Introduction

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By their nature, cardiac cells are exquisitely sensitive to oxidative stress. They die when unable to compensate adequately for the changed redox conditions caused by ischemia/reperfusion. The apoptotic cell death in cardiac tissue has been associated with oxidative stress and linked to the mitochondrial pathway (Bialik et al., 1999). This pathway involves depolarization of the mitochondrial membrane and opening of the permeability transition pore (PTP) (Kroemer and Reed, 2000). The resulting channel allows the passage of molecules 1.5 kDa (Bernardi et al., 1994; Kroemer and Reed, 2000). The released cytochrome c (Li et al., 1998), apoptosis inducing factor (Susin et al., 1996), Smac (Du et al., 2000), and procaspases (Susin et al., 1999) initiate the execution process that seals the fate of the cells.

Mitochondria are firmly integrated into the cellular control circuits and occupy a central position in cell survival/cell death decisions (Petit et al., 1996; Wei et al., 2001). The afferent signals affecting mitochondrial function are not fully understood. Among others they seem to include reactive oxygen species (ROS) and reactive nitrogen species as redox triggers of mitochondria, as well as activators of a number of upstream signaling molecules that are recruited to mitochondria. The subsequent generation of ROS may constitute a feed-forward loop to monitor the redox microenvironment and to initiate appropriate metabolic and bioenergetic responses (Vander Heiden et al., 2001). Although these are normal adjustments of healthy cells, ROS can also, in the extreme, initiate the suicide program. This is so because during oxidative stress mitochondria become susceptible to damage by ROS (Konno and Kako, 1992; Korshunov et al., 1997; Scanlon and Reynolds, 1998). The loss of membrane potential, _m, is thought to lead to the uncoupling of the respiratory chain. The consequent leakage of superoxide anions and hydrogen peroxide from the electron transport chain presumably amplifies the sudden redox stress until damage becomes irreversible.

Cells possess a number of defensive strategies to counter redox stress. These include vitamin A, although whether the mechanism is by pure antioxidant action or involves regulatory interactions with certain kinase signal pathways, as suggested by us (Hoyos et al., 2000; Imam et al., 2001), remains an open question. Because virtually all mammalian cells require vitamin A, we have investigated whether it plays a role in mitochondrial damage repair in cardiomyocytes. This assumption is in line with findings from our laboratory that retinol and its hydroxylated metabolites promote cell survival (O'Connell et al., 1996). We have shown that numerous cell types grow in culture as long as they have an adequate supply, but die of apoptosis when depleted of retinoids. Without nutritional replenishment the intracellular supply lasts on average 2 to 3 days (Eppinger et al., 1993). AR is a naturally occurring retinoid that antagonizes vitamin A action by displacing the latter from common binding sites in the regulatory domain of serine/threonine kinases (Hoyos et al., 2000). Therefore, when added exogenously AR produces an almost instant state of vitamin A deficiency, with the same consequences as those observed after nutritional vitamin A depletion, i.e., growth arrest and apoptosis. The proapoptotic capacity of AR has been connected to the production of reactive oxygen species (Chen et al., 1999). In this regard, mitochondria could be both a target of ROS and a powerful generator of ROS (Ambrosio et al., 1993). These considerations form the basis for the study presented here on the regulation of mitochondria by retinoids. We report that, depending on their chemical nature, retinoids can either ameliorate or exacerbate stress-related damage to mitochondria in cardiomyocytes.

	Materials and Methods

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Materials. AR was synthesized as described previously (Buck et al., 1993), purified by reversed phase high-performance liquid chromatography, and stored in methanol under argon at 70°C. All-trans-retinol was purchased from Sigma-Aldrich (St. Louis, MO) and phalloidin-BODIPI and 5,5',6,6'-tetrachloro-1,1',3,3'-tetraethylbenzimidazolylcarbocyanine iodide (JC-1) were from Molecular Probes (Eugene, OR). Electrophoresis reagents were from Bio-Rad (Cambridge, MA), cell culture reagents were from Invitrogen (Carlsbad, CA), and antibodies plus other reagents were from Sigma-Aldrich.

Cell Culture. Heart cells, cardiomyocytes and fibroblasts, were isolated from 2- to 3-day-old Sprague-Dawley rat pups, as described previously (Shubeita et al., 1992). Briefly, hearts were removed, stripped of atria and large vessels, minced, and digested with collagenase/pancreatine. For some experiments, myocytes were enriched by differential attachment to plastic plates. Otherwise, a purified population of cardiomyocytes was obtained by Percoll gradient centrifugation. Cardiac myocytes were cultured in 4:1 Dulbecco's modified Eagle's medium 199 containing 10% horse serum, 5% fetal calf serum, and antibiotics (100 units/ml penicillin, 100 µg/ml streptomycin). The cells were used within 6 days. Under these experimental conditions, most cells were spontaneously beating. For retinol depletion the cells were serum-starved for 72 h, as described previously (Eppinger et al., 1993). To replenish these cells with AR or retinol cultures were supplemented with the respective retinoids at 2 µM for 30 min at 37°C immediately before the experiment.

Monitoring of Mitochondrial Membrane Potential. The dual-emission, potential-sensitive probe JC-1 was loaded onto the cells 20 min before the experiment. The loaded cells were washed with PBS. Images were acquired on an LSM510 system (Carl Zeiss, Thornwood, NY). An argon laser with the 488-nm line was used for excitation. JC-1 exists as a monomer with emission maximum at 510 nm at low membrane potential, whereas at higher potentials it forms red fluorescent "J-aggregates" with emission maximum at 590 nm. JC-1 fluorescence was captured in the range of 505 to 545 nm for the monomeric form and in the range of 560 to 635 nm for the aggregate. The focal plane was set close to the nuclear center. Fluorescence intensity was calculated using MetaMorph software (Universal Imaging Corporation, Downingtown, PA). Image projections were created with the LSM510 software from individual optical sections of 1.6-mm focal depth. Images were taken from the same field at indicated time points. J-aggregate fluorescence intensity was used as a measure of mitochondrial membrane potential.

Filamentous Actin Staining and Fluorescence Microscopy. F-actin staining was performed as described previously (Korichneva and Hammerling, 1999) with minor modifications. The treated cells were fixed for 15 min with 3% paraformaldehyde in PBS. Cells were then washed with PBS and quenched in 0.1 mol/l glycine in PBS for 15 min. After an additional wash the cells were permeabilized with 0.2% Triton X-100 (w/v) in PBS containing 1% bovine serum albumin (BSA) for 10 min, and nonspecific binding sites were blocked in 2% BSA. The cells were then treated for 30 min with 0.5 µM BODIPI-conjugated phalloidin. All the above-mentioned incubations were performed at room temperature. For inspection by fluorescence microscopy, cell samples were mounted on glass slides in Vectashield (Vector Laboratories, Burlingame, CA) medium.

Isolation of Mitochondria. Hearts from adult mice were quickly removed from anesthetized animals. The organs were minced, washed, and homogenized in a Dounce glass homogenizer in 10 mM HEPES-KOH/1 mM EGTA buffer (pH 7.5) containing 250 mM sucrose and supplemented with protease and phosphatase inhibitors, 1 mM phenylmethylsulfonyl fluoride, 10 µg/ml leupeptin, 1 mM ortho-vanadate, and 20 mM NaF. The homogenates were spun down for 10 min at 2000g to discard the myofilaments. The supernatant was overlaid on 0.75 M sucrose in HEPES buffer and centrifuged for 30 min at 10,000g. Microscopy showed that the pellet constituted mostly of mitochondria. The mitochondrial pellet was resuspended in HEPES buffer for FACS analysis or treated as indicated and supplemented with 4× Laemmli electrophoresis buffer and boiled for 5 min.

FACS Analysis. Analytic flow cytometric measurements of membrane potential () of isolated mitochondria were performed using a FACScan flow cytometer with argon laser excitation at 488 nm. Mitochondria were gated using scatter parameters in the highest side scatter zone. Control treatment with FCCP confirmed the gating fully on the mitochondrial population. Green, orange, and red fluorescence intensities were detected in Fl1, Fl2, and FL3 filter sets, respectively. Ten thousand mitochondria in each sample were analyzed. The values of fluorescence intensities from the gated population were presented as a histogram. The mean values of orange fluorescence intensity were used as a measure of the value.

Electrophoresis and Western Blot Analysis. The released cytochrome c was separated on 10% SDS-polyacrylamide gel electrophoresis and electrophoretically transferred to polyvinylidene difluoride membranes, which were probed with the anti-cytochrome c antibodies. The proteins were revealed using the enhanced chemiluminescence detection system.

Statistical Analysis. Results are expressed as mean ± S.D. and compared by Student's t test.

	Results

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To monitor _m in cultured cardiac cells we used the dual-emission, potential-sensitive probe JC-1. It has been reported as the most reliable probe to reflect _m changes (Smiley et al., 1991; Mathur et al., 2000). The mitochondria of cardiac cells displayed heterogeneity with respect to their membrane potential determined by JC-1 staining. The images presented in Fig. 1 show myocytes and fibroblasts cocultured from the same heart. Cardiomyocytes were identified by their beating activity and typical morphology revealed by Nomarski optics (Fig. 1C). In these cells, the red fluorescence emitting JC-1 aggregates were localized to mitochondria showing a generally higher membrane potential than that of fibroblasts, which indicates higher metabolic activity associated with the beating function. Individual myocytes also showed heterogeneous filamentous mitochondrial structures containing both high- and low-charged mitochondria, and even regions of high and low potential within a long contiguous mitochondrion (Fig. 2).

View larger version (113K): [in this window] [in a new window]   Fig. 1.   Confocal images of cultured 6-day cells from neonatal rat hearts. Myocytes were distinguished by their beating activity in the presence of external Ca2+. The cells were loaded with dual-emission potential-sensitive fluorescent probe JC-1 for 30 min before the experiment. D, superimposed image; note that only myocytes accumulate the probe due to higher mitochondrial membrane potential. White and yellow arrows indicate myocytes and nonmyocytes, respectively. Representative from 10 cell preparations.

View larger version (56K): [in this window] [in a new window]   Fig. 2.   High-magnification confocal image of a single cardiomyocyte loaded with dual-emission potential-sensitive fluorescent probe JC-1. C, superimposed image. The cell displays clusters of highly charged mitochondria (arrow) equally distributed among organelles with green fluorescence. Red and green fluorescences together correspond to Mitotracker Red staining (data not shown). Representative from 10 cell preparations.

Taking mean red fluorescence as a measure of _m, we investigated how retinoids and retinol deficiency affect this parameter. The state of vitamin A deficiency can be reached by serum deprivation in the presence of retinoid carrier bovine serum albumin or by replacement of retinol from the binding sites by antagonist anhydroretinol (Eppinger et al., 1993). Retinoids, general components of serum, were extracted from the cells in serum-free medium repartitioning to albumin carrier present in the medium. Anhydroretinol induced a rapid decrease in cardiac _m, noticeable within minutes. The effect was signaled by a distinct decrease in red fluorescence of the aggregated JC-1 and a simultaneous increase in the green fluorescence emitted by monomeric JC-1 (Fig. 3A). The range of fluorescence intensity changes computed from cell images is displayed by intensity-coded plots. The drop in the intensity of red fluorescence was 2- to 3-fold. In a given cell, some mitochondria were depolarized, whereas others were not. A similar heterogeneity was reported in staurosporine-induced apoptosis in HL-60 cells (Salvioli et al., 2000). The doses corresponded to the ones that triggered morphological changes and cell death in many cell types (O'Connell et al., 1996; Korichneva and Hammerling, 1999) (Fig. 3B). This effect was specific for AR because retinol at similar concentrations did not affect basal _m values (Fig. 3C). Like in functional assays, retinol prevented the damage caused by AR (O'Connell et al., 1996; Korichneva and Hammerling, 1999). The mean red fluorescence corresponding to _m values were statistically analyzed as shown in Fig. 3D.

View larger version (31K): [in this window] [in a new window]   Fig. 3.   AR effect on the mitochondrial membrane potential. A, confocal images and intensity-coded plots of neonatal rat cardiomyocytes loaded with potential sensitive fluorescent probe JC-1. AR (2 µM) was added to the cells 4 min after the beginning of the experiment when fluorescent signal was stabilized. The cells were scanned at 4-min intervals. As potential drops, red fluorescence decreases, whereas green fluorescence increases. A typical of 15 experiments. B, dose dependence of AR effect on cardiac m. C, confocal images of the cells pretreated with 2 µM retinol. Red fluorescence of aggregates is shown. D, bar graph summarizing retinol and AR effects.

Oxidative treatment with hydrogen peroxide, like AR, depolarized cardiac mitochondrial membrane within minutes. To test the hypothesis that retinoids were linked to the observed ROS effects on mitochondria, cardiomyocytes were depleted of retinoids by prolonged culture in serum-free medium. One set of cultures was then replenished with a physiological concentration of retinol (2 µM), whereas a second set was maintained without retinol reconstitution. A more severe state of vitamin A deficiency was induced in a third set by exposure to 2 µM anhydroretinol for 30 min before oxidative stress induction by 100 µM hydrogen peroxide and monitoring the membrane potential over time. The results (Fig. 4) show that mild oxidative stress caused a precipitous drop in the membrane potential of vitamin A-deficient cardiomyocytes within 2 min of hydrogen peroxide administration, whereas cardiomyocytes with physiological vitamin A content were largely inured to oxidative stress. Cultures treated with AR for 30 min displayed the phenotype of oxidatively stressed cells with depolarized mitochondria membranes that did not suffer further damage by hydrogen peroxide exposure. FCCP (20 µM), a universal uncoupler of the electron transport chain, caused prompt (with 1 min) depolarization of vitamin A-sufficient mitochondria, similar in magnitude to anhydroretinol treatment alone, or to oxidative stress-induced depolarization of vitamin A-depleted cultures.

View larger version (16K): [in this window] [in a new window]   Fig. 4.   Time course of depolarization of cardiac m in response to H2O2 measured in primary culture of neonatal rat cardiomyocytes loaded with potential sensitive fluorescent probe JC-1. The cells had been retinol-depleted by culturing overnight in serum-free medium containing BSA and growth factors. Untreated, or reconstituted with retinol (ROL), or AR cardiomyocytes were then loaded with JC-1 and analyzed by confocal microscopy. m relative values correspond to red fluorescence intensity of JC-1. H2O2 induces rapid loss of m. Retinol protects mitochondrial membrane, whereas AR on its own dramatically decreases m. Values are mean ± S.D. from five measurements.

The rapidity with which cardiomyocyte mitochondria suffered damage by AR was surprising. To test whether other known forms of negative impact occurred with similar speed we monitored the morphological changes over time. We found that cultured cardiomyocytes treated with 2 µM AR developed morphological changes at the rate similar to that previously described for NIH 3T3 fibroblasts (Korichneva and Hammerling, 1999). They displayed volume increase, F-actin reorganization, membrane blebbing, and death in 18 h (Fig. 5). Actin reorganization as a hallmark of AR-triggered apoptosis was prominent in cardiomyocytes. The thick, short stick-like structures in the cytoplasm occurred after 2 h of treatment, changing later to strong F-actin accumulation in ruffle-like surface membrane structures. Still later, the actin cytoskeleton dissolved entirely and the cells shrank in size. Nontreated cells did not demonstrate any changes in F-actin morphology during cell culture (cells not shown).

View larger version (89K): [in this window] [in a new window]   Fig. 5.   Fluorescence images of cultured cells. Two-day neonatal mouse cardiac myocytes stained with BODIPY 581/591-phalloidin. A, control; cells were cultured in the presence of 1 µM AR for 6 (B), 8 (C and D) 12 (E), and 18 (F) h. Arrows indicate transformation of F-actin structures.

The fact that a single pharmacological agent (i.e., AR) caused pleotropic effects, such as mitochondrial damage and microfilament reorganization, is more in line with the model of disruption of the regulatory signals than with direct membrane toxicity. To gain further evidence in favor of this concept, we tested the pharmacological inhibition of the effect by acting on PTP. It is known that the classical PTP opening depends on the Ca²⁺ concentration (Ichas et al., 1997) and is sensitive to cyclosporin A binding (Crompton, 1999). The series of images shown in Fig. 6, A and B, demonstrates _m changes induced by AR or hydrogen peroxide in cardiomyocytes pretreated with cyclosporin A. Similarly, Ca²⁺ involvement was studied using the Ca²⁺-chelating agent BAPTA-AM. MetaMorph image analysis yielded time courses of red fluorescence intensity changes in pharmacologically pretreated cells (Fig. 6, C and D). We found that the AR-induced _m depolarization was blocked by cyclosporin A and BAPTA. In comparison with AR, the peroxide-triggered depolarization was only partially reversed by Ca²⁺ chelation with BAPTA and was completely insensitive to cyclosporin A pretreatment. It seems that AR acts on the classical PTP, whereas oxidation opens cyclosporin A-insensitive channels. Using a similar experimental approach, our results have been corroborated with H₂O₂-treated rat forebrain neurons, suggesting either a cyclosporin A-insensitive form of permeability transition or a separate mechanism (Scanlon and Reynolds, 1998).

View larger version (69K): [in this window] [in a new window]   Fig. 6.   Anhydroretinol and reactive oxygen affect permeability transition by different mechanisms. Series of confocal images of JC-1-loaded cardiomyocytes treated with AR (A) or hydrogen peroxide (B) after preincubation with cyclosporin A. C and D, time courses of the changes in red fluorescence of JC-1-loaded cardiomyocytes pretreated with the inhibitors of PTP, cyclosporin A, and BAPTA-AM. One of three similar experiments.

We further investigated whether retinoids influenced mitochondria directly. Intact mitochondria were isolated from whole heart using differential centrifugation in a sucrose gradient. The purified organelles were then loaded with JC-1 and analyzed by flow cytometry. According to Cozarizza et al. (1996) orange fluorescence JC-1 signals in the FACS analysis of isolated rat liver mitochondria and _m values measured by a tetraphenylphosphonium-selective electrode are linearly correlated. The Fig. 7A shows a histogram of JC-1 fluorescence intensity recorded in channel FL2, characteristic of intact mitochondria, indicating, however, heterogeneity in membrane potential among individual mitochondria. This heterogeneity was expected taking into account different sizes of isolated mitochondria. Also, it was similar to the distribution of fluorescence observed in intact cells. The orange fluorescence intensity decreased within 30 s after treatment with 2 µM anhydroretinol. Exposure to peroxide caused a very similar transition. The respective time courses of _m depolarization are shown in the Fig. 7B. Two control treatments were performed to relate _m decrease to the effects observed at mitochondrial permeability transition pore opening or uncoupling of oxidative phosphorylation, corresponding to treatment with the ligand of the benzodiazepine receptor, atractyloside and FCCP.

View larger version (17K): [in this window] [in a new window]   Fig. 7.   Direct effects of AR on isolated mitochondria from mouse hearts. A, overlaying histograms for FL2 orange fluorescence of intact mitochondria treated with different drugs. As demonstrated in primary cultures m is heterogeneous. B, time course shows that m decreases within 30 s after 2 µM AR addition to intact mitochondria. H2O2 at 100 µM induces similar pattern of change in m. The 2-min effect of atractyloside is shown for comparison. The fluorescent signal nearly disappeared in the mitochondria treated with FCCP. One of four similar experiments is shown. C, cytochrome c release from isolated mouse heart mitochondria was measured by Western blot with anti-cytochrome c antibodies. The time points represent periods of AR treatment.

The opening of PTP leads to mitochondria swelling, membrane rupture, and release of substances from the matrix. Because AR has been be causally linked to triggering the apoptotic program it was of interest to determine whether cytochrome c was released from isolated mouse heart mitochondria as a result of contact with AR. Analysis of Western blots with anti-cytochrome c antibodies confirmed the release of cytochrome c (Fig. 7C) after 15 min of AR treatment.

	Discussion

Top Abstract Introduction Materials and Methods Results Discussion References

We report that mitochondria of neonatal rat cardiomyocytes are regulated by retinoids. First, we confirmed and extended the observation that vitamin A depletion rendered mitochondria highly susceptible to oxidative stress. Presence of physiological concentrations of retinol prevented this effect. Second, blockade of vitamin A action by AR, which is known to displace the former from its cognate binding site on kinases, mimicked the results of oxidative stress. Like in functional experiments, retinol was able to prevent the AR effect. These data support the functional antagonism between retinol and AR previously demonstrated by us. The AR-induced _m depolarization was dependent on Ca²⁺ and sensitive to cyclosporin A, ruling out direct membrane effects.

Despite being the first vitamin discovered the full range of biological activities of vitamin A remains to be defined in general, and in cardiac tissue in particular. It has been noted that during myocardial infarction increased amounts of vitamin A are mobilized from the liver to the heart (Palace et al., 1999). This is remarkable because the body maintains strict steady-state vitamin A levels at other times. It is conceivable that vitamin A itself plays a role in the repair of infarct damage. Retinoids are known to act as survival factors of cardiac cells, because retinol reduced the incidence of heart disease and protected the heart from inflammation, oxidative damage, and degeneration (Swamidas et al., 1999). Conversely, chronic vitamin A deficiency in rats decreased the respiratory activity of their heart mitochondria (Estornell et al., 2000). Several arguments can be made to consider mitochondria as potential targets for protective intervention during reperfusion of the ischemic heart. Our findings that mitochondria constitute one of the multiple non-nuclear functional targets for retinoids are entirely congruent with the previous studies that had identified retinol as potent cell survival factor. Protein kinase C and mitogen-activated protein kinase pathways in a variety of cell types were identified as the intracellular targets of retinol, the very same pathways operating in stress management of cardiac cells.

A number of signaling and adapter proteins associated with mitochondrial membranes are subject to redox control. They include the components of the Raf/mitogen-activated protein kinase pathway (Wang et al., 1994), c-Jun NH₂-terminal kinase/stress-activated protein kinase pathway (Tournier et al., 2000), and protein kinase C (Majumder et al., 2000). In that respect, mitochondria can be viewed as signal integration and amplification stations. Some of these signaling proteins are characterized as retinoid binding proteins. cRaf and protein kinase C are among them (Hoyos et al., 2000; Imam et al., 2001). It is now well accepted that numerous non-nuclear targets exist for retinoids, even for retinoic acid, which is the classical nuclear receptor ligand (Marchetti et al., 1999). Isolated mitochondria behaved much like those of intact cells. Hence, it is possible that AR and retinol target a protein within the mitochondria rather than upstream regulatory factors. However, such molecular targets and binding sites for retinoids on mitochondria remain unknown.

The mitochondrial effects of retinoids are in line with their non-nuclear function addressed by us in a previous study (O'Connell et al., 1996). It corroborates recent finding that the synthetic retinoid 6-{3-(1-adamantyl)-4-hydroxyphenyl}-2-naphtalene carboxylic acid can trigger apoptosis through a mitochondrial pathway independently of nuclear involvement (Marchetti et al., 1999; Dawson et al., 2001). Similarly, mitochondrial permeability transition was a central feature of N-(4-hydroxyphenyl)retinamide-induced apoptosis in transformed cells (Hail and Lotan, 2000). Retinoic acids, e.g., all-trans-, 9-cis-, and 13-cis-retinoic acids, were also capable of inducing the membrane permeability transition (Rigobello et al., 1999). The ability of retinol to protect, at least in part, mitochondria from oxidative damage characterizes it as a component of general cellular defenses. Retinol and appropriate derivatives can be expected to find clinical applications for stabilization of the cellular redox state after ischemia/reperfusion and other conditions of oxidative stress.