Optical Imaging of Isolated Murine Ventricular Myocytes

All methods and procedures described in this protocol have been approved by the Institutional Animal Care and Use Committee (IACUC) of Case Western Reserve University.

1. Preparation of Solutions, Instruments, and Coverslips

NOTE: 1x solutions can be used for up to a month.

1. Make 10x Krebs-Henseleit buffer HEPES buffer without calcium (KHB-HB) by adding 68.96 g of NaCl, 3.57 g of KCl, 59.58 g of HEPES, 2.18 g of K₂HPO₄, 3.08 g of MgSO₄ and 19.82 g of glucose to 800 mL of double distilled water in a 1,000 mL flask. After contents are fully dissolved, bring up to volume in a 1,000 mL volumetric flask.
   NOTE: Traditional Krebs Henseleit solution uses sodium bicarbonate as a buffer and the solution in this protocol uses Krebs Henseleit solution with HEPES buffer. Solution is stable for 6 months if sterile filtered.
2. Make 10x Tyrode's solution by adding 86.51 g of NaCl, 0.552 g of NaH₂PO₄, 2.03 g of MgCl₂, 9.91 g of glucose, 4.03 g of KCl, 2.65 g of CaCl₂, and 35.76 g of HEPES to 800 mL of double distilled water in a 1000 mL flask. After contents are fully dissolved, bring up to volume in a 1000 mL volumetric flask.
   NOTE: Solution is stable for 6 months if sterile filtered.
3. Make 1x KHB-HB by measuring out 100 mL of the 10x stock and adding to 875 mL of double distilled water in a 1,000 mL flask. Place flask in 37 °C water bath. Once the solution has reached 37 °C, use NaOH to increase the pH to 7.39. After adjusting the pH, bring solution to volume in a 1000 mL volumetric flask. Sterile filter the solution using a vacuum filtration system.
4. Make 1x Tyrode's solution by measuring out 100 mL of the 10x stock and adding to 875 mL of double distilled water in a 1,000 mL flask. Place flask in 37 °C water bath. Once the solution has reached 37 °C, use NaOH to increase the pH to 7.39. After adjusting the pH, bring solution to volume in a 1,000 mL volumetric flask. Sterile filter using a vacuum filtration system.
5. Make 1x modified Tyrode's solution by measuring out 100 mL of the 10x stock and adding to 875 mL of double distilled water in a 1,000 mL flask. Dissolve 3.07 g of L-glutathione reduced in flask. Place flask in 37 °C water bath. Once the solution has reached 37 °C, use NaOH to increase the pH to 7.39. After adjusting the pH, bring solution to volume in a 1,000 mL volumetric flask. Sterile filter the solution using a vacuum filtration system.
6. Make 100 mM blebbistatin stock solution by adding 855 µL of dimethyl sulfoxide (DMSO) to 25 mg of powder. Aliquot out in 20 µL increments and store in a -80 °C freezer for up to six months.
7. Make stopping buffer by adding 2 g of bovine serum albumin (BSA) and 1 vial of aliquoted blebbistatin stock to 100 mL of 1x KHB-HB and sterile filter the solution using a vacuum filtration system.
8. Make plating buffer by adding 5 mL of fetal bovine serum and 1 vial of the aliquoted blebbistatin stock to 95 mL of M199 HEPES. Sterile filter the solution using a vacuum filtration system.
9. Make myocyte culture buffer by adding one 1 vial of the aliquoted blebbistatin stock and 4 mls penicillin-streptomycin to 396 mL of M199 (25 mM HEPES). Sterile filter the solution using a vacuum filtration system.
10. Autoclave 2 pairs of Dumont tweezers, 2 pairs of Iris curved scissors, 2 hemostats, one pair of plastic surgery forceps, 6 black braided silk 4-0 sutures arranged to be used as a surgical double-throw knot, and four 100 mL beakers.
11. Sterilize 22 x 22 mm² glass coverslips. First, place a single coverslip in each well of a six well plate. Afterwards, with the lid removed, turn on the UV lamp of the biosafety cabinet and expose the coverslips to UV light for 1 h.
12. Make working laminin stock solution by first thawing the bottle on ice. Add contents of one bottle to enough cold sterile phosphate-buffered saline (PBS) to reach a final concentration of 0.04 mg/mL. Aliquot out 1.3 µL into autoclaved 1.5 mL centrifuge tubes. Store at -80 °C.
    NOTE: Each tube has enough laminin for a single six well plate. Avoid multiple freeze thaw cycles.
13. Coat sterilized coverslip by first thawing the working laminin solution on ice. Using a P1000 pipette, aspirate 200 µL of laminin. Gently drag the pipette tip along one edge of the coverslip to allow capillary action to pull out a minuscule amount of laminin to facilitate coverslip attachment to the six well plate.
14. Then, expel the remaining laminin in the center of the coverslip. In a circular motion, spread the laminin droplet across the coverslip. Place in a 37 °C incubator at least 1 h and up to 24 h before the isolation.

2. Preparation of the Langendorff Apparatus

NOTE: The individual components of the Langendorff apparatus used in this protocol are listed in Table of Materials.

1. Turn on the circulating water bath. Set temperature so that perfusate has a temperature of 37 °C.
   NOTE: With the solution reservoirs set to a height of 60 cm, the circulating waterbirth needs to be set to 41 °C to have the perfusate be 37 °C. Unlike previously reported protocols, the height of the reservoir does not need to be changed.
2. Rinse the Langendorff apparatus with 70% ethanol followed by two rinses with autoclaved double distilled water. After rinsing, fill reservoir with KHB-HB and oxygenate with 100% oxygen.
3. Prime the system by allowing oxygenated KHB-HB to first flow into a 100 mL beaker. Once 50 mL of solution has flowed into the beaker, switch the 3-way stop-cock position to stop flow from the KHB-HB reservoir. Pour 50 mL of oxygenated KHB-HB from the beaker into the collagenase reservoir.
4. Let the KHB-HB drain from the digestion reservoir until 5 mL remains in collagenase reservoir. While priming collagenase reservoir, switch the 3-way stop cock repeatedly between reservoirs to allow the lines to degas. After the system is primed, remember to use the degassing trap located on top of the heating coil to allow any remaining air to exit the system.
5. Make the collagenase solution. For rats combine 100 mg of type II collagenase, 100 mL of oxygenated KHB-HB, and 2 vials of the blebbistain stock. For mice, combine 100 mg of type II collagenase, 40 mL of oxygenated KHB-HB, and 2 vials of the blebbistain stock. Once mixed, the solution should be stable for 1 h.
   NOTE: Myocyte viability can vary between type II collagenase lots. Take advantage of a collagenase sampling program to test a lot before bulk ordering.

3. Myocyte Isolation

1. Inject the animal with 1,000 U of heparin. Wait 5 min.
   NOTE: Mice and rats of any age can be used. However, in general the older or more diseased the animal, the lower the myocyte yield.
2. Sacrifice the animal by first anesthetizing it with isoflurane using the open-drop method (1 cc of Isoflurane per 500 cc volume) before euthanizing the animal with a pentobarbital mixture (150 mg/kg intraperitoneal).
3. Rapidly excise the heart by first grabbing the fur above the xiphoid process. With the iris scissors, make a small incision immediately below the xiphoid process and pull the fur upwards toward the head exposing the skin.
4. Grab the xiphoid process and cut the diaphragm exposing the thoracic cavity. Make a trap door incision, pull the sternum back using a hemostat, and use the curved forceps to excise the heart above the ascending aorta and place in cold KHB-HB.
5. Cannulate the heart using a stereo microscope and number 5 forceps. Make sure the heart is submerged and the cannula was primed before heart excision to prevent emboli. Confirm proper positioning of the cannula by visualizing the tip of the cannula approximately 1 mm above aortic insertion into the ventricle.
   NOTE: The faster the cannulation time, the better the myocyte yield.
6. Start the flow of KHB-HB by rotating the stopcock on the Langendorff. Connect the cannula to the Langendorff. Perfuse the heart for 5 min.
   NOTE: Since perfusion is supplied by a gravity-based system, flow through the heart will be a function of coronary artery compliance.
7. Switch perfusion from the KHB-HB reservoir to the digestion buffer reservoir. Once the digestion buffer reaches the heart, set a timer (5 min for mouse or 15 min for rat). Make sure to collect the perfusate in a sterile 100 mL beaker. Refill the digestion buffer reservoir as needed with the perfusate until the digestion time has expired.
8. After digestion, separate the chambers of the heart with forceps and the iris scissors in a sterile 100 mL beaker. Place each chamber into a separate well of a six well plate. Pour 5 mL of collagenase solution into each well.
9. Immediately start mincing the heart tissue using scissors. Tissue chunks should be approximately 1 mm³. Using sterile transfer pipettes, gently triturate the minced heart tissue. The solution should turn cloudy.
10. Once the tissue chunks become white and feathery, examine the cells using an inverted microscope. If the number of viable cells is greater than 80%, proceed to strain the cells into a 50 mL conical tube using a 100 µm cell strainer. Use a different tube and strainer for each chamber of the heart.
    1. If the number of viable cells is less than 80%, check the time it took to cannulate. If the cannulation time is over 5 min, try another heart. If not, assay new collagenase lots through the collagenase sampling program.
11. Pellet the cells by centrifuging at 215 x g for 2 min. The pellet should be compact and not loose. If the pellet is loose, the preparation contains many dead cells. In a tissue culture hood, resuspend the pellet in 10 mL of stopping buffer.
12. Pellet the cells by centrifuging at 215 x g for 2 min. The pellet should be compact and not loose. If the pellet is loose, the preparation contains many dead cells.
13. Resuspend the cells in 5 mL of plating buffer. Perform a cell count. Adjust the milliliters of plating buffer to reach a final myocyte concentration of 2 x 10⁴ cells per mL.
14. Remove the laminin-coated coverslips from the incubator. Aspirate the laminin droplet.
15. Plate 200 µL of myocyte suspension on each coverslip. Place in a 37 °C incubator (21% O₂, 5% CO₂) for 2 h to allow attachment. After 2 h, aspirate the unattached cells, add 2 mL of culture media, and culture for up to 4 days.

4. Fura-2 Dye Loading

1. Make a 2 mM fura-2 acetoxymethyl ester (fura-2 AM) stock solution by adding 25 µL of DMSO to 50 µg of fura-2 AM powder (1 vial). Aliquot out into 6 µL aliquots. Take 1 aliquot of fura-2 AM and add to 6 mL of plating medium. Vortex to mix.
2. Remove 1 six well plate of myocytes from the incubator. Aspirate media. Add 1 mL of fura-2 media mixture to each well. Cover plate with foil, leave the plate at room temperature, and wait 15 min.
3. Aspirate fura-2 media mixture and add 1 mL of Tyrode's solution to each well. Cover with foil. Wait 20 min at room temperature to allow for dye washout before imaging.

5. Fluo-4 Dye Loading

1. Make a 1.82 mM fluo-4 acetoxymethyl ester (fluo-4 AM) stock solution by adding 25 µL of DMSO to 50 µg of fluo-4 AM powder (1 vial). Aliquot out into 8.333 µL aliquots. Take 1 aliquot of fluo-4 AM stock and add to 6 mL of plating medium. Vortex to mix.
2. Remove 1 six well plate of myocytes from the incubator. Aspirate media. Add 1 mL of fluo-4 AM media mixture to each well. Cover plate with foil, leave the plate at room temperature, and wait 15 min.
3. Aspirate fluo-4 AM media mixture and add 1 mL of Tyrode's solution to each well. Cover with foil. Wait 20 min at room temperature to allow for dye washout before imaging.

6. Membrane Potential Dye Loading

1. Remove component A and component B from the membrane potential kit. In a 15 mL conical tube, combine 50 µL of component B and 5 µL of component A. Vortex to mix. Add 10 mL of plating media to the 15 mL conical tube containing the voltage dye mixture. Vortex to mix.
2. Remove 1 six well plate of myocytes from the incubator. Aspirate the media. Add 800 µL of the membrane potential dye mixture to each well. Cover plate with foil, leave the plate at room temperature, and wait 15 min.
3. Aspirate dye media mixture and add 1 mL of modified-Tyrode's solution to each well. Cover with foil.

7. Photometry and Charge Coupled Device Recordings

1. Turn on the equipment in the following order: microscope, arc lamp, hyperswitch, fluorescence interface system, Myocam power supply, field stimulator, and computer.
2. Make sure the excitation/emission filter sets are appropriate for the imaging dye.
   NOTE: Fura-2 is excited at 340 nm and 380 nm of light. It emits at 510 nm of light. Fluo-4 and the voltage membrane dye are excited at 485 nm of light and emit at 520 nm of light.
3. Prime the system by turning on the vacuum, fully opening the hose clamp, and gently plunging each 60 mL syringe being used in the manifold. For calcium recordings use Tyrode's solution. For voltage recordings use modified Tyrode's solution.
4. Turn heater on and set flow by adjusting the roller clamp on the perfusion tubing. Make recordings at 36 ± 1 °C.
5. Open the acquisition software. Make sure the parameters are set for the correct imaging dye.
6. In the dark, remove the foil from the six well plate and place a coverslip in the pacing chamber. Make sure the stimulator is off during this step. Focus on the myocytes using the 10x objective.
7. Once in focus, start pacing by field stimulating at 1 Hz, 0.2 V. Gradually increase the voltage until 1:1 pacing is obtained. Then increase the voltage until 1.5x the threshold is reached.
   NOTE: Because excitation-contraction coupling is temperature dependent, make sure the cells have been perfused for 15 min before recording. This allows for myocytes to recover from the shock of going from room temperature back to 37 °C as well as loosely attached cells to float away.
8. Switch from the 10x objective to the 40x objective. Focus in on a cell that is following a 1:1 pacing. Adjust the plastic shades so only one cell is in the field of view.
9. Using the software, place the area of interest box on well-defined sarcomeres. Start the acquisition software to initiate the excitation light. Using the neutral density filters, adjust the intensity setting accordingly to obtain an adequate SNR.