Simultaneous Isolation and Culture of Atrial Myocytes, Ventricular Myocytes, and Non-Myocytes from an Adult Mouse Heart

All research performed on mice reported in this paper has been reviewed and approved by the SDSU Institutional Animal Care and Use Committee and it conforms to the Guide for the Care and Use of Laboratory Animals published by the National Research Council.

1. Preparation of isolation and culture media and plating

1. Prepare 1 L of heart perfusion media prior to use by adding Joklik modified minimum essential media (MMEM) to 1 L of sterile water. Adjust the pH to 7.36 with 10 N NaOH, then filter through a 0.2 µm filter and store at 4 °C for up to 2 weeks.
   NOTE: Joklik MMEM consists of 112 mM NaCl, 5.4 mM KCl, 1 mM MgCl₂, 9 mM NaH₂PO₄, and 11.1 mM D-glucose. This is supplemented with 10 mM HEPES (2.38 g/L), 30 mM taurin (3.75 g/L), 2 mM D-l-carnitine (0.4 g/L), 2 mM creatine, and 10 mM butanedione monoxime (1.01 g/L).
2. Prepare digestion buffer just prior to perfusion, as follows: supplement 50 mL of heart perfusion media with 6.25 µL of 100 mM CaCl₂ (see step 1.3) and collagenase type 2 (enzymatic activity ~310–320 U/mg dw).
   NOTE: Weigh the animal prior to sacrifice. The amount of collagenase type 2 supplemented into the digestion buffer is dictated by the weight of the animal (2.25 mg of collagenase type 2 per 1 g of body weight).
3. To prepare 100 mM CaCl₂, add 1.47 g of CaCl₂ to 100 mL of molecular biology grade water. Stir until dissolved, then pass through a 0.2 µm filter and store at room temperature for up to 2 months.
4. Prepare myocyte stopping buffer 1 by supplementing 90 mL of heart perfusion buffer with 10 mL of fetal bovine serum (FBS) and 125 µL of 100 mM CaCl₂. Pass through a 0.2 µm filter and store at 4 °C for up to 2 weeks.
5. Prepare myocyte stopping buffer 2 by supplementing 114 mL of heart perfusion buffer with 6 mL of FBS and 150 µL of 10 mM CaCl₂. Pass through a 0.2 µm filter and store at 4 °C for up to 2 weeks.
6. Prepare atrial myocyte plating medium just prior to perfusion by supplementing 95 mL of Dulbecco’s modified Eagle medium (DMEM) with 4 mL of FBS, 1 mL of 100x pen/strep-glutamine, 1 mL of 100x insulin-transferrin-selenium, and 1 mL of 10 µM dexamethasone. Pass through a 0.2 µm filter and store at 37 °C until plating.
7. Prepare ventricular myocyte plating medium by supplementing 95 mL of minimum essential medium (MEM) with 4 mL of FBS, 1 mL of 100x pen/strep-glutamine, 10 mL of 1 M HEPES solution, 1 mL of 100x insulin-transferrin-selenium, and 0.1 g of butanedione monoxime. Pass through a 0.2 µm filter and store at 4 °C for up to 2 weeks.
8. Prepare ventricular myocyte maintaining medium by supplementing 99 mL of MEM with 1 mL of 100x insulin-transferrin-selenium, 0.1 mg/mL bovine serum albumin (BSA), 10 mL of 1 M HEPES solution and 1 mL of 100x pen/strep-glutamine. Pass through a 0.2 µm filter and store at 4 °C for up to 2 weeks.
9. To prepare laminin-coated experimental plates and slides, thaw mouse laminin stock solution (1.19 mg/mL). Add 10 µL of laminin stock solution to every 1 mL of DMEM, and mix. Coat experimental plates and slides evenly and store in a 37 °C, 5% CO₂ incubator for at least 1 h prior to perfusion to allow for equilibration.
   NOTE: Coated experimental plates and slides can be stored at 4 °C for up to 2 days.

2. Isolation apparatus

1. Prior to each isolation, clean the tubing and other components of the system with three complete washes of 70% EtOH by filling the bubble trap to the top (close off the bottom stopcock and keep the top stopcock open).
2. Rinse the complete system 3x with sterile H₂O by filling the bubble trap to the top (close off the bottom stopcock and keep the top stopcock open).
3. Rinse out the complete system with heart perfusion buffer and fill the bubble trap halfway with media.
4. Set the circulating water bath to 37 °C.
5. Set a water bath for media to 37 °C.
6. Remove any air bubbles in the peristaltic pump tubing.
7. Adjust the flow rate of the peristaltic pump to 3 mL/min.

3. Surgical procedure (non-survival)

1. Soak surgical instruments in 70% EtOH for at least 5 min.
2. Place the heart perfusion media, myocyte stopping buffers, and digestion buffer in the 37 °C water bath.
3. Place the atrial myocyte plating medium, ventricular myocyte plating medium, and ventricular myocyte maintaining medium in a 37 °C, 5% CO₂ incubator 1 h prior to use and loosen the caps to allow equilibration.
4. Inject 10-week-old male or female C57b6/j mice intraperitoneally (i.p.) with 0.35 mL of heparin, diluted in phosphate buffered saline (PBS) to 100 IU/mL. Allow the drug to take effect for about 10 min.
   NOTE: If two hearts are to be subjected to this isolation procedure, the second mouse can be anesthetized and administered heparin at this point in the first procedure. To minimize stress on the animal, light anesthesia can be administered using 2% isoflurane/oxygen mixture in a hermetically sealed induction chamber.
5. Anesthetize the animal with a 2% isoflurane/oxygen mixture and inject i.p. with pentobarbital (0.3 mL from 10 mg/mL stock), then prepare the chest by swabbing with 70% EtOH.
6. Prepare a tied 5-0 silk suture to be ready to ligate the heart to the perfusion cannula. Mount cannula right next to surgical microscope.
7. Quickly open the chest by first making a midline skin incision, from mid-abdomen to the jaw, then entering the peritoneum with the large scissors, clearing the diaphragm away by blunt dissection. Then, cut away the rib cage using the scissors with cuts up the chest wall on the lateral aspect of both sides.
8. Snip away fibrous connections between the heart and chest wall (including thymus). Then, cut away the rib cage all together. Using the small forceps and scissors, gently lift the heart by the apex and expose the posterior aspect of the heart.
9. Explant the heart by dissecting immediately inferior to the innominate artery on the ascending aorta and immediately place the heart in ice-cold PBS or cold heart perfusion media. Subsequently, quickly dissect away remaining tissue from the explanted heart in ice-cold heart perfusion media, exposing the ascending aorta.
   NOTE: It is useful to explant the heart with the thymus intact to use as an anatomical landmark.
10. Clean the area surrounding the aorta of excess tissue using micro-dissecting forceps and scissors. Position the aorta onto the cannula using fine-tipped forceps and secure with a 5-0 silk suture.
    NOTE: The best placement usually occurs with the aorta extending about 2 mm up on the cannula.
11. Perfuse the cannulated heart with heart perfusion media at a flow rate of 3 mL/min for 4 min. Then switch the media from heart perfusion media to digestion buffer for 15.0−17.5 min of perfusion (Figure 1A).
    NOTE: Coronary flow rate will likely increase, indicating effective tissue digestion (i.e., pale, swollen heart).
12. Collect 8 mL of digestion buffer flow through during the final minutes of perfusion for later use in step 5.4.
13. Remove the heart from the cannula and place it on a 60 mm plastic culture dish. Remove excess tissue (i.e., aorta, veins) and submerge the heart in 2.5 mL of digestion buffer in preparation for mechanical separation.
    NOTE: At this point, the atria are dissected away, and one experimenter should conduct the atrial cell isolation protocol, while a second experimenter should conduct the ventricular cell isolation.

4. Atrial cell isolation and culture

1. Dissect the atria away from the heart and place it into a 30 mm plastic culture dish. Submerge it in 0.75 mL of digestion buffer for mechanical separation. Keep the ventricles in the 60 mm dish (step 3.12) and carry out the separate isolation methods simultaneously (section 5, Figure 1B).
   NOTE: At this point, the atria and ventricles can undergo further separation should there be a need to isolate the left- and right-side cells.
2. Begin to mince and tease the atria apart, initially with fine-tip surgical scissors and followed by fine forceps for further mincing. Avoid agitating the tissue, and do not rapidly pull apart muscle fibers.
3. Using a sterile transfer pipette tip, continue to gently mix and dissociate the tissue for 15 min. Every 5 min, observe atrial myocyte disassociation from tissue under a 10x objective brightfield microscope. As tissue becomes further digested, continue gently mixing and dissociating tissue using a sterile transfer pipette tip with a smaller pore size.
4. Transfer cell suspension to a 2 mL sterile micro-centrifuge tube. Rinse the 30 mm plate with 0.75 mL of 37 °C myocyte stopping buffer 1 and combine with the cell suspension (end volume = 1.5 mL).
5. Allow the atrial myocytes to sediment by gravity for 10 min at room temperature. Gently agitate the cell suspension to allow for myocytes to sediment to the bottom of the conical tube, forming a visible pellet.
6. Centrifuge the cell suspension for 5 min at 20 x g. Carefully remove the supernatant, which contains the non-myocytes, and transfer to a 15 mL polypropylene conical tube using a sterile pipette tip without disturbing the pellet of atrial myocytes.
7. Centrifuge the non-myocyte fraction for 5 min at 20,000 x g. Aspirate the supernatant and resuspend the non-myocyte pellet in 10 mL of DMEM supplemented with 10% fetal calf serum (FCS).
8. Count non-myocytes using a hemocytometer or other method, then plate as per experimental needs, or further isolate into individual specific cell populations via fluorescence-activated cell sorting (Figure 1C).
9. Resuspend the pellet of isolated atrial myocytes from step 4.6 in 1 mL of atrial myocyte plating medium and apply 10 µL of this suspension onto a hemocytometer. Perform a cell count of rod-shaped myocytes per field.
10. Aspirate laminin from precoated experimental plates/slides and resuspend the isolated atrial myocytes in the appropriate volume of atrial myocyte plating medium supplemented with 25 µM blebbistatin. Plate at the desired density per experimental needs (Figure 1C).
    NOTE: Typical plating density for the long-term culturing described here is 5 x 10⁵ cells/chamber on four-chamber (1.7 cm²) glass slides.

5. Ventricular cell isolation and culture

1. Begin to mince and tease heart apart ventricular tissue, first with fine-tip surgical scissors and followed by fine forceps for further mincing (Figure 1B). Avoid agitating the tissue by rapidly pulling apart muscle fibers.
2. Transfer the cell suspension to a 15 mL polypropylene conical tube. Rinse the plate with 2.5 mL of 37 °C myocyte stopping buffer 1 and combine with the cell suspension (end volume = 5 mL).
3. Using a sterile transfer pipette tip, continue to gently mix and dissociate the tissue for 4 min. Apply 10 µL of this cell suspension onto the slide and visualize the presence of rod-shaped myocytes to ensure the quality of isolation.
4. Pass the cell suspension through a 100 µm sterile nylon filter into a 50 mL polypropylene conical tube. Use 2 mL of digestion buffer collected in step 3.12 to wash any remaining cells off the sterile nylon filter.
5. Allow the ventricular myocytes to sediment by gravity for 6 min at room temperature. Gently agitate the filtered cell suspension to allow for myocytes to sediment at the bottom of the conical, forming a visible pellet.
6. Without disturbing the pellet of ventricular myocytes, carefully remove the supernatant (non-myocytes) and transfer to a 50 mL polypropylene conical tube using a sterile pipette tip. Centrifuge the non-myocyte fraction for 5 min at 20,000 x g. Aspirate the supernatant and resuspend the non-myocyte pellet in 10 mL of DMEM supplemented with 10% FCS.
7. Count non-myocytes using a hemocytometer and resuspend in an appropriate volume of DMEM supplemented with 10% FCS. Then, plate according to experimental needs, or further isolate into individual specific cell populations via fluorescence-activated cell sorting (Figure 1C).
8. Resuspend the isolated ventricular myocytes in 2 mL of myocyte stopping buffer 2 and apply 10 µL of cell suspension onto hemocytometer. Perform a cell count of rod-shaped myocytes per field.
9. Reintroduction of Ca²⁺ using a stepwise paradigm
   NOTE: Calcium reintroduction steps are specific for ventricular myocytes and should not be performed for atrial myocytes, as this will result in cell death.
   1. Add 50 µL of 10 mM CaCl₂ to the ventricular myocyte cell suspension. Mix well and incubate for 4 min at room temperature.
   2. Add an additional 50 µL of 10 mM CaCl₂ to the ventricular myocyte cell suspension. Mix well and incubate for 4 min at room temperature.
   3. Add an additional 100 µL of 10 mM CaCl₂ to the ventricular myocyte cell suspension. Mix well and incubate for 4 min at room temperature.
   4. Add 80 µL of 100 mM CaCl₂ to the ventricular myocyte cell suspension. Mix well and incubate for 4 min at room temperature.
10. Remove laminin coating from plates or slides and resuspend the isolated ventricular myocytes in an appropriate volume of ventricular myocyte plating medium according to experimental needs (Figure 1C).
    NOTE: Typical plating density for the long-term culturing described here is 5 x 10⁵ cells/chamber on four-chamber (1.7 cm²) glass slides. Avoid plating at a density of greater than 75% confluency, as cellular over-densification promotes cell clumping, thereby inhibiting attachment to the plate. Furthermore, large numbers of cells will be lost during the first medium change. Plate cells quickly after isolation, as extracellular Ca²⁺ promotes hypercontraction and loss of viable myocytes.
11. Allow the ventricular myocytes to settle and adhere for at least 1 h. Subsequently change the media to ventricular myocyte maintaining medium supplemented with 25 µM blebbistatin.
    NOTE: Ventricular myocytes can be cultured up to 96 h after plating in µM blebbistatin. Experiments should be conducted in ventricular myocyte maintaining medium in the absence of blebbistatin.