Measuring mitochondrial function in intact cardiac myocytes

Abstract

Mitochondria are involved in cellular functions that go beyond the traditional role of these organelles as the power plants of the cell. Mitochondria have been implicated in several human diseases, including cardiac dysfunction, and play a role in the aging process. Many aspects of our knowledge of mitochondria stem from studies performed on the isolated organelle. Their relative inaccessibility imposes experimental difficulties to study mitochondria in their natural environment-the cytosol of intact cells-and has hampered a comprehensive understanding of the plethora of mitochondrial functions. Here we review currently available methods to study mitochondrial function in intact cardiomyocytes. These methods primarily use different flavors of fluorescent dyes and genetically encoded fluorescent proteins in conjunction with high-resolution imaging techniques. We review methods to study mitochondrial morphology, mitochondrial membrane potential, Ca(2+) and Na(+) signaling, mitochondrial pH regulation, redox state and ROS production, NO signaling, oxygen consumption, ATP generation and the activity of the mitochondrial permeability transition pore. Where appropriate we complement this review on intact myocytes with seminal studies that were performed on isolated mitochondria, permeabilized cells, and in whole hearts.

Figure 1. Schematic overview of mitochondrial structure and transport mechanisms

For details and abbreviations see text.

Figure 2. Measurements ofΔΨ_m and mPTP activity in permeabilized ventricular myocytes and isolated mitochondria

A, ΔΨ_m monitored with the membrane potential sensitive dye TMRM in permeabilized rabbit ventricular myocytes upon inhibition of ETC complex I with rotenone and subsequent exposure to ETC complex II substrate succinate. The mitochondrial uncoupler FCCP completely depolarized Ψ_m. B, mPTP activity monitored with mitochondria entrapped calcein (620 Da) in permeabilized cat ventricular myocytes. Opening of mPTP was induced by enhanced ROS formation (prevented by superoxide and peroxynitrite scavenger MnTBAP, 50 μM) resulting from Ca²⁺-induced stimulation of mtNOS in the absence of L-arginine. Opening of mPTP resulted in the loss of calcein and a decrease of fluorescence. The pore forming antibiotic alamethicin induced maximum calcein release. Cyclosporine A (CsA, 5 μM) blocked mPTP opening. C, mPTP openings recorded from isolated mitochondria with TMRE. PTP opening can occur as brief incomplete (arrows) and prolonged maximal depolarizations. D, simultaneous monitoring of mPTP activity as changes in ΔΨ_m (TMRE) and loss of calcein in isolated mitochondria. Brief incomplete depolarizations (arrow) did not cause loss of calcein, however prolonged and complete depolarizations were accompanied by loss of calcein. The decline of calcein fluorescence was preceded by a transient increase (asterisk) caused by calcein dequenching. (Panel B: from Dedkova and Blatter, The Journal of Physiology, 2009, with permission; panels C and D: from Hüser, Rechenmacher and Blatter, Biophysical Journal, 1998, with permission).

Figure 3. Measurements of [Ca²⁺]_m with compartmentalized fluo-3 in single permeabilized ventricular myocytes

A, top: changes in fluo-3 fluorescence following membrane permeabilization with digitonin and increase in extramitochondrial [Ca²⁺] ([Ca²⁺]_em). Images a to c (bottom) were taken at times indicated by arrows. B, changes of [Ca²⁺]_m in response to rapid cytosolic Ca²⁺ transients. a and b, technique used to generate rapid changes in [Ca²⁺]_em to simulate cytosolic Ca²⁺ transients. Cells were placed in the laminar flow of zero-Ca²⁺ solution (1 mM EGTA) and computer controlled pressure ejections of 10 μM Ca²⁺-containing solution from a glass micropipette. c, changes of [Ca²⁺]_m in a permeabilized myocyte in response to stimulation with Ca²⁺ pulses applied at 0.5 Hz (pulse duration = 0.5 s; [Na⁺]_em = 40 mM; [Ca²⁺]_pip = 10 μM). (From Sedova, Dedkova & Blatter, Am. J. Physiol. Cell Physiol., 2006; Am. Physiol. Soc., used with permission)

Figure 4. Monitoring mitochondrial redox potential in intact ventricular myocytes by measuring FAD autofluorescence

Top, from left to right: confocal images of ventricular myocyte FAD autofluorescence (excitation 488 nm, emission 510–525 nm) before (cntrl) and after exposure to ETC complex II substrate methyl succinate (Met-Succinate), ETC complex IV inhibitor sodium cyanide (NaCN), and mitochondrial uncoupler FCCP. Bottom, time course of changes in FAD/FADH₂ fluorescence recorded from the regions of interest shown on top. Exposure to NaCN represents maximal reduction of FAD (i.e., 0% fluorescence from FAD with maximal formation of FADH₂), while exposure to FCCP reflects maximal oxidation of FAD.

Figure 5. Measurements of the mitochondrial reactive oxygen species (ROS) generation with DCF in intact ventricular myocytes

Top, from left to right: confocal images of ventricular myocytes loaded with ROS-sensitive dye DCF (excitation 488 nm, emission 510–525 nm) before (cntrl) and after exposure to hydrogene peroxide (H₂O₂; 1 – 100 nM). Bottom, time course of DCF fluorescence in response to 1, 10 and 100 nM H₂O₂.

Figure 6. Measurements of mitochondrial NO production with DAF-2

A, from left to right: confocal images of a ventricular myocyte loaded simultaneously with DAF-2 (green, emission 510–525 nm) before (cntrl) and after cell permeabilization with digitonin, and MitoTracker Red (red, emission >590 nm). The right panel shows colocalization of DAF-2 and MitoTracker Red represented by shades of yellow. B, confocal images of a permeabilized DAF-2 loaded cell under unstimulated conditions ([Ca²⁺]_em = 0.1 μM), after an increase in [Ca²⁺]_em to 2 μM, and after subsequent addition of the exogenous NO donor Sper/NO. Insets revel NO signals from individual mitochondria. C, time course of DAF-2 fluorescence recorded from a subcellular region of interest (≤40 μm²) under conditions shown in A and B. Treatment with the NO scavenger hemoglobin (Hb, 10 μM) prevented the Ca²⁺- and Sper/NO-induced increase in DAF-2 fluorescence. (From: Dedkova and Blatter, The Journal of Physiology 2009; with permission).