Isolation of Human Atrial Myocytes for Simultaneous Measurements of Ca²⁺ Transients and Membrane Currents

Experimental protocols need to be approved by the local ethics committee and all patients need to give written informed consent. Our research was approved by the ethics committee of the Medical Faculty Mannheim, University of Heidelberg (#2011-216N-MA) and was performed in compliance with all institutional, national and international guidelines for human welfare. All patients gave written informed consent.

0. Obtaining Human Atrial Tissue

During routine cannulation procedures in patients undergoing open-heart surgery for cardiopulmonary bypass grafting, the tip of the right atrial appendage is usually removed and can be used for isolation of atrial cardiomyocytes. After excision the tissue sample is transferred immediately into a 50 ml Falcon tube with sterile Ca²⁺-free transport solution containing 2,3-butanedione monoxime (Table I; BDM, contractile inhibitor, preventing myocyte contracture). With transport times between 30 and 45 min, we did not recognize a clear advantage of cooling or oxygenation with respect to both number and quality of isolated atrial cardiomyocytes. In general transport to the lab should be as quick as possible. However, if longer transportation time cannot be avoided, transport at 4 °C in oxygenated solution might be advantageous.

1. Prearrangements

1. Switch on the thermocirculator, which controls the temperature of the jacketed beaker (Table II). Make sure the temperature within the beaker equals 37 °C. Cover the beaker with a glass lid to maintain a constant temperature within the beaker throughout the whole isolation process.
2. Weigh the Collagenase I and Protease XXIV into three glass beakers and store it at room temperature. The enzymes will be diluted just before use.
3. Store 20 ml of Ca²⁺ free solution in a Petri dish at 4 °C.
4. Store 250 ml of Ca²⁺ free solution in a water bath at 37 °C.

2. Cleaning of the Tissue

1. Transfer the tissue sample together with the transport solution into a Petri dish (~10 cm diameter) and remove fatty tissue roughly using scissors.
2. Weigh the tissue sample. Between 200 and 600 mg are used for cell isolation. The remaining tissue can be frozen in liquid nitrogen for later biochemical analysis.
3. Transfer the tissue sample into the Petri dish containing 20 ml Ca²⁺ free solution at 4 °C (see step 1.3) and chop the tissue sample into small chunks of approximately 1 mm³ in size.
4. The following steps should be executed at 37 °C and under continuous gassing with 100% O₂. A pipette tip can be placed onto the oxygen tube to avoid strong bubbling and generate continuous air flow.
5. Transfer the tissue chunks together with the Ca²⁺-free solution into the jacketed beaker and stir carefully for about 3 min with a magnetic stirring bar.
6. Stop stirring and allow the tissue chunks to settle down for a few seconds. Carefully strain the supernatant through a nylon mesh (200 μm, Table II). Return the restrained tissue chunks from the mesh into the beaker using forceps.
7. Refill the beaker with Ca²⁺ free solution and repeat the washing step 2.4 and 2.5 two times.

3. First Enzymatic Digestion

1. Re-suspend the tissue chunks with 20 ml Enzyme solution E1 (Table I) containing collagenase and protease and stir carefully for 10 min.
2. Add 40 μl of 10 mM CaCl₂-solution to obtain a final concentration of 20 μM Ca²⁺.
3. After 35 min strain the supernatant carefully through a nylon mesh (200 μm, Table II) in a way that most of the tissue chunks remain in the beaker. Return restrained tissue chunks from the mesh into the beaker.

4. Second Enzymatic Digestion

1. Resuspend the tissue chunks again with 20 ml enzyme solution E2 containing collagenase I only (Table I). Add 40 μl of 10 mM CaCl₂-solution immediately to obtain a final concentration of 20 μM Ca²⁺.
2. During the second digestion use scissors to further chop tissue clots occasionally.
3. After 5 min take a sample using a Pasteur pipette to check the dissociation of cells. Repeat this every 2-3 min until rod-shaped, striated cardiomyocytes appear.
4. Stop stirring and allow the tissue chunks to settle down for about 20-30 sec.
5. Strain the supernatant carefully through a nylon mesh (200 μm, Table II) into a 50 ml Falcon tube (Tube A). Return the restrained tissue chunks from the mesh into the beaker.
6. Re-suspend the tissue chunks in the beaker with 20 ml storage solution (Table I)¹⁰ and further dissociate the cells by gentle mechanical trituration using a 20 ml serological pipette with dispenser. Try to avoid bubble formation since bubbles are detrimental to cell survival and quality.
7. Strain the supernatant carefully through a nylon mesh (200 μm, Table II) into a 50 ml Falcon Tube (Tube B).

5. Final Preparation and Adjustment of Final Ca²⁺ Concentration

1. Centrifuge both Falcon tubes A and B (see step 4.5 and 4.7) at 95 x g for 10 min.
2. Remove the supernatant from both tubes carefully by aspiration. Make sure not to disturb the pellet. Discard the supernatant.
3. Re-suspend both pellets in 1.5 ml storage solution (Table I) each (room temperature).
4. Add two times 7.5 μl of 10 mM CaCl₂ solution to each Falcon and stir carefully. Incubate for 10 min after each step.
5. Add 15 μl of 10 mM CaCl₂ solution to obtain a final Ca²⁺ concentration of 0.2 mM.

6. Loading of Myocytes with the Fluorescent Ca²⁺-indicator Fluo-3 AM (Figure 1)

1. Transfer 1.5 ml of cell suspension (tube A and/or tube B) into a 2 ml microcentrifuge tube (Eppendorf tube).
2. The following steps should be executed under consideration of the light sensitivity of the fluorescent Ca²⁺ Indicator Fluo-3.
3. Dissolve 50 μg of the membrane permeable acetoxymethyl ester derivative of Fluo-3 (Fluo-3 AM, Table III) in 44 μl of the Pluronic F-127 (Table III) stock solution (20% w/v in anhydrous DMSO) to get a 1 mM Fluo-3 AM stock solution, which can be stored at -20 °C for at most 1 week.
4. Add 15 μl of the Fluo-3 AM stock solution to the microcentrifuge tube containing 1.5 ml of cell suspension (see step 6.1) and agitate carefully.
5. Incubate the cell suspension for 10 min in an optically opaque box.
6. Briefly centrifuge at about 6,000 rpm.
7. Discard the supernatant and re-suspend the pellet in 1.5 ml bath solution (Table IV).
8. Leave the cell suspension for about 30 min for de-esterification before beginning with experiments.

7. Simultaneous Patch-clamp and Epifluorescent Ca²⁺ Measurements

Since patch-clamp measurements are not the major topic of this review, we refer the interested reader to other publications providing a more in depth description of this technique.^11-14 For the sake of completeness we provide a brief summary of a protocol to measure action potentials or L-type Ca²⁺ currents, both together with simultaneous Ca²⁺-transient recordings.

During experiments myocytes are superfused at 37 °C with bath solution (Table IV) using a rapid perfusion system (Octaflow IITM, ALA Scientific Instruments, NY). For voltage-clamp experiments, K⁺ currents are blocked by adding 4-aminopyridine (5 mmol/L) and BaCl₂ (0.1 mmol/L) to the bath solution. Borosilicate glass microelectrodes are used and should have tip resistances of 2-5 MΩ when filled with pipette solution (Table V). In addition to the Fluo-3 AM loading of the myocytes (see step 6), Fluo-3 is also included in the pipette solution (Table V). Fluorescence is excited at 488 nm and emitted light (<520 nm) converted 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 experiment.¹² Both electrical signals and epifluorescent Ca²⁺ signals are recorded simultaneously. Action potentials are stimulated at 0.5 Hz in current-clamp mode using 1 msec current pulses of 1.2x threshold strength. L-type Ca²⁺-currents are measured in voltage-clamp mode using a holding potential of -80 mV and a 100-msec ramp-pulse to -40 mV to inactivate the fast Na⁺-current, followed by a 100-msec test-pulse to +10 mV at 0.5 Hz.