Isolation of High Quality Murine Atrial and Ventricular Myocytes for Simultaneous Measurements of Ca²⁺ Transients and L-Type Calcium Current

Cardiac arrhythmias are common and one of the current major healthcare challenges since they affect millions of people worldwide. Arrhythmias are associated with high morbidity and mortality^1,2 and represent the underlying cause for the majority of sudden cardiac deaths³. Up to date treatment options have improved patient survival but are still mainly symptomatic treatments rather than targeting the underlying mechanisms. Thus, these treatments have limited efficacy and may frequently cause severe side effects^4,5,6. An improvement of current treatment options requires insight into the underlying pathophysiology, creating the need for suitable models to study. Small animal models - and specifically mouse models - play a crucial role in arrhythmia research as they allow to study key mechanisms of arrhythmogenesis, for example the genetic impact on cellular electrophysiology, ion channel function or calcium handling^7,8.

For this purpose, isolated atrial and ventricular cardiomyocytes of sufficient quantity and viability are required. A broad spectrum of different isolation approaches to obtain atrial and ventricular myocytes has been previously described^{9,10,11,12,13} and some groups have presented data from simultaneous measurements of L-type current and calcium current induced calcium transients from either atrial¹⁴ or ventricular¹⁵ murine cardiomyocytes. However, to our best knowledge there is no data available of atrial as well as ventricular measurements from one animal. Researchers focus on a broad variety of topics ranging from electrophysiology to proteomics, functional studies as cell contractility or protein interactions, mitochondrial function, or genetics – all in need of isolated cardiomyocytes. Many of the published protocols thus have not been specifically developed for patch clamp studies, leading to limited yields and insufficient cell quality for patch clamp studies. Thus, simultaneous patch clamp and calcium transient measurements of atrial and ventricular cells isolated from one animal cannot be performed with established protocols.

Isolation of murine – especially atrial – myocytes for patch clamp experiments remains challenging. This article provides a simple and fast method for the isolation of high-quality murine atrial and ventricular myocytes via retrograde enzyme based Langendorff perfusion, which subsequently allows simultaneous measurements of both net membrane current and current induced calcium transients from one animal. This article elaborates a protocol for the isolation of atrial and ventricular myocytes derived from wild type mice and mice carrying genetic mutations. This protocol can be used for male and female mice alike. The myocyte isolation, images, and representative results described below were obtained from wild type C57Bl/6 mice at the age of 6 (± 1) months. Nevertheless, this protocol has successfully been used for mice at various ages ranging from 2 to 24 months with different genotypes. Figure 1 shows the isolation setup and a close-up of a cannulated heart during enzyme perfusion.

All animal procedures were approved by the Lower Saxony Animal Review Board (LAVES, AZ-18/2900) and were conducted in accordance with all institutional, national, and international guidelines for animal welfare.

1. Prearrangements

1. Prepare 1 L of 10x perfusion buffer (Table 1), 500 mL of 1x perfusion buffer (Table 2), 50 mL of digestion buffer (Table 3), 10 mL of stop buffer (Table 4), 1 L of Tyrode solution (Table 5), 10 mL of each calcium step solution (Tyrode solution with glucose and respective amount of calcium as indicated), 1 L of 4-AP solution (Table 5), 100 mL of pipette solution (Table 6) and 5 mL of pluronic acid according to the provided recipes.
   NOTE: Bath solutions (Tyrode and 4-AP solution) can be prepared (without glucose) in advance and stored at +4 °C, glucose is added on the experimental day. Pipette solution can be stored at -20 °C, calcium indicator is added on the experimental day and solution then stored on ice until further use. 10x perfusion buffer can be stored at room temperature, 1x perfusion buffer, digestion buffer and stopping buffer should be freshly prepared on experimental day.
2. Turn on the water bath and roller pump.
3. Prefill Langendorff apparatus with perfusion buffer; make sure it is air-free.
4. Prepare aortic cannula by fixing it under the dissection microscope, connect with a 1 mL syringe filled with perfusion buffer and clear air by rinsing the cannula.
   NOTE: It is crucial to avoid any air inside the perfusion system, as this will directly affect coronary perfusion and thus digestion effectiveness. A bubble trap might be added to the setup if necessary, to safely avoid any air trapping.
5. Prepare Petri dishes with enough perfusion buffer for organ collection and microscopy (buffer should securely cover the entire organ, a few millilitres – depending on the used Petri dish size – should be enough).
6. Prepare 3 Petri dishes with digestion buffer for tissue dissociation and microscopy, buffer should cover the organ for dissection under the microscope within the respective Petri dish, amount depends on the used Petri dish size. For the dissociation use 3 mL for ventricular tissue and 1.5 mL for atrial tissue.

2. Organ harvest

1. Inject mouse with 0.1 mL of heparin (1,000 U/mL) i.p. using a 1 mL syringe with a 27 G cannula and wait for 5-10 min.
2. Place the mouse into an induction chamber along with a small tissue soaked in approximately 500 µL of isoflurane. The animal should not be in contact with the tissue. To avoid that, one can use a plastic biopsy-embedding cassette to cover the tissue. Once the animal is fully anesthetized, check for toe pinch reflex and as soon as it is not present anymore, quickly euthanize the mouse by cervical dislocation.
3. Place the mouse on a platform on its back (e.g., on Styrofoam covered with a paper towel) and fix the paws down with cannulas to hold it in place.
4. Remove fur and skin covering the chest and part of the abdomen with a clear cut from jugulum towards symphysis and open the abdomen right under the xiphoid without injuring any organ structure using scissors. Lift the sternum with surgical forceps and cut the diaphragm with scissors along the edge of the ribs, then cut the ribs in medial axillary line and remove the rib cage to expose the heart.
5. Carefully remove the pericardium using blunt forceps and quickly remove the heart by lifting it with blunt forceps from below and by cutting the large vessels with one single cut using scissors.
6. Put the heart into room temperature perfusion buffer and cannulate the aorta with a blunt end needle under the microscope as quickly as possible.
   NOTE: Remove any lung tissue and fatty tissue attached to the organ without losing too much time on it. While cannulating, make sure that the end of the needle does not extend through the aortic valve, as this will impair results by preventing buffers from entering the coronary arteries.
7. Tie the heart with a piece of suturing silk firmly to the needle and disconnect from syringe.
   NOTE: The entire procedure from obtaining the heart (the moment when the large vessels are cut) until suturing the aorta to the needle should take as little time as possible. It is recommended not taking longer than 90-180 s from removing the heart until start of perfusion.

3. Enzymatic digestion

1. After aortic cannulation, immediately connect the cannulated heart to the Langendorff apparatus avoiding any air entering the system.
   NOTE: It can help to have a hanging drop of perfusion buffer at the bottom of the Langendorff apparatus as well as a drop of perfusion buffer sitting on the top of the needle in order to avoid any air entering the system.
2. Perfuse the heart with perfusion buffer for 1 min at a temperature of exactly 37 °C and a perfusion rate of exactly 4 mL/min.
   NOTE: In order to have a temperature of 37 °C at the tip of the perfusion needle, water bath temperature has to be set slightly above at approximately 40 °C. This should be tested regularly by measuring the temperature at the perfusion tip.
3. Switch perfusion to digestion buffer and perfuse for exactly 9 min at a temperature of exactly 37 °C and a perfusion rate of exactly 4 mL/min.
4. Transfer the digested heart to a Petri dish with enough digestion buffer to keep it fully covered. Then carefully dissect the atria and ventricles under the microscope.
5. Transfer the atria into a Petri dish with 1.5 mL of digestion buffer and the ventricles into another Petri-dish with 3 mL of digestion buffer.
6. Atrial dissection
   1. Carefully, but without loss of time, pull the atria apart into tiny pieces using blunt forceps.
   2. Dissolve the tissue by carefully pipetting up and down using a 1,000 µL pipette tip, which has previously been cut to widen the tip opening.
   3. Transfer the solution to a 15 mL centrifuge tube and add an equivalent amount of stop buffer (1.5 mL) by carefully pipetting down at the side of the tube to end the reaction.
   4. Carefully pass all 3 mL of cell/tissue-solutions through a 200 µm nylon mesh to remove remaining larger tissue pieces that have not been fully digested.
      NOTE: A successful digestion will leave almost no solid chunks.
7. Ventricular dissection
   1. Quickly chop the ventricular tissue into tiny pieces using dissection scissors and pipette up and down to dissolve. Use another 1,000 µL pipette tip for pipetting up and down, it may be shortened to widen the opening.
   2. Transfer cell/tissue-solution into a 15 mL centrifuge tube and add an equivalent amount of stop buffer (3 mL) by carefully pipetting down at the side of the tube to end the reaction.
   3. Carefully pass all 6 mL of cell/tissue-solution through a 200 µm nylon mesh to remove larger tissue pieces that have not been fully digested.
      NOTE: A successful digestion will leave almost no solid chunks.
8. Leave both tubes (atrial and ventricular cell suspension) on the bench at room temperature for 6 min to settle.
9. Centrifuge both 15 mL tubes at 5 x g for 2 min.

4. Calcium reintroduction

NOTE: The following steps are identical for both atrial and ventricular cells (unless otherwise mentioned) and are performed at room temperature.

1. Discard the supernatant using a plastic Pasteur pipette and carefully resuspend pellet in 10 mL of calcium free Tyrode solution.
2. Leave cells for 8 min for sedimentation.
3. Centrifuge at 5 x g for 1 min (only atrial cells, sedimentation is enough for ventricular cells).
4. Discard supernatant and carefully resuspend pellet in 10 mL of Tyrode solution with 100 µM calcium concentration.
5. Leave cells for 8 min for sedimentation.
6. Centrifuge at 5 x g for 1 min (only atrial cells, sedimentation is enough for ventricular cells).
7. Discard the supernatant and carefully resuspend pellet in 10 mL of Tyrode solution with 400 µM calcium concentration.
8. Leave cells for 8 min for sedimentation.
9. Centrifuge at 5 x g for 1 min (only atrial cells, sedimentation is enough for ventricular cells).
10. Discard the supernatant and carefully resuspend the pellet in 1 mL (atrial)/ 5 mL (ventricular) of Tyrode solution with 1 mM calcium concentration.

5. Loading of myocytes with fluorescent calcium-indicator Fluo-3 AM

NOTE: Due to the light sensitivity of the fluorescent calcium indicator, the following steps should be executed protected from light (e.g., by covering tubes with aluminium foil).

1. Prepare Fluo-3 AM stock solution by adding 44 µL of 20% pluronic F-127 in anhydrous DMSO to 50 µg of Fluo-3AM (can be stored at -20 °C protected from light).
2. Add 10 µL of Fluo-3 AM stock solution to 1 mL of cell suspension and incubate for 10 min at room temperature protected from light.
3. Centrifuge at 5 x g for 1 min.
4. Discard the supernatant using a plastic Pasteur pipette and resuspend the pellet in a reasonable amount of bath solution to obtain a good working concentration (1-5 mL of bath solution depending on the cell density).
5. Leave for 30 min for de-esterification before starting with experiments.

6. Simultaneous patch-clamp and epifluorescent Ca²⁺-transient measurements as previously described¹⁶

NOTE: Patch clamp measurements are not the topic of this article, the interested reader may be referred to major publications providing in depth descriptions of this method^{17,18,19,20,21,22}. Nevertheless, for a better overall understanding, a summary on a protocol to measure L-type calcium currents along with current induced calcium transients is provided.

1. Transfer myocytes into a cell chamber and superfuse with bath solution at 37 °C.
2. Block potassium currents by adding 4-aminopyridine and barium chloride to the bath solution as indicated in Table 5.
3. Ensure that borosilicate microelectrodes have a tip resistance of 2-5 MΩ filled with pipette solution (Table 6).
4. Setup measurements to allow for simultaneous recording of both electrical signals and epifluorescence at the same time. Voltage clamp mode is used to measure L-type Ca²⁺-current with a protocol holding the cell at -80 mV and a 600 ms ramp-pulse to -40 mV to inactivate the fast Na⁺-current, followed by a 100 ms test-pulse to +10 mV at 0.5 Hz (Figure 2).
5. Use excitation at 488 nm, emitted light at <520 nm to detected and convert to [Ca²⁺]_I assuming
   
   Where k_d = dissociation constant of Fluo-3 (864 nM), F = Fluo-3 fluorescence; F_max = Ca²⁺-saturated fluorescence obtained at the end of each experiment¹⁹.

Isolation yield is determined after calcium reintroduction by pipetting 10 µL of cell suspension onto a microscope slide. More than 100 viable, rod-shaped, non-contracting cells/10 µL for atrial cell isolation and more than 1,000 viable, rod-shaped, non-contracting cells/10 µL for ventricular cell isolation are considered as sufficient yield and are commonly obtained using this protocol. Atrial cells obtained with this protocol showed a variety of different cell types containing cells of the cardiac conduction system, especially the sinus node, as well as different myocytes from the right and left atrium as well as the atrial appendages. Since these regions are not dissected separately, various cell morphologies are the result. All atrial cells are small, with cell capacitances ranging from approximately 35 pF– 100 pF, some are more filiform than others. Figure 3 panel A shows a typical cell from the atrial working myocardium loaded with calcium sensitive dye, ready for measurements obtained with this protocol. Ventricular myocytes are more rod-shaped and larger with cell capacitances ranging from 100 to around 400 pF, Figure 3 panel B shows a typical ventricular cell from the working myocardium loaded with calcium sensitive dye, ready for measurements obtained with this protocol. Figure 4 shows representative examples of L-type calcium current measurements with simultaneous cytosolic calcium transients from one atrial myocyte (panel B and C) and one ventricular myocyte (panel E and F).

Figure 1: Langendorff-apparatus. (A) Photograph of the isolation setup with view of the custom designed heat exchanger and jacketed heart chamber. (B) Close-up of jacketed heart chamber with cannulated heart in place. Please click here to view a larger version of this figure.

Figure 2: Stimulation protocol. Voltage-clamp protocol (0.5 Hz) used to measure total net membrane current (I_M), predominantly reflecting L-type Ca²⁺ current and Ca²⁺ current induced Ca²⁺ transient. Please click here to view a larger version of this figure.

Figure 3: Isolated murine cardiomyocytes. (A) An example of an atrial cardiomyocyte loaded with Fluo-3, ready for patch-clamp experiments. (B) An example of a ventricular cardiomyocyte loaded with Fluo-3, ready for patch-clamp experiments. Please click here to view a larger version of this figure.

Figure 4: Representative recordings. (A) Voltage-clamp protocol (0.5 Hz). (B) Total net membrane current (I_M), predominantly reflecting L-type Ca²⁺ current in a murine atrial myocyte. (C) L-type Ca²⁺ current triggered Ca²⁺ transient in a murine atrial myocyte. (D) voltage-clamp protocol (0.5 Hz). (E) Total net membrane current (I_M), predominantly reflecting L-type Ca²⁺ current in a murine ventricular myocyte. (F) L-type Ca²⁺ current triggered Ca²⁺ transient in a murine ventricular myocyte. Please click here to view a larger version of this figure.

Table 1: 10x perfusion buffer (1,000 mL).

Table 2: 1x perfusion buffer (500 mL).

Table 3: Digestion buffer (50 mL).

Table 4: Stop buffer (10 mL).

Table 5: Bath solutions.

Table 6: Pipette solution.

Table 7: Common problems and how to solve them.

This article provides an easy and functional way to obtain high quality atrial and ventricular myocytes from the same mouse for patch-clamp studies with simultaneous calcium transient recordings. The quality of the obtained data highly depends on the quality of the cell isolation. As mentioned above, many methods to isolate murine cardiomyocytes have been described previously^9,10,11,12. The isolated cells are used for various analyses ranging from patch clamp experiments to contractility studies, morphological studies, proteomics, mitochondrial function research and many more. Each individual purpose requires isolated cells optimized differently. Due to this, none of the protocols led to high quality isolations suitable for patch-clamp experiments with simultaneous measurement of calcium handling properties of atrial and ventricular myocytes from one mouse. To change this, the adaption of numerous protocols described before resulted in the development of this protocol. Some steps are mechanically challenging, some are to be done in a specific fashion and some are simply crucial to isolation quality. These steps will be discussed below.

Yield and quality of cell isolation decrease with animal age, possibly due to increased tissue fibrosis^11,23,24 making individual adjustments of digestion time and enzyme amount necessary. The older an animal, the more fibrotic tissue is to be expected. The indicated digestion times and enzyme amounts are a good starting point for normal sized, healthy mice at the age of 6 months, but will need adaption to individual needs. It is suggested to apply more enzyme and longer digestion times when using older mice or transgenic mice favouring fibrosis. As previous work has emphasized, rapid and air-free cannulation as well as immediate perfusion of the heart are absolutely critical for good cell quality¹¹. It is recommended not taking longer than 90-180 seconds from obtaining the heart and putting it into room temperature perfusion buffer to start of perfusion – the faster, the better. It is helpful to set up all the needed equipment in one room to avoid long distances. If further shortening of the ischemic interval is desired, it is possible to use another method of narcosis and subsequent euthanization. Fully anesthetize the animal as described above, apply a sufficient amount of fentanyl (or an equivalent analgesic medication in accordance to your guidelines for animal welfare) i.p. and move the animal to a platform with an isoflurane vaporizer and an anaesthesia mask suitable for mice, providing constant isoflurane and oxygen flow. Afterwards proceed as described above. Since the animal has a normal circulation until the large vessels are disconnected with a single cut, this method will reduce the no-flow time significantly and can thus improve isolation quality if needed.

After harvesting the heart, it is critical to place the aortic cannula at the right depth to secure coronary perfusion. It is, therefore, necessary to cut the aorta at the aortic arch or at the descendent aorta when removing the heart to leave at least 5 mm of the aorta for cannulation allowing to tie the suturing silk distal from the coronary arteries but before the first aortic side-branches.

During the isolation process, multiple cell transfers are necessary and pipetting with small pipette tips will cause high shear stress to the cells. Possible solutions to reduce shear stress are pipetting slowly and carefully, as well as cutting pipette tips by some millimetres in a 45° angle to widen the tip opening. Further reduction of shear stress can be achieved by melting the cut pipette tips to soften the edges. Isolation yield of the ventricular isolation usually is so high that shear stress reduction is not obligatory. Depending on one’s individual goal, it can even be more useful to expose ventricular cells to shear stress to select for the ’fittest’ cells. However, for the atrial isolation this procedure is critical, as atrial cells are particularly vulnerable after digestion. Whenever adding solutions, it is recommended to avoid pipetting directly onto the cells, but rather trying to pipet slowly at the tube wall.

The choice of collagenase will significantly affect isolation results. Since there are not only differences in enzyme preparations but also a relevant batch-to-batch variation in enzyme activity, titration of collagenase amount and digestion time are necessary when starting to work with a new batch of enzyme or a new strain of mice. However, the amount and time indicated in the tables above mark a good starting point for individual optimization⁹. Most companies provide small enzyme samples to test individual batches, which makes batch selection much more convenient.

Calcium transients with typical amplitudes and normal monophasic decay can be obtained if cells are isolated without the use of EGTA²⁵ as previously described by Voigt et al.^9,16. This protocol, therefore, uses low calcium concentrations and avoids EGTA during isolation process. To protect cells from the Ca²⁺ paradox phenomenon²⁶, calcium is reintroduced stepwise and slowly until a final concentration of 1 mM. Due to higher intracellular sodium concentrations, rodent cardiomyocytes are especially prone to calcium overload and slow and stepwise reintroduction is crucial²⁷. The need of calcium free solutions represents one of the major limitations of all Langendorff-based rodent cardiomyocyte isolations. Calcium reintroduction always leads to a significant loss of viable cells. Slow and stepwise reintroduction as described here nevertheless, leads to sufficient yield with good quality to obtain measurements from each animal reliably.

It is important to assess cell yield and quality right after calcium reintroduction. Although final evaluation takes place while patching the isolated cells, one can already judge cell quantity and quality by having a look under the microscope prior to loading cells with calcium sensitive dye. Good quality cells are clearly rod-shaped, have a homogenous membrane and are non-contracting. Contraction is a sign of low cell quality as it indicates a loss of membrane integrity and might be caused by overdigestion or application of high shear stress. Another sign of overdigestion is – besides many cell shadows in relation to viable cells – an overly nice-looking membrane that has been cleared of many surface proteins and is extremely vulnerable when one tries to approach the cell with the pipette tip for seal formation. Exemplary healthy, good quality cells are shown in Figure 3. (Panel A: atrial myocyte, Panel B: ventricular myocyte). Isolations with high yields can lead to cells immune to seal formation and vice versa. In the end, the ultimate quality test regardless of cell looks and yield, is the answer to a simple question: ‘Is it possible to perform the desired measurements with high quality results in the isolated cells of each isolation?’. The protocol provided above makes a positive answer possible.

Many isolation protocols use the uncoupling agent Blebbistatin to obtain higher yields and better-quality cells. This protocol purposely avoids Blebbistatin, taking into consideration the published evidence of its influence on cardiac electrophysiology^28,29 in optical mapping experiments. The use of uncoupling agents in electrophysiologic studies must be treated with caution and should be limited to circumstances where uncoupling is mandatory.

Cells obtained with this protocol can be used for approximately six hours after loading with Fluo-3 with atrial cells being more vulnerable to time. Consequently, it is recommended to study atrial cells before ventricular cells if the same researcher processes them.

One limitation of this protocol – as true for most Langendorff-based isolation protocols – is that it is technically difficult. Clearly, mouse hearts are small, and all technical aspects require a lot of exercise. This means that an experienced researcher will obtain better results. Some protocols use a constant pressure to perfuse the heart. This has the advantage that due to digestion and consecutive loss of tissue resistance the perfusion will accelerate, and this helps to define the optimal digestion time, nevertheless the acceleration can harm the tissue und decrease cell quality significantly. In a constant flow approach as used here, there are no clear benchmarks for digestion times and it can often be hard to stop the digestion at an optimal time, which marks a relevant limitation. Another limitation is that in this protocol isolated cells were only tested for suitability to the experiments described above. It is likely that these cells can be used for different analysis as well, but this still must be proven. A first step into that direction was usage of this cells to visualize action potentials by loading them with VF2.1CI, a recently derived voltage sensitivity dye with superior kinetics to previously used voltage sensitive dyes according to a protocol published by Seibertz et al.³⁰. To help with troubleshooting, Table 7 shows common isolation problems and how to approach them.

Taken together, the described isolation protocol offers a working approach to the simultaneous isolation of murine atrial and ventricular cardiomyocytes, enabling the researcher to obtain an atrial and ventricular electrophysiological phenotype of a single mouse. This removes statistical obstacles posed by previous isolation protocols only isolating either atrial or ventricular cells at high quality and helps to reduce the number of animals needed for a specific study. This can be especially important if interventions as for example the implantation of osmotic minipumps filled with expensive agents are part of the experiments and it can be a vital factor in animal welfare considerations.