Isolation, Culture, and Functional Characterization of Adult Mouse Cardiomyoctyes

Summary

Here we describe the isolation of adult mouse cardiomyoctyes using a Langendorff perfusion system. The resulting cells are Ca2+-tolerant, electrically quiescent and can be cultured and transfected with adeno- or lentiviruses to manipulate gene expression. Their functionality can also be analyzed using the MMSYS system and patch clamp techniques.

Abstract

The use of primary cardiomyocytes (CMs) in culture has provided a powerful complement to murine models of heart disease in advancing our understanding of heart disease. In particular, the ability to study ion homeostasis, ion channel function, cellular excitability and excitation-contraction coupling and their alterations in diseased conditions and by disease-causing mutations have led to significant insights into cardiac diseases. Furthermore, the lack of an adequate immortalized cell line to mimic adult CMs, and the limitations of neonatal CMs (which lack many of the structural and functional biomechanics characteristic of adult CMs) in culture have hampered our understanding of the complex interplay between signaling pathways, ion channels and contractile properties in the adult heart strengthening the importance of studying adult isolated cardiomyocytes. Here, we present methods for the isolation, culture, manipulation of gene expression by adenoviral-expressed proteins, and subsequent functional analysis of cardiomyocytes from the adult mouse. The use of these techniques will help to develop mechanistic insight into signaling pathways that regulate cellular excitability, Ca²⁺ dynamics and contractility and provide a much more physiologically relevant characterization of cardiovascular disease.

Introduction

Murine models of cardiovascular disease have served as effective tools for elucidating fundamental disease mechanisms^1,2 as well as for identifying potential therapeutic targets^1,3. In particular, the use of both murine models of acquired heart disease (such as pressure-overload)^4,5 and transgenic mouse models have advanced our understanding of heart disease^6-8. The use of cell culture techniques to study signaling cascades^3,9,10 and alterations in individual proteins that underlie cellular excitability and excitation-contraction coupling in the heart^11-13 at the level of the single cell have complemented the in vivo mouse models. However, the lack of adequate cell lines that reflect adult CM structure and function has been a significant limitation. Investigators have sought to overcome this by studying individual proteins, such as ion channels, in heterologous expression systems¹⁴, and while these studies have provided us with useful information in terms of ion channel biophysics or protein trafficking, inadequate representation of the native microenvironment of CMs is a significant limitation. Secondly, since most of these heterologous cells do not have a mature contractile apparatus, it has not been possible to study contractile function and the complex interplay between cellular excitability and contraction. For this reason, researchers have turned to primary cardiac cell cultures for many of their in vitro functional studies. Finally, isolated cardiomyocyte studies allow assessment of contractile function without the confounding factors of multicellular preparation including the effect of scar or fibrosis and fiber orientation.

Primary neonatal rat ventricular cardiomyocytes (NRVMs) are relatively easy to culture, can be infected with adenoviruses and lentiviruses to manipulate gene expression¹⁵, and have therefore been used successfully¹, but have limitations of their own. Although they provide a physiologic microenvironment¹ and have been the workhorse of the signaling field, substantial differences between the morphology and subcellular organization of NRVMs and adult cardiomyoctyes make them an inadequate model for the investigation of ionic fluxes and excitation-contraction coupling in the adult heart. Most notably NRVMs lack a definitive t-tubular subsystem⁴. Since Ca²⁺ flux and dynamics are critically dependent on mature t-tubular and sarcoplasmic reticulum (SR) structure⁶, Ca²⁺ dynamics and functional studies of the cardiac contractility in NRVMs are not an accurate reflection of these critical processes in adult cardiomyocytes. Further, some components of signaling pathways differ between neonatal and adult mice⁹, thereby providing another limitation for studying disease processes and their impact on cellular excitability and contractility in NRVMs. Finally, the distribution of the contractile machinery leads to multidirectional and non-uniform cell shortening limiting the accuracy of the contractile measurements.

The use of isolated adult cardiomyocytes provides therefore a more accurate in vitro modeling system. The extraordinary growth of knowledge made possible by the genetic manipulation of mice underlines the significance of obtaining functional isolated cardiomyocytes from mice. In fact, the characterization of adult CMs isolated from mouse models has shed light on many biological and pathological events. Isolated CMs from transgenic mouse models have allowed for studies of the gain or loss of function of proteins on the contractile properties of single cells^2,16, and viability in disease models such as ischemia/reperfusion^17,18, thereby complementing information gained from in vivo studies on these mice. Use of isolated adult CMs from murine models of acquired heart disease^3,19,20 (such as transverse aortic constriction-induced pressure overload, that mimics hypertension or aortic valve stenosis) or exercise^5,21 (for modeling physiological hypertrophy) allows for examination of the interaction of signaling cascades implicated in these processes with cellular excitability and excitation-contraction coupling at the level of the single cell. Furthermore, the ability to manipulate gene expression using adenoviral-driven gene expression in adult CMs affords us the opportunity to dissect the components of complex signaling pathways.

From an electrophysiological perspective, whole-cell voltage and current clamp experiments on isolated adult CMs have been critical in elucidating the nature of ionic fluxes at baseline and in various disease states. Because of the complex structure of the cellular membrane and the differential protein scaffolding structures between adult CMs and NRVMs or heterologous cell lines, the ability to patch adult cells gives a much better representation of the effects of certain membrane proteins, structural proteins, and ion channel interacting partners on the electrophysiological components of the adult heart.

Despite such prominent advantages in studying adult murine cardiomyocytes, isolating and culturing adult mice cardiomyocytes has been challenging, urging the need for a systematic and accurate description of the methodology to isolate viable mouse cardiomyocytes and to maintain them in culture to allow further genetic manipulation using viral vectors. Previous studies have used either acutely isolated mouse adult CMs or cultured rat adult CMs. The latter are easier to culture than adult mouse CMs, and most experiments manipulating gene expression in vitro have used rat adult CMs. Few studies have successfully altered and investigated functional gene expression in mouse adult CMs, presenting a large limitation in the scope of experiments. Therefore, here we present in detail such methodologies, modified from previous investigations, for the isolation^7,8,22, culture^3,10,15,23, adenoviral infection^11-13,15, and functional analysis of adult mouse ventricular cardiomyoctyes. This isolation protocol results in Ca²⁺-tolerant, excitable cardiomyocytes that we have successfully cultured for up to 72 hr and transiently transfected with adenovirus. The functionality of these isolated cells can be assessed using the MMSYS imaging system^14,24 and patch clamp, which will also be discussed.

Protocol

1. Cardiomyocyte Isolation

Materials (Figure 1)

Microdissecting forceps
Tissue forceps
Delicate hemostatic forceps
Hemostatic forceps
Microdissecting, serrated, curved forceps
Operating scissors, straight
Operating scissors, curved
15 ml Falcon tubes (5)
60 mm Petri dish
Phosphate Buffered Saline (PBS)
Nylon Mesh - 400 μm pore size
Small funnel
Wax coated, braided silk 4-0, 19 mm, 7-10 cm long
Non-hypodermic needle, blunt end, 24 G or commercial animal-feeding needle (24 x 1 in, W/1-1/4)
Heparin sodium
Ketamine/xylazine mixture
Pasteur Pipettes
OptiVision Dissecting Goggles
IonOptix MMSYS system

Note: If culturing cells, see section 2 on cell culture before beginning this protocol.

Note: All mice were cared for in a barrier facility and sacrificed according to approved IACUC regulations, practices, and procedures.

Note: Prior to beginning the procedure, the entire system should be preheated to ensure that all the elements of the Langendorff system (Figure 2A, Figure 3) are warmed to 37 °C to allow for proper and complete collagenase activity.

1. Inject adult mice (~12 weeks) with 150 USP units heparin sodium into the abdominal space.
2. Anesthetize with an injection of a ketamine/xylazine mixture at a weight-dependent dose (Table 1).
   1. 80:12 mg/kg ketamine:xylazine recipe: 2.6 ml ketamine (100 mg/ml) + 0.4 ml xylazine (100 mg/ml) + 37 ml PBS. Final: ketamine = 6.5 mg/ml; xylazine = 1 mg/ml
3. Secure the sedated mouse to the operating pad, excise the heart, and place into a dish of room temperature PBS or 0.9% saline (Figure 1J). Physiologic saline allows the intact heart to contract for better extrusion of blood in the chamber and easy identification of the aorta prior to step 1.6. Mice were checked to ensure that they are deeply anesthetized via loss of hind limb toe pinch reflex and slowing of respiratory rate prior to onset of thoracotomy.
4. Use OptiVision dissecting goggles (Figure 1A) to identify and expose the aorta. Removing the tissue around the aorta, although it does facilitate the visualization of the vessel, is not necessary for the optimization of the cell isolation if the aortic root is clearly visible.
5. Prime the pump with perfusion buffer (Table 2). Temperature should be held at 37 °C for the length of the procedure using a controlled, circulating water bath (Figure 2D).
6. Mount the heart onto the Langendorff Apparatus (Figure 2A). Using two micro-dissecting forceps (Figures 1D-E), prime the aorta around the non-hypodermic, blunt end needle (24 G) with silicone tube at the distal tip. Alternatively a commercial animal-feeding needle (24 x 1 in, W/1-1/4) can be used. The silicone tube on the 24 G needle or the animal feeding needle will allow for securing the suture over the groove. Place the aorta on the needle as a sock. (Figure 2C).
   1. Note: It is important to be sure that the end of the needle rests in the ascending aorta, ideally in the aortic root distal to the right innominate, but that it does not extend through the aortic valve into the left ventricle, as this will severely limit the ability of the buffers to effectively perfuse through the heart via the coronary arteries.
7. Secure the heart to the needle by tying a small length of suturing silk (7-10 cm long) (Figure 1K) around the top of the aorta, ensuring that the tie is at the level of the ascending aorta, below the right innominate artery, but is above the end of the needle.
8. Lower the heart into the interior of the conical glass (Figure 2B) to help maintain a suitable ambient temperature.
9. Perfuse the heart with perfusion buffer for 5 min at a rate of 1 ml/min and clamp the glass chamber outflow to allow the perfusate to collect in the conical glass and envelope the heart. The schematic for digesting the heart is shown in Figure 3.
   1. At this point you should see the volume of the heart increase. Additionally, blood in cardiac tissue will be replaced by perfusate, causing tissue pallor.
10. Release the outflow clamp to drain the perfusate. Stop the pump to move the tubing from the perfusion buffer container to the enzyme buffer container (Figure 2E). Restart the pump to begin to perfuse the heart with enzyme buffer (Table 2) at 1 ml/min. Clamp the outflow again to allow the enzyme buffer to envelope the heart.
    1. Here, as the enzyme is perfusing the heart and digesting the connective tissue, the heart will become paler and the structural integrity will diminish.
    2. To determine when to cut the ventricles from the digested heart, lift the heart out of the conical glass, gently squeeze the heart with forceps and collect a few drops of perfusate in a small Petri dish and check to see whether or not single cells are dropping.
    3. For the novice it is helpful to connect the perfusion line with a manometer. When the heart is getting digested the pressure will rapidly decline, as the connective tissue does not hold the resistance. The manometer is also useful for the novice to ensure that the position of the needle is above the aortic valve and not in the ventricle. Low pressure will indicate that the needle is in the ventricular chamber and high pressure indicated that the needle touches the valve.
11. Cut the ventricles from the heart into a dish full of transfer buffer (A) (Table 2) when the ventricular pressure begins to drop dramatically or when you are able to see single cells in the perfusate. This usually takes ~7-10 min.
12. Again using micro-dissecting forceps (Figure 1C), mince the ventricles into small pieces to dissociate. A successful digestion will leave almost no solid chunks of tissue after mincing, with most of the tissue becoming amorphous upon dissociation.
    1. To further dissociate the tissue, pipette the solution in/out using a plastic Pasteur pipette (Figure 4) from which the tip had been cut at an ~45° angle. This modification serves to increase the inner diametric space of the pipette tip thus decreasing the amount of shear stress on the cells as the tissue is triturated.
    2. Before dissecting the tissue with the forceps it is possible to separate the individual chambers and separately dissociate atrial, left ventricular or right ventricular cells.
13. Secure the squared mesh (Figure 1M) to a small funnel (Figure 1L) using clamps and place this filtering apparatus into a 15ml Falcon tube (Figure 1I).
14. Use the modified Pasteur pipette to remove the cell solution from the dish and filter the solution into the Falcon tube.
15. Allow the live cells to settle to the bottom of the tube (live cell sedimentation should take 2-4 min).
16. Aliquot 2.5ml of each of the four calcium solutions (Table 3) into 4 separate 15ml falcon tubes.
17. Again using a standard Pasteur pipette, carefully remove the cell pellet from the bottom of the tube and transfer the cells to the first of the calcium solutions.
18. Allow the cells to settle to the bottom of the tube and repeat step 1.17 in the subsequent calcium solutions until the cells are in the last solution.
19. Do not let the cells settle in the fourth calcium solution. Cap the tube and turn it on its side.

2. Cell Culture

1. Before the isolation, plate 1 ug/ml natural mouse laminin onto glass coverslips and incubate for at least 1 hr at 37 °C and 2% CO₂. Also allow your plating and culture media (Table 4) to warm in the incubator at 37 °C and 2% CO₂. This allows the dissolved CO₂ in the media to equilibrate.
   1. Do not remove the top from the media. Simply loosen the top to avoid contamination.
2. Allow the isolated cells to pellet at the bottom of the Falcon tube.
3. Remove the pellet using a standard plastic Pasteur pipette and resuspend in the appropriate amount of plating media.
4. Plate the cells on the laminin-coated coverslips in the appropriate sized dish and incubate in 37 °C and 2% CO₂ for at least 30-60 min to allow the cells to adhere to the coverslips.
5. **If transfecting with adenovirus, dilute the virus in an appropriate amount of plating media and replace the plating media in the dish with the virus-containing media. Let the virus incubate on the cells for 2 hr in 37 °C and 2% CO₂.**
6. Remove the plating media from the dish and replace with culture media.
7. Carefully change the culture media each day so as not to disturb the cells on the coverslips.

Using this procedure, cells have been successfully cultured for up to 72 hr. Images of cultured and GFP transfected cells can be found in Figure 5.

3. MMSYS System

1. Power the MMSYS system ensuring that the arc lamp is initiated first.
2. Connect the tubes from the pump to the appropriate inlet and outlets of the MMSYS chamber (Figure 6).
3. Prime the system with calcium buffer (B) (Table 2).
4. Place an appropriately sized glass coverslip into the chamber and fasten (Figure 6E). Most MMSYS chambers use either 22 mm or 25 mm square coverslips. Consult your MMSYS user manual to determine the appropriate coverslip for your chamber.
5. Transfer 500 μl of the cell solution to a small Eppendorf tube.
6. Add 0.5 μl Fura2-AM (stock solution of 1 μg/μl) to the small tube of cells and allow them to incubate in the dark at RT for 5-7 min. This allows the Fura2-AM to "load" into the cells through rapid passive diffusion through the cell membrane.
   1. Let the Fura-2 loaded cells pellet to the bottom of the tube, and wash the excess Fura with 500 μl of calcium buffer B.
7. Turn off the room lights.
8. Using a standard Pasteur pipette, drop 1-2 drops (depending of the cell concentration) of the "loaded" cell solution into the center of the chamber (Figure 6C) and allow the cells to settle onto the coverslip for 5 min.
   1. The density of the cells in the chamber should be such that single non-overlapping cells can be easily viewed in the MMSYS data acquisition platform.
9. Carefully begin to flow calcium buffer (B) through the chamber.
10. Increase the chamber temperature to 37 °C.
    1. IMPORTANT: Do not turn on the heating apparatus until the flow has been initiated. This can severely damage the feedback thermocoupler (Figure 6A).
11. Make sure that the chamber is connected to the MyoPacer via the connection wires (Figure 6B) and begin to pace the cells 5 Hz at 1.5x the pacing threshold for the cells.
12. Choose a cell that is beating at the correct frequency and move it into the framing aperture.
13. Adjust the camera and framing aperture dimensions so that the entire cell is in the center of the window, directed horizontally, with the sarcomeres apparent.
    1. For cell length adjust the red and green (right and left) photodiode indicators to the edge of the cell. Adjust the contrast to optimize the black/white contrast at the edges of the cell. Refer to the Ionoptix manual for further details.
14. Place the box in an area of the cell containing well-defined sarcomeres and adjust the focus to optimize the peak of the power spectrum (red). Also adjust the brightness of the microscope to optimize the blue smoothing window and black contrast information.
15. Adjust the green threshold bars to capture as much of the red power spectrum (peak) and as little background as possible.
16. Begin recording.
17. After enough data has been recorded, click "Pause" to temporarily stop recording and, ensuring that neither the focus nor dimensions of the viewing window are altered, move the microscope's stage to an area of the coverslip in which no cells or cellular material can be seen. Length of recording varies depending on the investigator's experimental protocol.
    1. Note: Clicking "Stop" will terminate the recording and no background will be able to be recorded for that cell.
18. Click "Resume" to record a background measurement.
19. After enough background has been recorded click "Stop" to terminate the recording.
20. Click "File >> Save" to save all the traces for that cell in a single .zpt file.
21. Repeat steps 3.12 - 3.20 until enough cells have been recorded.
22. Consult the IonOptix-MMSYS User Manual 12 for instructions on analysis of the recorded data. Characteristic recorded data of wild-type cardiomyocytes are shown in in Figure 7.

4. Patch Clamp

Patch clamping of different cell types have been well described previously in this journal^25-28. We therefore focus on some critical parameters for successful patch clamping of adult cardiomyocytes isolated using our protocol described.

1. Using a pipette puller and borosilicate glass (with filament: O.D. 1.5 mm, I.D. 0.86 mm - 10 cm length) pull a pipette that exhibits ~2.0-3.0 MΩ when filled with pipette solution (Table 5).
2. Fire-polish the pipette tip gently using a Narishige or other vertical fire polisher. The resistance of the patch pipette should be 2-4 MO after fire polishing.
3. Turn on the computer, amplifier (Axopatch 200B), A-D converter (Digidata 1440A, Axon Instruments) and Micromanipulator (MP-285). For whole cell patch clamping, we start with standard parameters as previously described.
4. For cultured adult CMs, remove them from the incubator and gently wash the coverslip on which the CMs are plated with PBS at room temperature.
5. Gently remove the coverslip and place it in the perfusion chamber on the inverted microscope (Nikon TE 2000).
   1. Ensure that the inlet for the perfusion system and outlet (for suction) are just above the surface of the coverslip.
6. Start perfusing with the appropriate extra-cellular solution (Table 5).
7. For freshly dissociated adult CMs, place a collagen-coated coverslip in the perfusion system as above.
   1. Since the cells will be in solution, gently replace the solution with extracellular buffer.
8. Using the modified Pasteur pipette (Figure 4), gently resuspend the cells in the extracellular buffer, take an aliquot and place it on the coverslip.
9. Let the cells attach for 10-15 min before starting the perfusion as in 4.4.
10. Back-fill the whole cell patch pipette with intra-cellular buffer using a Microfil (World Precision Instruments, 28 G) needle. Ensure that there are no air bubbles in the tip of the pipette; gently tap to allow the air bubbles to float to the surface of the solution.
11. Attach the filled patch pipette to the pipette-holder, ensuring that there is contact between the microelectrode and the internal solution.
    1. We use silver chloride electrodes, but take care of 'chloriding' the silver wires at least once a week by immersing overnight in household bleach.
12. Gently lower the pipette into the bath, ensuring that the bath electrode is immersed at all times in the bath/extra-cellular solution. Offset any liquid junction potentials at this time. Large junction potentials may be a sign of deteriorating silver chloride on the electrodes.
13. Healthy cylindrical adult CMs are identified in the microscope and centered in the field of vision. As previously described, a combination of moving the micromanipulator and adjusting the focus allows for close proximity of the patch pipette and the surface of the CM.
    1. Once they are in the same focal plane, we use a combination of visualization of the cell and test pulse (available in the Axopatch 200B).
    2. Once the resistance of the pipette decreases in half (suggesting that the pipette is in contact with the surface of the cell), and the cell continues to look cylindrical, we establish gigaohm seal using gentle suction.
    3. For CMs, we prefer mouth suction to establish negative pressure. If a gigaseal is successfully obtained, we again use gentle rhythmic mouth suction to 'break-in' to the cell to establish whole cell patch mode.
14. Typical resting potentials of healthy CMs are in the order of -85 to -90 mV. Once a gigaseal is obtained, measure the resting membrane potential using current clamp with I=0 pA.
    1. If the resting membrane potential is depolarized this can indicate a damaged cell or a poor seal.
15. Because the CMs are large cells with a lot of surface area, careful attention needs to be paid to capacitance compensation, especially when rapidly activating currents such as the sodium channel needs to be studied. If adequate capacitance compensation cannot be obtained using the built in compensation settings on the pulse generator, prepulses at sub-threshold voltage and opposite polarity may have to be applied and the resultant current subtracted from the recorded signal. Such a protocol is easily programmed in Axopatch.
16. Once whole cell patch mode has been established, wait 15 min to allow for equilibration and to ensure stability of the seal prior to beginning voltage or current clamp protocols. We use standard protocols that have been described previously in this and other journals and will not be elaborated upon.

Results

The isolation of adult cardiomyoctyes results in rod-shaped, striated, and quiescent (not spontaneously beating) cells (Figure 5A). Dead cells will look rounded and no striations will be present. Quiescent cells can be cultured and transfected with adenovirus to manipulate gene expression (Figures 5B and 5C). After 24 hr of culture, the morphology of the live cells does not change, they are still Ca²⁺-tolerant, and they can be paced by field stimulation. With ...

Discussion

In this report, we have described the techniques necessary for successful isolation and culture of adult CMs from the mouse heart. Our technique allows for subsequent study of CM function and excitability using the methods described above. The critical parameter for studying functionality of adult CMs is the health and quality of the isolated CMs. As described above, our techniques allow for a high yield of functional cells that are amenable to manipulation of gene expression using adenoviral/lentiviral infections in ...

Materials

Name	Company	Catalog Number	Comments
Sodium Chloride	Sigma	S7653
Potassium Chloride	Sigma	P9333
Magnesium Chloride	Sigma	M8266
HEPES	Sigma	H3375
Sodium Phosphate Monobasic	Sigma	S8282
D-glucose minimum	Sigma	G8270
Taurine	Sigma	T0625
2,3-Butanedione monoxime	Sigma	B0752
Collagenase B	Roche Applied Science	11088807001
Collagenase D	Roche Applied Science	11088858001
Protease XIV from Streptomyces griseus	Sigma	P5147
Albumin from Bovine Serum	Sigma	A2153
Calcium Chloride	Sigma	C8106
Minimum Essential Media	Sigma	51411C
Albumin solution from bovine serum	Sigma	A8412
L-glutamine	Sigma	G3126
Penicillin-Streptomycin	Sigma	P4333
Insulin-transferrin-sodium selenite media supplement	Sigma	I1884
Cesium Chloride	Sigma	289329
Glutamate	Sigma	G3291
Adenosine 5'-triphosphate magnesium salt	Sigma	A9187
Ethylene glycol-bis(2-amin–thylether)-N,N,N',N'-tetraacetic acid	Sigma	E3889
Cesium Hydroxide Solution	Sigma	232041
Tetraethylammonium hydroxide solution	Sigma	86643
OptiVisor optical glass binocular visor	Dohegan Optical Company Inc.	N/A
Tissue forceps, 5.5", 1x2 teeth	Roboz Scientific	RS-8164
Moloney forceps - 4.5" (11.5 cm) long slight curve, serrated	Roboz Scientific	RS-8254
Dumont #3 Forceps, Dumostar, tip size 0.17 x 0.10mm	Roboz Scientific	RS-4966
Packer Mosquito Forceps 5" Straight Flat	Roboz Scientific	RS-7114
Micro Dissecting Scissors 4.5" Curved Sharp/Sharp	Roboz Scientific	RS-5917
Micro Dissecting Scissors 3.5" Straight Sharp/Sharp20mm	Roboz Scientific	RS-5907