Isolation and Kv Channel Recordings in Murine Atrial and Ventricular Cardiomyocytes

The overall goal of this procedure is to extract living atrial and ventricular cardiomyocytes from genetically engineered mice for use in whole cell patch clamp recording and other experiments. This is accomplished by first carefully, but rapidly excising the heart of an anesthetized animal. The second step is to connect the excised and cannulated heart to the LOR apparatus.

Next, the heart is retro greatly perfused with collagenase solution To loosen the connecting tissue, the final step is to dissociate the tissue by gently pulling it apart with fine forceps and to suspend the cells with an enlarged fire polished plastic pipette tip. After that, cells are resuspended stepwise with calcium rich solution. Ultimately, whole cell patch clamp technique is used to show differences in voltage gated potassium channel currents.

The main advantage of this method is that you will receive a good amount of HL and ventricular cardiomyocytes with good characteristics for further experiments. Generally, scientists new to this method will struggle because it needs a lot of practice to precisely perform this procedure and get really good results. I, after injecting the anesthetics and 250 IU heparin, ensure that the mouse is deeply anesthetized by pinching the tail to check the flight reflex.

Next, transfer the mouse to the operating table in supine position. Then make an incision on the skin and the abdominal wall below the xiphoid. To perform clamshell thoracotomy.

Extend the cut to both sides along the costal arch. Subsequently cut the ribs in the medial axillary line and deflect the rib cage upwards. After that, open the pericardium and locate the great vessels.

Gently press the coddle heart to better display the aorta. Then clamp the aorta using a pair of forceps. Lift the heart up carefully, then place a pair of scissors below the heart and dissect all connecting vessels with one single cut.

Make sure to preserve a large enough part of the ascending aorta for Lor cannulation. Next, transfer the excised heart immediately to a Petri dish filled with ice, cold and pre solution.One. In this procedure, cannulate the aorta with a 1.8 French steel cannula and make sure to avoid air embolism.

Next, fix the aorta to the cannula with a surgical suture. After that, flush the coronaries with one milliliter of solution.One. Connect the cannula to a langor apparatus to avoid extended ischemia reperfusion injury to the myocardium.

Make sure to perform the procedures from thoracotomy to l endorf cannulation within 120 seconds. Then after pre-filling the warming coil of the L endorf system with calcium free solution, two. Perfuse the heart with 10 milliliters of this solution at four milliliters per minute.

Subsequently, perfuse the heart with collagenase solution. Three for eight minutes at four milliliters per minute. Now transfer the heart to a prewarm 100 millimeter Petri dish containing low calcium solution.Four.

Carefully remove the aortic and other non-cardiac tissue with scissors. Then separate the atria and the ventricles and keep the atrial and ventricular tissue immersed in small amount of solution.Four. To isolate the atrial cardiomyocytes, transfer the atria into a separate prewarm 100 millimeter culture dish.

Dissociate the tissue by gently pulling it apart with fine forceps and ensure that the tissue is almost completely dissociated. Next, use a one milliliter pipette with an enlarged fire polished plastic pipette tip to suspend the cells in one milliliter of solution. Four for atrial cells and five milliliters of solution.

Four for ventricular cells for five minutes. After that, separate the cells from the debris using a cell filter of 200 micron mesh size. Centrifuge the cells in solution four for one minute for atrial cells, and for two minutes for ventricular cells At room temperature at 16 times G in a cell culture hood, discard the supernatant and resuspend the pellet in five milliliters of solution.

Five for the atrial cells and around 20 to 25 milliliters. For the ventricular cells, gently shake the tube to resolve the cell pellet and centrifuge again for one minute at room temperature at 16 times G after the second centrifugation, discard the supernatant again and resuspend the pellet in five milliliters of solution. Six for the atrial cells and around 20 to 25 milliliters for the ventricular cells.

Then allow the sedimentation of the cells for 10 minutes. In a cell culture hood, remove the supernatant, then count the cells subsequently. For electrophysiology studies add 25 to 50 milliliters of solution six to the cells for electrophysiology studies.

Keep the myocytes in solution six in a 50 milliliter tube and inhibit sedimentation. For immunofluorescent staining. Prepare a cell culture dish with glass cover slips coated with laminate solution at a final concentration of 50 micrograms per milliliter.

Laminin in PBS incubate the cell culture dish for two hours at 37 degrees Celsius. Next, remove the laminate solution. Before plating the myocytes under a microscope, plate the cells and control the cell density.

Let the myocytes adhere to the cover slip for one hour at 37 degrees Celsius in 2%CO2. After an hour, remove the solution and start immediately with the standard staining procedure protocol. Then incubate the myocytes with fixative.

For example, 4%PFA in PBS at pH 7.5 for 10 minutes at room temperature. Then follow with three PBS washing steps for five minutes each to perme the cells and to inhibit unspecific antibody binding. Incubate the myocytes with 10%serum 0.3%Triton 0.2%BSA in PBS for 30 minutes at room temperature.

After that, incubate them with the primary antibody for one hour at 37 degrees Celsius and wash them as described before. Subsequently incubate with the secondary antibody for one hour at room temperature to counterstain the nuclei and alpha actinin. Incubate the secondary antibody with DPI and fluorochrome conjugated phin for one hour at room temperature.

After washing transfer, the glass cover slips carefully to the saline treated microscope Slides. Then embed the cells in the fluorescence mounting medium for whole cell patch clamp recordings. Transfer the healthy appearing cardiomyocytes to the perfusion chamber filled with a defined volume of extracellular bath solution.

Use the proper patch clamp equipment and patch pipettes with resistances of three to five mega ohms when filled with intracellular solution For standard patch clamp recordings, after a cell has been patched and a giga seal has been established, use a standard protocol to measure the potassium currents by evoking the outward potassium currents with 4.5 second voltage. Steps from minus 60 millivolts to 50 millivolts in 10 millivolt increments. Make sure leak currents are always smaller than 100 pico amps.

To differentiate different potassium currents, use specific inhibitors such as four amino paridine, hetero potto, toin, or tetraethyl ammonium. Stock solutions should be prepared in extracellular bath solution and applied directly to the closest possible vicinity of the cell via a micro tip. After baseline recordings.

An equilibration for two to three minutes should be allowed before starting the recordings after drug application At the end, analyze the data offline using P clamp 10.3 software or comparable software. This figure shows an exemplary ventricular cardiomyocyte stained with DPI and LOR 4 88 Phin. And here are the exemplary traces of the whole cell patch clamp recording.

This figure shows an exemplary atrial cardiomyocyte stained with DPI and fluorochrome conjugated phin. And here are the exemplary traces of the whole cell patch clamp recording Once mastered, this technique can be done in more or less one and a half hour if it's performed properly. After watching this video, you should have a good understanding of how to isolate atrial and ventricular cardiomyocytes of genetically engineered mice.