Electromechanical Assessment of Optogenetically Modulated Cardiomyocyte Activity

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 (Regierungspräsidium Freiburg, X-16/10R, Germany).

1. Solutions for cell isolation

1. Prepare the solutions for the cell isolation with water of the following requirements (Table 1) and according to the ionic compositions listed in Table 2.
   NOTE: CaCl₂ and MgCl₂ are added from 1 M stock solutions.

Table 1: Water requirements.

Table 2: Solutions for CM isolation.

2. Adjust all solutions to pH 7.4 at 37 °C and check osmolarity.
   NOTE: Dissolve the enzymes (Collagenase type 2 and Protease XIV) directly before heart excision. Oxygenate all solutions prior to use.

2. Preparation of the Langendorff-perfusion setup

NOTE: The used setup is custom-made. As depicted in Figure 2, the setup consists of three water jacketed reservoirs (1-3), one spiral counter-flow heat exchanger (4) and a water jacketed perfusion vessel (5).

Figure 2: Langendorff-perfusion setup optimized for rabbit cell isolation. (1-3) Water jacketed reservoirs with (1) physiological saline solution, (2) low calcium, high potassium solution and (3) enzyme-containing cardioplegic solution. (4) Spiral counter-flow heat exchanger and (5) water jacketed collecting tank. The inflow of the water jacketed system is the spiral heat exchanger (temperature of solutions leaving the perfusion cannula at the end of the heat exchanger should be constant at 37 °C), followed by the perfusion vessel and the three reservoirs. All solutions are oxygenated (dashed line). Please click here to view a larger version of this figure.

1. Switch on the pump of the water bath to circulate water at 38 °C in the heat exchange system and preheat all solutions to 37 °C.
   NOTE: The temperature at the outflow of (4) must be controlled and constant at 37 °C.
2. Fill the three reservoirs with the respective solution and wash each line (black) with the corresponding solution. Fill the main (blue) line at the end without air bubbles using solution (1).
   NOTE: Oxygenate the solutions prior (10 min) and during use. Fill the line from reservoir (3) to the tap with low calcium, high potassium solution.
3. Prepare a suture to tie the heart around the aorta at the cannula.

3. Cell isolation

1. Prepare the following syringes.
   1. For sedation/anesthesia: Mix 0.5 mL/kg body weight esketamine hydrochloride (25 mg/mL) and 0.2 mL/kg body weight xylazine hydrochloride (2%).
   2. Fill two syringes with 12 mL of NaCl solution (0.9%).
   3. Prepare 6 mL of 12.5 mg/mL Na-thiopental, dissolved in 0.9% NaCl solution.
   4. Fill 0.2 mL of esketamine hydrochloride (25 mg/mL) in a syringe.
   5. Dilute 0.2 mL of Na-heparin (5,000 IU/mL) in 1 mL of 0.9% NaCl solution (end-concentration 1,000 IU/mL).
2. Sedate/anaesthetize rabbits (9-10 weeks, New Zealand white rabbit, female or male, ~2 kg) via intramuscular injection of esketamine hydrochloride and xylazine hydrochloride (step 3.1.1).
   NOTE: Rabbits need at least 10 min to be fully anaesthetized; exact duration depends on their body weight. Confirm anesthesia with the loss of the righting reflex.
3. Shave the chest and the ears where the veins are located.
4. Insert a flexible cannula into the ear vein, fix it with tape and flush it with 0.9% NaCl solution.
5. Inject 1 mL of Na-heparin solution intravenously and flush with 0.9% NaCl solution.
6. Inject 0.2 mL of esketamine hydrochloride, flush again with 0.9% NaCl solution and inject Na-thiopental until apnea.
   NOTE: Rabbit should not respond to the pedal withdrawal reflex.
7. Open the chest at the left side and remove the pericardium.
8. Start the timer when the heart is excised and wash the heart twice in physiological saline solution.
   NOTE: Use scissors with round tips to prevent accidental damage to cardiac tissue.
9. Cannulate the aorta in a bath with physiological saline solution and keep all tissue in solution. Switch on the Langendorff-perfusion system (physiological saline solution (1), speed 24 mL/min).
10. Transfer the heart to the Langendorff-perfusion setup, connect the aorta to the perfusate nozzle, and tightly tie the heart with the suture around the aorta to the cannula (< 1 min).
    NOTE: Pre-fill the cannula with physiological saline solution, ensure that no air bubbles enter the cannula during transportation from the cannulation site to the Langendorff setup, connect bubble-free.
11. Perfuse the heart until all blood is washed out (2-3 min).
12. Switch to low calcium, high potassium solution (2). Perfuse for 2 more min after the heart has stopped beating and switch to enzyme solution.
13. Start recirculating the enzyme solution, after 2 min from start of digestion, back into the reservoir. Decrease speed to 16 mL/min after 5 min of digestion.
14. When the tissue appears soft (40-50 min of digestion), cut the heart off the cannula and separate the left ventricle.
15. Release cells by mechanical dissociation (gently pulling apart the tissue with a pipette and a fine forceps to hold the tissue) in blocking solution.
16. Filter the cell suspension through a mesh (pore size of 1 mm²) and centrifuge for 2 min at 22 x g (gravitational acceleration).
17. Remove the supernatant containing non-myocytes and re-suspend CM in blocking solution.

4. Culturing of CM

NOTE: Perform the following steps under sterile conditions.

1. Dilute Laminin (from Engelbreth-Holm-Swarm murine sarcoma basement membrane, 1 mg/mL) 1:10 in sterile phosphate buffered saline (without Ca²⁺/Mg²⁺) to a final concentration of 100 µg/mL.
2. Prepare culture medium in M199-Medium with the supplements as indicated in Table 3.

Table 3: Cell culture medium.

3. Sterile-filter solution (0.22 µm) and add 5% Fetal Bovine Serum.
4. For patch-clamp experiments autoclave coverslips ø 16 mm, thickness No. 0, coat them with 100 µg/mL laminin directly before culturing.
5. For carbon fiber experiments, coat the Petri dish surface with poly(2-hydroxyethyl methacrylate) (poly-HEMA, 0.12 g/mL in 95:5 EtOH:H₂0) and let it solidify.
   NOTE: Cells do not 'stick' to poly-HEMA coated Petri dishes; this is crucial for their friction-less contraction in cell mechanics studies.
6. After re-suspended CM have settled (~10-15 min), remove the supernatant, and then re-suspend CM in culture medium.
7. Count CM with a Neubauer chamber and seed at a target density of 17,500 cells/mL either on laminin coated coverslips or in poly-HEMA coated Petri dishes.
8. Incubate cells at 37 °C, 5% CO₂ for 3-4 hours. Exchange the medium (37 °C) of coverslip seeded cells.
9. Add adenovirus (type 5) coding for GtACR1-eGFP at a multiplicity of infection (MOI) of 75 and start functional experiments after 48 hours.
   NOTE: After transduction keep the cells in the dark. Use red illumination when working with blue or green light-activated proteins. A commercially available adenoviral delivery system (see Table of Materials) is used to clone the genes encoding GtACR1-eGFP into the adenoviral vector. The insert of interest, here GtACR1-eGFP, is PCR amplified and then combined with an adenoviral vector including a CMV promoter in an IN-Fusion Cloning reaction. The CMV (human cytomegalovirus) promoter is commonly used to drive overexpression of transgenes in mammalian cells. eGFP is an enhanced green fluorescent protein derived from Aequorea victoria with an excitation maximum of 488 nm and an emission maximum at 507 nm. Adenovirus (type 5) was externally produced at Charité-Universitätsmedizin Berlin, Institut für Pharmakologie, Berlin, Prof. Dr. Michael Schupp.
   CAUTION: Adenoviral transduction is categorized as BSL-2 safety level work, and appropriate safety measures are legally required.

5. Functional experiments

NOTE: Recordings are performed using an inverted fluorescence microscope. Filter the transmission light by a red band-pass filter (630/20 nm) in the condenser to avoid co-activation of GtACR1.

1. Patch-clamp setup
   1. Use an amplifier in combination with an analogue-to-digital converter. Use a data acquisition software to record current and voltage data (see Table of Materials).
      NOTE: The recorded data are digitized at 10 kHz and filtered at 5 kHz.
2. Carbon fiber setup
   1. Use a camera to detect carbon fiber position and sarcomere length by tracking changes in optical contrast (carbon fibers appear as darker structures, overlaid on the striated cell pattern). A schematic representation of the setup is shown in Figure 3.
      NOTE: Sarcomere length is calculated in real time using a fast Fourier transform (FFT) of the power spectrum of the striation pattern.

Figure 3: Scheme depicting experimental setup for carbon fiber measurements. (Drawing is not at scale). Two carbon fibers are attached on a cell and their position is controlled by a piezo positioner. The pacer is used for electrical field stimulation. Multi-color LEDs are coupled into the epifluorescence port of the inverted microscope for illumination of cells in the object plane. LED power is controlled via a dedicated control box, which receives digital pulses via the digital output of the digital-analogue-converter (DAC). The DAC communicates via analogue output with the fluorescence system interface. A black-and-white camera (774 pixels by 245 lines) for cellular imaging is connected to the computer to track sarcomere length and carbon fiber bending. Please click here to view a larger version of this figure.

3. Timed illumination
   1. Provide light for fluorescence microscopy and activation of light-gated ion channels via an external custom-built LED control box, comprising three LEDs of different color (460 nm, 525 nm, 640 nm, see Table of Materials).
   2. Modify the microcontroller and graphical user interface (GUI) code for the control box to allow control of the LED via external Time to Live (TTL) pulses, generated in data acquisition software protocols (see Table of Materials). Transmit TTL pulses to the LED control box via the digital-analogue-converter.
   3. Drive the LED and choose the number of pulses via the GUI. Upon receiving the command from the GUI the microcontroller starts a process on a new core. In this process the TTL input as well as a control switch set from the GUI will be continuously checked.
   4. When the TTL input is positive, the microcontroller turns the LED on and then resumes checking the TTL input. Once the TTL signal returns to zero, the microcontroller turns the LED off and reduces the number of pulses left by one. If at any point the control switch is false or the number of pulses is zero, the microcontroller stops this process until a new command is received from the GUI.
   5. Directly couple the LEDs into the backport of the microscope.
4. Determination of light intensity in the object plane
   1. Measure the illuminated area with a stage micrometer (objective magnification 40x, A = 0.8 mm²).
   2. Use an optical power meter (see Table of Materials).
   3. Define the settings for the experiments: excitation wavelength (525 nm), objective magnification (40x), excitation filter (530/20 nm) or mirror, and read out the light power [W] at various LED-input voltages.
   4. Calculate the light intensity [W/mm²] by dividing the light power [W] by the illuminated area [mm²] (here: 0.8 mm²).
      NOTE: Measure the actual light power with the respective protocols in step 5.6 to check if short light pulse durations of 10 ms reach and long durations hold the set value (Supplemental Figure 1).
5. Preparation for patch-clamp experiments
   1. Prepare the following external and internal solutions (Table 4; for water requirements see Table 1).
   2. Adjust the osmolarity with glucose to 300 ± 5 mOsmol/L. Aliquot the internal solution and store at -20 °C.
      NOTE: Keep the internal solution on ice for the day of the recording. Keep the external solution at room temperature. The here described patch-clamp solutions were based on previously used solutions and the Cl^– concentration was changed to lower, more physiological levels⁷. For characterization of ion selectivity of the respective optogenetic actuator, we suggest to vary the concentrations of major ions (e.g., Cl^–, Na⁺, K⁺, H⁺) in the extra- and intracellular solutions¹⁹.
   3. Take off the recording electrode from the pipette holder and remove the silver chloride layer from the silver wire with very fine sandpaper.
      NOTE: Do this step at the beginning of each measuring day.
   4. Connect the wire to the positive pole of a 1.5 V battery and immerse in 3 M KCl solution for silver chloride coating for 10 min.
      NOTE: The negative pole is connected to a reference silver wire immersed in the 3 M KCl solution.
   5. Prepare the measuring chamber: put silicon grease on the frame of the measuring chamber and place a coverslip (diameter: 50 mm, thickness No. 0) on the top of the frame that the chamber is sealed.
   6. Put a reference silver/silver-chloride pellet electrode in the bath and connect it with the head stage.
   7. Pull 1.7 – 2.5 MΩ patch pipettes from soda lime glass capillaries (outer diameter: 1.55 mm, inner diameter: 1.15 mm) with a micropipette puller (see Table of Materials).
   8. Start data acquisition software and adjust the membrane test (pulse 10 mV for 15 ms, baseline 0 mV).

Table 4: Patch-clamp solutions.

6. Protocols for patch-clamp measurements
   1. Record photoactivation protocol in the V-clamp mode at a holding potential of -74 mV. Use light pulses of 300 ms.
      NOTE: We suggest performing V-clamp recordings close to the resting membrane potential of cultured CM (established in I-clamp; in our hands between -79 mV and -77 mV both for transduced and non-transduced CM¹⁹). Freshly isolated cells show a mean resting membrane potential of -79 mV (Supplemental Figure 2, all values after correction for liquid junction potential).
   2. Record AP in I-clamp mode at 0 pA.
      1. For electrical pacing, inject current pulses of 10 ms (ramp from 0 pA to the set value within 10 ms), 0.25 Hz and find the threshold to elicit AP. Record AP by current injections of 50% more than the threshold.
      2. For optical pacing use light pulses of 10 ms, 0.25 Hz at the minimal light intensity to elicit reliable AP.
   3. Record photoinhibition in I-clamp mode at 0 pA. Elicit AP as described in step 5.6.2.1 and apply sustained light for 64 s at 4 mW/mm² after 15 electrically triggered AP.
      NOTE: Figure 6F shows a photoinhibition protocol where during sustained light higher current injections are applied. Starting from 1.5 times the threshold (here: 0.7 nA) the injected current was increased in steps of 0.1 nA (final level: 2.2 nA). At all tested current amplitudes, sustained light application inhibited AP generation.
      1. As a control experiment, pause electrical stimulation for 64 s without light application.
7. Patch-clamp experiments
   NOTE: Perform the following experiments in the dark (red light can be used for blue/green light-activated tools).
   1. Place coverslip with cells in measuring chamber with external solution and select fluorescent CM.
      NOTE: eGFP-positive cells can be detected using a blue LED (460 nm) in combination with a band-pass excitation filter (450 nm – 490 nm), a 510 nm dichroic mirror and a 515 nm long-pass emission filter. If other fluorescent tags are used, use corresponding LED and fluorescence filter sets. If a high transduction efficiency is achieved (in our hands >99% with the GtACR1 adenovirus), there is no need to check eGFP fluorescence before the functional experiments; this avoids potential GtACR1 pre-activation.
   2. Fill the patch pipette with internal solution. Ensure that there are no air bubbles in the tip.
   3. Attach pipette to the pipette holder, inserting the recording silver-chloride coated silver wire in the internal solution.
   4. After reaching the cell-attached configuration, switch to whole-cell mode in the data acquisition software with a holding potential of -74 mV. Rupture the membrane by gently applying negative pressure to access the whole-cell configuration. This is indicated by an immediate increase in the measured capacitance.
   5. Run the protocols described in section 5.6.
8. Carbon fiber technique
   1. Produce carbon fibers.
      1. Use glass capillaries with the following parameters: outer diameter: 2.0 mm, inner diameter: 1.16 mm, length: 100 mm (see Table of Materials). Using a micropipette puller, pull the glass capillary into two pipettes of the same length (total taper length ~11 mm, Figure 5) to a final inner diameter of ~30 µm.
         NOTE: Settings used for the first and second pull are 85.2% (proportion to the maximum output of the puller) and 49.0%, respectively (will depend on the puller, type and age of the filament).
      2. Bend the pipettes up to 45° with a self-made micro forge using settings of 12 V, 24 A (see Figure 4 for details of the pipette bending setup).
         1. Align the capillary (2) on the red line in the orientation circle (5), keep the positioning of the capillary constant so the length of the bend part is always the same after the center of the orientation circle (radius of 4.5 mm).
         2. Bend the capillary up to 45° (green line) by pushing down the tip of the capillary with the bender (3) and forge by heating-up the filament (4) until the capillary captures the 45° angle even after the bender is removed.
      3. Fit the carbon fibers (provided from Prof. Jean-Yves Le Guennec) into the fine tip of the glass capillary under a stereo microscope. Use fine forceps with soft tubing at the end to increase grip and decrease the risk of damaging the fibers.
         NOTE: These fibers are characterized by microstructures, which increase the contact surface between fibers and cells, thereby improving adhesion²⁰.
      4. Cut the carbon fibers at a length of 2 mm and use super glue (cyanoacrylate) to fix the fiber to the front section of the capillary.
         NOTE: The longer the fibers are, the more they bend upon application of the same force.
   2. Calibrate carbon fibers.
      1. Calibrate the carbon fibers using a force transducer with a sensitivity of 0.05 mN/V and a force range of 0 – 0.5 mN (see Table of Materials).
         NOTE: This setup is custom-made in order to measure compression instead of "pull".
      2. Attach the capillary with the carbon fiber to a holder that is controlled by a micromanipulator and a piezo motor.
      3. Place the tip of the fiber in contact with the force sensor, but without producing any force and move the piezo motor in steps of 10 µm (total movement of 60 µm) towards the sensor and read out the measured voltage (E) in Volt.
         NOTE: Make sure the force transducer is contacted by the very tip of the free end of the carbon fiber.
      4. Repeat these measurements three times.
      5. Use Formula 1 to calculate the force for each piezo position (ΔΕ difference of measured voltage between the piezo motor steps):
         
         NOTE: The sensitivity of the force transducer is dependent on the model of the transducer (here: 0.05 mN/V = 50 µN/V). The gain can be set at the controller.
      6. Plot the force [µN] against the piezo position. The slope corresponds to the fiber stiffness [µN/µm].
   3. Record force of contracting CM.
      NOTE: Perform the following experiments in the dark (red light can be used for blue/ green light-activated tools).
      1. Coat the surface of the measuring chamber with poly-HEMA. Fill the measuring chamber with external bath solution and put a few drops of the cultured cell suspension in the chamber (step 4.9).
      2. Attach capillaries with carbon fibers to the stage micromanipulator. Select GtACR1-expressing CM by checking the ability to induce contractions by application of short green-light pulses. Align the carbon fibers near-horizontally to the surface of the measuring chamber.
      3. Lower the first fiber onto the cell surface. Attach the second fiber parallel to the first fiber at the other end of the CM. The ideal alignment is near-perpendicular to the cell axis.
         NOTE: Attach the fiber by gently pushing the cell to the bottom surface. Release the pressure before attaching the second fiber. Do not stretch the cell by attaching the second fiber.
      4. After both fibers are attached on the cell, lift the fibers, so the cell has no contact to the chamber surface anymore and is able to contract without any friction.
      5. Focus the sarcomeres in the data acquisition software (see Table of Materials) and set the sarcomere length tracking window (Figure 7A I (3)) between the fibers.
         NOTE: Resulting FFT power spectrum (Figure 7A I (2)) shows ideally one sharp peak, representing the average sarcomere length.
      6. Track fiber bending using the edge detection module. Set the detection areas with the red and green window and define a threshold (red and green horizontal line) at the first derivative of the light intensity trace (Figure 7A III).
      7. Start to optically pace the cell at 0.25 Hz (if possible, try faster pacing rates) and track sarcomere length and fiber bending.
         NOTE: Fiber holder position, LED trigger and electric stimulation pulses are controlled via the data acquisition software (see Table of Materials).
      8. After recording at least 15 optically elicited contractions, field-stimulate the cell electrically (see Table of Materials). Find the threshold to elicit contractions and record by applying 1.5 times of the threshold voltage.
      9. For the inhibition protocol, apply electrical stimuli to elicit contractions and then expose to sustained light of 64 s (at various light intensities).

Figure 4: Pipette bending setup. (1) The micromanipulator on the left side is used to control the position of the capillary, and a second micromanipulator on the right is used to bend it. (2) Capillary. (3) Bender. (4) Microforge. (5) Orientation circle. Please click here to view a larger version of this figure.

Figure 5: Pipette with carbon fiber. Please click here to view a larger version of this figure.

6. Data analysis

1. Patch-clamp recordings
   NOTE: Correct all recorded and command voltages for the liquid junction potential after the experiment. Determine liquid junction potential in the data acquisition software by using the tool junction potential calculator (for the stated patch-clamp solutions in Table 4: 14.4 mV at 21 °C). Subtract the liquid junction potential from the recorded/command voltage.
   1. For I-clamp AP recordings, check electrical pacing versus optical pacing. Calculate the AP duration (APD) at 20 and 90% repolarization with a custom-written script (Supplemental Material). Determine resting membrane potential and AP amplitude.
      NOTE: Determine the APD as described in Wang K. et al.²¹ The script to load .abf files is generally accessible via the following link: https://de.mathworks.com/matlabcentral/fileexchange/22114-fcollman-abfload. Average APD values for at least 6 AP.
   2. For V-clamp photoactivation, check if baseline is at 0 pA. If not adjust baseline to zero. Analyze the recorded current triggered by 300 ms light pulses at -74 mV. Transfer the data to data analysis software and determine the peak and average stationary current.
2. Carbon fiber experiments
   1. Contraction recordings during optical pacing: Load the recorded data in the data acquisition software and read out the baseline and maxima of the carbon fiber bending and the sarcomere length changes.
      NOTE: Average values for 10 contractions of a stable recording.
   2. Measure the cell width and calculate the cross-sectional area of the cell assuming an elliptical cross-section (Figure 7B II).
      NOTE: The formula for the area of an ellipse is A = π·a·b (Formula 2) where a is the distance from the center to the vertex and b is the distance from the center to the co-vertex. In our case this means a = (width of the cell)/2 and b = (thickness of the cell)/2. According to Nishimura et al.²² the thickness of CM can be estimated to be one third of the cell width so that A = π·(1/2)·width·(1/2)·thickness = π·(1/4)·width·(1/3)·width = π·(1/12)·width².
   3. Calculate the end-systolic force (F):
      
   4. Calculate the end-systolic cell deformation (ESD):
      
      NOTE: Further contractile parameters can be analyzed: resting sarcomere length, time to peak, time to 90% relaxation, fractional sarcomere shortening, maximum velocity of contraction and relaxation (see software acquisition manual).