Optical Imaging of Isolated Murine Ventricular Myocytes

Heart failure is the leading cause of death for both men and women in the world. New evidence suggest alteration in cardiac and systemic metabolism contribute to the pathology of heart failure. Rapidly identify which metabolites affect excitation-contraction coupling is still a challenge.

The traditional approach of isolating cardiac myocyte from adult mouse is a time-consuming and challenging process. The innovative method in this article makes a stringent process more efficient and easier to perform. Further, using fluorescent probe, we can perform many phenotypic assessments of how chemicals can induce cardiac toxicity or dysfunction at the cellular level.

By identifying which molecules alter myocyte function, new therapeutics can be identified for the treatment of heart failure. Using transgenic mice, we can probe different genes to determine which can prevent and contribute to the heart failure. This information will be essential in the future curing of heart failure.

Demonstrating the procedure will be Matthew Klos and a student Shreyas Suresh. To begin, thaw the working laminin solution on ice. Using a P1000 pipette, aspirate 200 microliters of laminin and gently drag the pipette tip along one edge of the sterilized coverslip to allow capillary action to pull out a minuscule amount of laminin for facilitating coverslip attachment to the six-well plate.

Then expel the remaining laminin in the center of the coverslip in a circular motion. Spread the laminin droplet across the coverslip. Place the coverslips in a 37 degree Celsius incubator for at least one hour and up to 24 hours before the isolation.

To isolate myocytes from a prepared heart in cold KHB-HB, place the sample under a stereo microscope. To prevent emboli, prime the aortic cannula by submerging it in the KHB-HB solution and using a 20 milliliter syringe to force the solution through. Make sure the heart is submerged and the cannula is primed before heart excision.

With a 5 forceps, cannulate the heart. Confirm proper positioning of the cannula by visualizing the tip of the cannula approximately one millimeter above aortic insertion into the ventricle. Then start the flow of KHB-HB by rotating the stopcock on the Langendorf.

Connect the cannula to the Langendorf to perfuse the heart for five minutes. Next, rotate the stopcock to switch the perfusion from the KHB-HB reservoir to the digestion buffer reservoir. Once the digestion buffer reaches the heart, set a timer.

Collect the perfusate in a sterile 100 milliliter beaker. Refill the digestion buffer reservoir as needed with the perfusate until the digestion time has expired. After digestion, in a sterile 100 milliliter beaker, separate the chambers of the heart with forceps and Iris scissors.

Place each chamber into a separate well of a six-well plate. Pour five milliliters of collagenase solution into each well. Immediately use scissors to mince the heart tissue into chunks of approximately one cubic meter.

Using a sterile transfer pipette, gently triturate the minced heart tissue with digestion buffer. Once the tissue chunks become white and feathery, place the plate under an inverted microscope to examine the cells. If the number of viable cells is greater than 80%proceed to strain the cells into a 50 milliliter conical tube using a 100 micron cell strainer.

If the number of viable cells is less than 80%check the time it took to cannulate. If the cannulation time is over five minutes, try another heart. If not, assay new collagenase slots through the collagenase sampling program.

Next, pellet the cells by centrifuging at 215 times g for two minutes. 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 compact pellet in 10 milliliters of stopping buffer. Pellet the cells again by centrifuging at 215 times g for two minutes. Resuspend the cells in five milliliters of plating buffer.

Perform a cell count on a cyto counter and adjust the volume of plating buffer to reach a final myocyte concentration of two times 10 to the four cells per milliliter. Now remove the laminin-coated coverslips from the incubator. Aspirate the laminin droplet if present.

Plate 200 microliters of myocyte suspension on each coverslip. Place the coverslips in a 37 degree Celsius incubator with 21%oxygen and 5%carbon dioxide for two hours to allow attachment. After two hours, take out the coverslips and aspirate the unattached cells.

Add two milliliters of culture media and culture in the incubator at 37 degrees Celsius for up to four days. First, remove component A and component B from the membrane potential kit. In a 15 milliliter conical tube, combine 50 microliters of component B and five microliters of component A to form a voltage dye mixture.

Vortex to mix. Add 10 milliliters of plating media to the 15 milliliter conical tube containing the voltage dye mixture to form the membrane potential dye mixture. Again, vortex to mix.

Then remove one six-well plate of myocytes from the incubator. Aspirate the media. Add 800 microliters of the membrane potential dye mixture to each well.

Cover the plate with foil and leave the plate at room temperature for 15 minutes. After that, aspirate the dye media mixture from each well and add one milliliter of modified Tyrode's solution to each well. Cover with foil.

Turn on the equipment in the order of microscope, arc lamp, HyperSwitch, fluorescence interface system, MyoCam power supply, field stimulator, and computer. Make sure the excitation emission filter sets are appropriate for the imaging dye. For example, fura-2 is excited at 340 nanometers and 380 nanometers of light.

It emits at 510 nanometers of light. Fluo-4 and the voltage membrane dye are excited at 485 nanometers of light and emit at 520 nanometers of light. Prime the recording system by turning on the vacuum, fully opening the hose clamp, and gently plunging each 60 milliliter syringe with Tyrode's solution in the manifold.

For calcium recordings, use standard Tyrode's solution. For voltage recordings, use modified Tyrode's solution. Turn on the in line heater.

Maintain the temperature so the perfusate in the chamber is at 36 plus or minus one degree Celsius for at least 15 minutes. Next, open the acquisition software. Make sure the parameters are set for the correct imaging dye.

Turn the stimulator off. In the dark, remove the foil from the six-well plate and place a coverslip in the pacing chamber. Place the plate under a microscope and focus on the myocytes using the 10X objective.

Once in focus, start pacing by field stimulating at one hertz and 0.2 volts. Gradually increase the voltage until a one-to-one pacing is obtained. Then increase the voltage until 1.5 times the threshold is reached.

Switch from the 10X objective to the 40X objective. Focus in on a cell that is following a one-to-one pacing. Adjust the plastic shades so only one cell is in the field of view.

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 signal-to-noise ratio.

Here, the representative calcium and sarcomere shortening traces recorded from C57BL/6 mouse myocytes using fura-2 are shown. A quantification of ensemble averaged data obtained from C57BL/6 mice and their transgenic litter mates shows no difference between the groups except for the relaxation time at 10 hertz pacing. The voltage tracing recorded from a C57BL/6 mouse myocyte paced at 10 hertz was post processed to obtain a usable signal.

The ensemble averaged action potential after a low pass Butterworth or a Savitzky-Golay digital filter was applied show subtle differences in the shape of the action potentials. The traces recorded from rat myocytes paced at one hertz shows in addition to the voltage signal being lower than the calcium signal, the contraction kinetics are different as well. This is because calcium dyes buffer calcium while voltage dyes do not.

As with the calcium transient, myocytes demonstrated pacing dependent changes in their optical action potential duration. Drug-induced prolongation of the action potential was demonstrated as well. The last 11 seconds of a 20-second recording indicated that prolonged exposure of myocytes to blue light leads to triggered activity.

When attempting this procedure, the most important thing to remember is to use the lowest intensity light possible so you don't damage the myocytes. Myocyte isolated using our approach can also be used for immunohistochemistry or conducted for Western blot analysis. The beauty of this technique, it allows researchers to rapidly assess how the variety of molecules and genes help excitation-contraction coupling using a single system.