 mW/mm2 is 245 pA. AP were triggered either electrically (Figure 6B) or optically (Figure 6C) with current injections 1.5 times the threshold, or short depolarizing light pulses of 10 ms, respectively. Analyzing APD values, electrically paced CM show an APD 20 of 0.24 &#177; 0.08 s and an APD 90 of 0.75 &#177; 0.17 s, whereas optically paced CM show an APD 20 of 0.31 &#177; 0.08 s and an APD 90 of 0.81 &#177; 0.19 s (SE, n = 5, N = 2, in the here presented example APD 20electrical = 0.17 s; APD 20optical = 0.27 s and APD 90electrical = 0.61 s; APD 90optical = 0.68 s; Figure 6D). Optically paced CM show a slower AP onset (Figure 6D). CM activation was inhibited upon sustained illumination (for 64 s, 4 mW/mm2) by polarizing the membrane potential towards the reversal potential of chloride, here -58 mV (Figure 6E). Higher current injections than 1.5 times the threshold do not elicit AP generation (Figure 6F). Generated peak forces were determined from carbon fiber bending (Figure 7B,C,E). The CM generated 232 &#181;N/mm2 upon electrical pacing (Figure 7B) and 261 &#181;N/mm2 following optical pacing (Figure 7C). Prolonged green-light pulses inhibit contractions (Figure 7E). Following optical inhibition for 64 s, reoccurring contractions generate a lower contractile force, and force values recover towards baseline after ~10 contractions (pacing at 0.25 Hz, Figure 7D) in keeping with diastolic calcium loss from rabbit CM.
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/60490/60490fig6a.jpg" /> 
Figure 6: Representative patch-clamp recordings of electrically and optically paced/inhibited CM. (A) Representative photocurrent at -74 mV using a light pulse of 300 ms, 4 mW/mm2. IP indicates the peak current. The insert shows a GtACR1-eGFP positive cell. (B) Representative AP recording at 0 pA using a current ramp of 10 ms, 0.6 nA to electrically pace the CM. (C) Representative AP recording at 0 pA using light pulses of 10 ms, 0.4 mW/mm2. (D) Top graph shows the overlay of the 10th AP of electrically (blue) and optically (green) activated CM. AP were aligned by the maximum change in membrane potential (dV/dt max). Bottom graph shows the difference of membrane potential between optically and electrically triggered AP (Eoptical-Eelectrical). (E) Electrically triggered AP were inhibited under sustained light of 64 s, 4 mW/mm2. (F) AP are inhibited by higher current injections than 1.5 times the threshold (from 0.7 nA in steps of 0.1 nA to 2.2 nA) under sustained light. <a href="https://www.jove.com/files/ftp_upload/60490/60490fig6alarge.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/60490/60490fig7a.jpg" /> 
Figure 7: Representative data from carbon fiber recordings of optically and electrically paced/inhibited CM. (A) Display in the data acquisition software. Image (I) shows the measured CM with the window for calculating sarcomere length. Cell width is labelled in orange. (1) Range of relevant frequencies. (2) FFT power spectrum shows the frequency of the sarcomere spacing on the cell. The average sarcomere length is calculated from the peak frequency. (3) Sarcomere length tracking window. (4) Intensity trace. (5) The intensity trace multiplied by a Hamming window is the windowed intensity trace. Scheme (II) shows the elliptical cross-section of the cell. Width in orange and thickness in dashed blue. Image (III) shows the position of the carbon fibers with the respective detection boxes, left in red and right in green. (6) Intensity trace. (7) First derivative of intensity trace (see data acquisition software manual). (B) Representative trace of electrically elicited contractions. Panel (I) shows the sarcomere length shortening, panel (II) the distance between the two carbon fibers. (C) Representative trace of optically elicited contractions (525 nm, 0.25 Hz, 10 ms, 6 mW/mm2). Panel (I) shows the sarcomere length shortening, panel (II) the distance between the two carbon fibers. (D) Generated peak force from contraction 1 to 11 after a pause caused by inhibition of AP generation. (E) Representative trace of optical inhibition of contractions under sustained illumination (525 nm, 64 s, 6 mW/mm2). Panel (I) shows the sarcomere length shortening, panel (II) the length between the two carbon fibers. <a href="https://www.jove.com/files/ftp_upload/60490/60490fig7alarge.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1">
	<tbody>
		<tr>
			<td>A</td>
			<td>area</td>
		</tr>
		<tr>
			<td>ACR</td>
			<td>anion channelrhodopsin</td>
		</tr>
		<tr>
			<td>AP</td>
			<td>action potential</td>
		</tr>
		<tr>
			<td>APD</td>
			<td>action potential duration</td>
		</tr>
		<tr>
			<td>CFU</td>
			<td>colony forming unit</td>
		</tr>
		<tr>
			<td>ChR</td>
			<td>channelrhodopsin</td>
		</tr>
		<tr>
			<td>CM</td>
			<td>cardiomyocyte</td>
		</tr>
		<tr>
			<td>eGFP</td>
			<td>enhanced green fluorescent protein</td>
		</tr>
		<tr>
			<td>ESD</td>
			<td>end systolic cell deformation</td>
		</tr>
		<tr>
			<td>EU</td>
			<td>endotoxin units</td>
		</tr>
		<tr>
			<td>F</td>
			<td>force</td>
		</tr>
		<tr>
			<td>FFT</td>
			<td>fast Fourier transform</td>
		</tr>
		<tr>
			<td>GtACR</td>
			<td>Guillardia theta anion channelrhodopsin</td>
		</tr>
		<tr>
			<td>GUI</td>
			<td>graphical user interface</td>
		</tr>
		<tr>
			<td>I-clamp</td>
			<td>current-clamp</td>
		</tr>
		<tr>
			<td>IU</td>
			<td>international units</td>
		</tr>
		<tr>
			<td>MOI</td>
			<td>multiplicity of infection</td>
		</tr>
		<tr>
			<td>poly-HEMA</td>
			<td>poly(2-hydroxyethyl methacrylate)</td>
		</tr>
		<tr>
			<td>V-clamp</td>
			<td>voltage-clamp</td>
		</tr>
	</tbody>
</table>
Table 5: List of abbreviations.
<img alt="Supplemental Figure 1" src="/files/ftp_upload/60490/60490suppfig1a.jpg" /> 
Supplemental Figure 1: Light intensity measurements with optical power meter. (A) Measurement of 10 ms light pulses at 4 mW/mm2. (B) Measurement of sustained illumination of 64 s at 4 mW/mm2. <a href="https://www.jove.com/files/ftp_upload/60490/60490suppfig1alarge.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Supplemental Figure 2" src="/files/ftp_upload/60490/60490suppfig2a.jpg" /> 
Supplemental Figure 2: Properties of freshly isolated CM and their structural adaptation in culture. (A) AP recording of a freshly isolated CM (APD 20 of 1.11 &#177; 0.34 s, APD 90 of 1.96 &#177; 0.32 s, n = 7, N = 2). Mean resting membrane potential of -79.3 &#177; 0.8 mV (n = 7, N = 2). (B) Carbon fiber recording of an electrically paced freshly isolated CM. Mean peak force of 205 &#177; 78 &#181;N/mm2 (n = 7, N = 2). (C) Confocal images of a freshly isolated CM (I); untransduced (II) and transduced (III) CM after 48 hours in culture. <a href="https://www.jove.com/files/ftp_upload/60490/60490suppfig2alarge.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplemental Material: MatLab script to determine APD and resting membrane potential. <a href="https://www.jove.com/files/ftp_upload/60490/SupplementalMaterial.zip" target="_blank">Please click here to download this file.</a>

Watch this Scientific Journal Video about Electromechanical Assessment of Optogenetically Modulated Cardiomyocyte Activity at JoVE.com

Electromechanical Assessment of Optogenetically Modulated Cardiomyocyte Activity

We present a protocol for evaluating the electromechanical effects of GtACR1 activation in rabbit cardiomyocytes. We provide detailed information on cell isolation, culturing and adenoviral transduction, and on functional experiments with the patch-clamp and carbon-fiber techniques.

Over the past two decades, optogenetic tools have been established as potent means to modulate cell-type specific activity in excitable tissues, including ...

electromechanical-assessment-optogenetically-modulated-cardiomyocyte

Research

JoVE Journal

Medicine

7.9K Views.  Medical Center-University of Freiburg. Het gebruik van het protocol als biofysisch effect van verschillende optogenetische actuatoren op cardiomyocyteactiviteit kan worden getest om te helpen bij de ontwikkeling van optogenetische experimenten in hartweefsel en hele harten. Door de patchklemtechniek te combineren met sarcomere tracking en koolstofvezel ondersteunde krachtmetingen, kunnen we de effecten van GtACR1-fotoactiviteit op de elektra en mechanica van ventriculaire cardiomyocyten bestuderen. Optogenetische remming kan mogel...

Video: Electromechanical Assessment of Optogenetically Modulated Cardiomyocyte Activity

Simultaneous Electrical and Mechanical Stimulation to Enhance Cells' Cardiomyogenic Potential

Cardiovascular diseases are the leading cause of death in developed countries. Consequently, the demand for effective cardiac cell therapies has motivated researchers in the stem cell and bioengineering fields to develop in vitro high-fidelity human myocardium for both basic research and clinical applications. However, the immature phenotype of cardiac cells is a limitation on obtaining tissues that functionally mimic the adult myocardium, which is mainly characterized by mechanical and electrical signals. Thus, the purpose of this protocol is to prepare and mature the target cell population through electromechanical stimulation, recapitulating physiological parameters. Cardiac tissue engineering is evolving toward more biological approaches, and strategies based on biophysical stimuli, thus, are gaining momentum. The device developed for this purpose is unique and allows individual or simultaneous electrical and mechanical stimulation, carefully characterized and validated. In addition, although the methodology has been optimized for this stimulator and a specific cell population, it can easily be adapted to other devices and cell lines. The results here offer evidence of the increased cardiac commitment of the cell population after electromechanical stimulation. Electromechanically stimulated cells show an increased expression of main cardiac markers, including early, structural, and calcium-regulating genes. This cell conditioning could be useful for further regenerative cell therapy, disease modeling, and high-throughput drug screening.

Here we present a protocol for training a cell population using electrical and mechanical stimuli emulating cardiac physiology. This electromechanical stimulation enhances the cardiomyogenic potential of the treated cells and is a promising strategy for further cell therapy, disease modeling, and drug screening.

Cardiovascular diseases are the leading cause of death in developed countries. Consequently, the demand for effective cardiac cell therapies has motivated ...

Bioengineering

Optical Imaging of Isolated Murine Ventricular Myocytes

The ability to isolate adult cardiac myocytes has permitted researchers to study a variety of cardiac pathologies at the single cell level. While advances in calcium sensitive dyes have permitted the robust optical recording of single cell calcium dynamics, recording of robust transmembrane optical voltage signals has remained difficult. Arguably, this is because of the low single to noise ratio, phototoxicity, and photobleaching of traditional potentiometric dyes. Therefore, single cell voltage measurements have long been confined to the patch clamp technique which while the gold standard, is technically demanding and low throughput. However, with the development of novel potentiometric dyes, large, fast optical responses to changes in voltage can be obtained with little to no phototoxicity and photobleaching. This protocol describes in detail how to isolate adult murine myocytes which can be used for cellular shortening, calcium, and optical voltage measurements. Specifically, the protocol describes how to use a ratiometric calcium dye, a single-excitation calcium dye, and a single excitation voltage dye. This approach can be used to assess the cardiotoxicity and arrhythmogenicity of various chemical agents. While phototoxicity is still an issue at the single cell level, methodology is discussed on how to reduce it.

We present the methodology for the isolation of murine myocytes and how to obtain voltage or calcium traces simultaneously with sarcomere shortening traces using fluorescence photometry with simultaneous digital cell geometry measurements.

The ability to isolate adult cardiac myocytes has permitted researchers to study a variety of cardiac pathologies at the single cell level. While advances ...

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

Conventional intracellular microelectrode techniques to quantify cardiomyocyte electrophysiology are extremely complex, labor intensive, and typically carried out in low throughput. Rapid and ongoing expansion of induced pluripotent stem cell (iPSC) technology presents a new standard in cardiovascular research and alternate methods are now necessary to increase throughput of electrophysiological data at a single cell level. VF2.1Cl is a recently derived voltage sensitive dye which provides a rapid single channel, high magnitude response to fluctuations in membrane potential. It possesses kinetics superior to those of other existing voltage indicators and makes available functional data equivalent to that of traditional microelectrode techniques. Here, we demonstrate simplified, non-invasive action potential characterization in externally paced human iPSC derived cardiomyocytes using a modular and highly affordable photometry system.

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 ...

Impact of Intracardiac Neurons on Cardiac Electrophysiology and Arrhythmogenesis in an Ex Vivo Langendorff System

Since its invention in the late 19th century, the Langendorff ex vivo heart perfusion system continues to be a relevant tool for studying a broad spectrum of physiological, biochemical, morphological, and pharmacological parameters in centrally denervated hearts. Here, we describe a setup for the modulation of the intracardiac autonomic nervous system and the assessment of its influence on basic electrophysiology, arrhythmogenesis, and cyclic adenosine monophosphate (cAMP) dynamics. The intracardiac autonomic nervous system is modulated by the mechanical dissection of atrial fat pads-in which murine ganglia are located mainly&#8212;or by the usage of global as well as targeted pharmacological interventions. An octapolar electrophysiological catheter is introduced into the right atrium and the right ventricle, and epicardial-placed multi-electrode arrays (MEA) for high-resolution mapping are used to determine cardiac electrophysiology and arrhythmogenesis. F&#246;rster resonance energy transfer (FRET) imaging is performed for the real-time monitoring of cAMP levels in different cardiac regions. Neuromorphology is studied by means of antibody-based staining of whole hearts using neuronal markers to guide the identification and modulation of specific targets of the intracardiac autonomic nervous system in the performed studies. The ex vivo Langendorff setup allows for a high number of reproducible experiments in a short time. Nevertheless, the partly open nature of the setup (e.g., during MEA measurements) makes constant temperature control difficult and should be kept to a minimum. This described method makes it possible to analyze and modulate the intracardiac autonomic nervous system in decentralized hearts.

Impact of Intracardiac Neurons on Cardiac Electrophysiology and Arrhythmogenesis in an Ex Vivo Langendorff System

Here, we present a protocol for the modulation of the intracardiac autonomic nervous system and the assessment of its influence on basic electrophysiology, arrhythmogenesis, and cAMP dynamics using an ex vivo Langendorff setup.

Since its invention in the late 19th century, the Langendorff ex vivo heart perfusion system continues to be a relevant tool for studying a broad spectrum ...

In Vitro Assessment of Cardiac Function Using Skinned Cardiomyocytes

In this article, we describe the steps required to isolate a single permeabilized (&#34;skinned&#34;) cardiomyocyte and attach it to a force-measuring apparatus and a motor to perform functional studies. These studies will allow measurement of cardiomyocyte stiffness (passive force) and its activation with different calcium (Ca2+)-containing solutions to determine, amongst others: maximum force development, myofilament Ca2+-sensitivity (pCa50), cooperativity (nHill) and the rate of force redevelopment (ktr). This method also enables determination of the effects of drugs acting directly on myofilaments and of the expression of exogenous recombinant proteins on both active and passive properties of cardiomyocytes. Clinically, skinned cardiomyocyte studies highlight the pathophysiology of many myocardial diseases and allow in vitro assessment of the impact of therapeutic interventions targeting the myofilaments. Altogether, this technique enables the clarification of cardiac pathophysiology by investigating correlations between in vitro and in vivo parameters in animal models and human tissue obtained during open heart or transplant surgery.

This protocol aims to describe step-by-step the technique of extraction and assessment of cardiac function using skinned cardiomyocytes. This methodology allows measurement and acutemodulation of myofilament function using small frozen biopsies that can be collected from different cardiac locations, from mice to men.

In this article, we describe the steps required to isolate a single permeabilized (&#34;skinned&#34;) cardiomyocyte and attach it to a force-measuring ...

Optocardiography and Electrophysiology Studies of Ex Vivo Langendorff-perfused Hearts

Small animal models are most commonly used in cardiovascular research due to the availability of genetically modified species and lower cost compared to larger animals. Yet, larger mammals are better suited for translational research questions related to normal cardiac physiology, pathophysiology, and preclinical testing of therapeutic agents. To overcome the technical barriers associated with employing a larger animal model in cardiac research, we describe an approach to measure physiological parameters in an isolated, Langendorff-perfused piglet heart. This approach combines two powerful experimental tools to evaluate the state of the heart: electrophysiology (EP) study and simultaneous optical mapping of transmembrane voltage and intracellular calcium using parameter sensitive dyes (RH237, Rhod2-AM). The described methodologies are well suited for translational studies investigating the cardiac conduction system, alterations in action potential morphology, calcium handling, excitation-contraction coupling and the incidence of cardiac alternans or arrhythmias.

The objective of this study was to establish a method for investigating cardiac dynamics using a translational animal model. The described experimental approach incorporates dual-emission optocardiography in conjunction with an electrophysiological study to assess electrical activity in an isolated, intact porcine heart model.

Small animal models are most commonly used in cardiovascular research due to the availability of genetically modified species and lower cost compared to ...

A Cardiac Microphysiological System for Studying Ca2+ Propagation via Non-genetic Optical Stimulation

In vitro cardiac microphysiological models are highly reliable for scientific research, drug development, and medical applications. Although widely accepted by the scientific community, these systems are still limited in longevity due to the absence of non-invasive stimulation techniques. Phototransducers provide an efficient stimulation method, offering a wireless approach with high temporal and spatial resolution while minimizing invasiveness in stimulation processes. In this manuscript, we present a fully optical method for stimulating and detecting the activity of an in vitro cardiac microphysiological model. Specifically, we fabricated engineered laminar anisotropic tissues by seeding human-induced pluripotent stem cell-derived cardiomyocytes (hiPSC-CMs) generated in a 3D bioreactor suspension culture. We employed a phototransducer, an amphiphilic azobenzene derivative, named Ziapin2, for stimulation and a Ca2+ dye (X-Rhod 1) for monitoring the system&#39;s response. The results demonstrate that Ziapin2 can photomodulate Ca2+ responses in the employed system without compromising tissue integrity, viability, or behavior. Furthermore, we showed that the light-based stimulation approach offers a similar resolution compared to electrical stimulation, the current gold standard. Overall, this protocol opens promising perspectives for the application of Ziapin2 and material-based photostimulation in cardiac research.

Using light to control cardiac cells and tissue enables non-contact stimulation, thereby preserving the natural state and function of the cells, making it a valuable approach for both basic research and therapeutic applications.

In vitro cardiac microphysiological models are highly reliable for scientific research, drug development, and medical applications. Although widely ...

Advanced Cardiac Rhythm Management by Applying Optogenetic Multi-Site Photostimulation in Murine Hearts

Ventricular tachyarrhythmias are a major cause of mortality and morbidity worldwide. Electrical defibrillation using high-energy electric shocks is currently the only treatment for life-threatening ventricular fibrillation. However, defibrillation may have side-effects, including intolerable pain, tissue damage, and worsening of prognosis, indicating a significant medical need for the development of more gentle cardiac rhythm management strategies. Besides energy-reducing electrical approaches, cardiac optogenetics was introduced as a powerful tool to influence cardiac activity using light-sensitive membrane ion channels and light pulses. In the present study, a robust and valid method for successful photostimulation of Langendorff perfused intact murine hearts will be described based on multi-site pacing applying a 3 x 3 array of micro light-emitting diodes (micro-LED). Simultaneous optical mapping of epicardial membrane voltage waves allows the investigation of the effects of region-specific stimulation and evaluates the newly induced cardiac activity directly on-site. The obtained results show that the efficacy of defibrillation is strongly dependent on the parameters chosen for photostimulation during a cardiac arrhythmia. It will be demonstrated that the illuminated area of the heart plays a crucial role for termination success as well as how the targeted control of cardiac activity during illumination for modifying arrhythmia patterns can be achieved. In summary, this technique provides a possibility to optimize the on-site mechanism manipulation on the way to real-time feedback control of cardiac rhythm and, regarding the region specificity, new approaches in reducing the potential harm to the cardiac system compared to the usage of non-specific electrical shock applications.

This work reports a method for controlling the cardiac rhythm of intact murine hearts of transgenic channelrhodopsin-2 (ChR2) mice using local photostimulation with a micro-LED array and simultaneous optical mapping of epicardial membrane potential.

Ventricular tachyarrhythmias are a major cause of mortality and morbidity worldwide. Electrical defibrillation using high-energy electric shocks is ...

Measurement of Heart Contractility in Isolated Adult Human Primary Cardiomyocytes

The evaluation of changes in heart contractility is essential during preclinical development for new cardiac- and non-cardiac-targeted compounds. This paper describes a protocol for assessing changes in contractility in adult human primary ventricular cardiomyocytes utilizing the MyoBLAZER, a non-invasive optical method that preserves the normal physiology and pharmacology of the cells. This optical recording method continuously measures contractility transients from multiple cells in parallel, providing both medium-throughput and valuable information for each individual cell in the field of view, enabling the real-time tracking of drug effects. The cardiomyocyte contractions are induced by paced electrical field stimulation, and the acquired bright field images are fed to an image-processing software that measures the sarcomere shortening across multiple cardiomyocytes. This method rapidly generates different endpoints related to the kinetics of contraction and relaxation phases, and the resulting data can then be interpreted in relation to different concentrations of a test article. This method is also employed in the late stages of preclinical development to perform follow-up mechanistic studies to support ongoing clinical studies. Thus, the adult human primary cardiomyocyte-based model combined with the optical system for continuous contractility monitoring has the potential to contribute to a new era of in vitro cardiac data translatability in preclinical medical therapy development.

This protocol describes how to measure contractility in adult human primary cardiomyocytes from donor hearts with the MyoBLAZER system, a reliable platform for assessing drug-induced changes in contractility during preclinical development.

The evaluation of changes in heart contractility is essential during preclinical development for new cardiac- and non-cardiac-targeted compounds. This ...

Subtype-specific Optical Action Potential Recordings in Human Induced Pluripotent Stem Cell-derived Ventricular Cardiomyocytes

Cardiomyocytes generated from human induced pluripotent stem cells (iPSC-CMs) are an emerging tool in cardiovascular research. Rather than being a homogenous population of cells, the iPSC-CMs generated by current differentiation protocols represent a mixture of cells with ventricular-, atrial-, and nodal-like phenotypes, which complicates phenotypic analyses. Here, a method to optically record action potentials specifically from ventricular-like iPSC-CMs is presented. This is achieved by lentiviral transduction with a construct in which a genetically-encoded voltage indicator is under the control of a ventricular-specific promoter element. When iPSC-CMs are transduced with this construct, the voltage sensor is expressed exclusively in ventricular-like cells, enabling subtype-specific optical membrane potential recordings using time-lapse fluorescence microscopy.

Here we present a method to optically image action potentials, specifically in ventricular-like induced pluripotent stem cell-derived cardiomyocytes. The method is based on the promoter-driven expression of a voltage-sensitive fluorescent protein.

Electromechanical Assessment of Optogenetically Modulated Cardiomyocyte Activity

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