From the Consiglio Nazionale delle Ricerche Unit for the Study of
Biomembranes, and the Departments of Biological Chemistry
and
Biomedical Sciences, University of Padova, Viale Giuseppe
Colombo 3, I-35121 Padova, Italy, and the ¶ Department of
Biochemistry and Molecular Biology, University of Bari, Via Ovabona 4, I-70125 Bari, Italy
Received for publication, July 31, 2000, and in revised form, October 27, 2000
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The opening of the mitochondrial permeability
transition pore (PTP) has been suggested to play a key role in various
forms of cell death, but direct evidence in intact tissues is still lacking. We found that in the rat heart, 92% of NAD+
glycohydrolase activity is associated with mitochondria. This activity
was not modified by the addition of Triton X-100, although it was
abolished by mild treatment with the protease Nagarse, a condition that
did not affect the energy-linked properties of mitochondria. The
addition of Ca2+ to isolated rat heart mitochondria
resulted in a profound decrease in their NAD+ content,
which followed mitochondrial swelling. Cyclosporin A(CsA), a PTP
inhibitor, completely prevented NAD+ depletion but had no
effect on the glycohydrolase activity. Thus, in isolated mitochondria
PTP opening makes NAD+ available for its enzymatic
hydrolysis. Perfused rat hearts subjected to global ischemia for 30 min
displayed a 30% decrease in tissue NAD+ content, which was
not modified by extending the duration of ischemia. Reperfusion
resulted in a more severe reduction of both total and mitochondrial
contents of NAD+, which could be measured in the coronary
effluent together with lactate dehydrogenase. The addition of
0.2 µM CsA or of its analogue MeVal-4-Cs (which does not
inhibit calcineurin) maintained higher NAD+ contents,
especially in mitochondria, and significantly protected the heart from
reperfusion damage, as shown by the reduction in lactate dehydrogenase
release. Thus, upon reperfusion after prolonged ischemia, PTP opening
in the heart can be documented as a CsA-sensitive release of
NAD+, which is then partly degraded by glycohydrolase and
partly released when sarcolemmal integrity is compromised. These
results demonstrate that PTP opening is a causative event in
reperfusion damage of the heart.
Depending on the duration and severity of myocardial ischemia,
reperfusion can result in either recovery of contractile function or
rapid transition toward tissue necrosis (for review see Refs. 1-3).
Paradoxically, both events require coupled mitochondrial respiration
(4). Indeed, cyanide (5) or 2,4-dinitrophenol (6) largely reduce the
release of intracellular enzymes, the marker of cell death induced by
postischemic reperfusion. However, after more than 25 years, the
specific mechanisms underlying these phenomenological observations have
yet to be elucidated.
A large body of experimental evidence suggests that a suboptimal
mitochondrial function could produce low levels of ATP, which in the
presence of even a modest rise in [Ca2+]i might
cause hypercontracture in isolated cardiomyocytes (7) and sarcolemma
rupture in intact hearts (8, 9). Such a sequence of events could be set
in motion by the opening of the mitochondrial
PTP,1 a high conductance
channel located in the inner mitochondrial membrane (10). The open
probability of this channel is regulated by several factors including
mitochondrial membrane potential difference ( PTP opening is likely to alter several metabolic pathways linked to
energy metabolism, and results from a classic study suggest that
NAD+ catabolism may be one of them (17). Indeed, the
content of mitochondrial pyridine nucleotides was drastically reduced
upon Ca2+ addition, a condition that could have induced PTP
opening. Here we show that in RHM opening of the PTP causes the release
of mitochondrial NAD+ followed by its hydrolysis by a
CsA-insensitive NAD+ glycohydrolase localized outside the
matrix space. Furthermore, we document that in the intact heart during
postischemic reperfusion mitochondrial NAD+ content is
severely decreased in a process that is largely reduced by PTP
inhibitors, suggesting that the hydrolysis of mitochondrial NAD+ directly reflects PTP opening in situ. The
maintenance of mitochondrial NAD+ is thus associated with a
significant protection from myocyte death, indicating that the PTP
plays a key role in this process.
Perfusion of Isolated Hearts--
All aspects of animal care and
experimentation were performed in accordance with the Guide for the
Care and Use of Laboratory Animals, published by the National
Institutes of Health (NIH Publication No. 85-23, revised in 1996), and
the national laws of Italy concerning the care and use of laboratory
animals and were approved by the Ethical Committee of the University of
Padova. Hearts excised from male Wistar rats (weighing 180-200 g) were
perfused by the nonrecirculating Langendorff technique as previously
described (18). Hearts were not stimulated, and the flow was maintained at 12 ml/min throughout all the perfusion protocols except during ischemia, which was induced by the complete abolition of coronary flow
for periods ranging from 30 to 90 min. Left ventricular wall temperature was maintained at 36-37 °C irrespective of coronary flow by suspending the heart in a water-jacketed chamber.
Mitochondria Isolation and Swelling Assay--
Mitochondria were
isolated by conventional procedures of differential centrifugation
(19). Freshly excised rat hearts were homogenized by Ultra-Turrax in a
medium containing 0.18 M KCl, 10 mM EDTA, 0.5%
fatty acid-poor bovine serum albumin, 10 mM Hepes, pH 7.4. To remove EDTA and albumin, mitochondrial pellets were washed twice
with 0.18 M KCl, 10 mM Hepes, pH 7.4 (18). In a separate set of experiments aimed at characterizing the localization of
the mitochondrial NAD+ glycohydrolase, mitochondria were
isolated after the incubation of the whole tissue homogenate with
Nagarse as previously described (20).
Mitochondrial swelling was monitored as the changes in absorbance at
540 nm as previously described (21). Incubations were carried out at
25 °C with 0.25 mg of mitochondrial protein/ml in the RB medium,
0.25 M sucrose, 10 mM Tris-Mops, 0.05 mM EGTA, pH 7.4, 5 mM pyruvate, 5 mM malate, and 1 mM Pi-Tris. PTP
opening was induced by the addition of 0.25 mM
Ca2+.
Metabolite and Enzyme Assays--
NAD+ and CoASH
were measured after perchloric acid extraction. To achieve this, the
hearts were freeze-clamped with aluminum tongues cooled in
liquid nitrogen, and 0.3 g of freeze-clamped tissue (stored at
In neutralized HClO4 extracts, NAD+ was
determined fluorometrically with alcohol dehydrogenase (22), and
CoASH was assayed with an enzymatic cycling method (23).
The mitochondrial hydrolysis of endogenous FAD was measured as the
increase in fluorescence (excitation and emission wavelengths at 450 and 520 nm, respectively) in the supernatant of mitochondria pelleted
after the various incubation protocols (24). The fluorescence increase
is caused by the release of the hydrolytic products, namely flavin
mononucleotide and riboflavin.
The activity of mitochondrial NAD+ glycohydrolase was
measured by monitoring the enhancement in fluorescence emission caused by the hydrolysis of
During postischemic reperfusion, 1-ml samples of the effluent were
collected at 1-min intervals for the first 5 min and at 5-min intervals
until the end of the reperfusion protocol. LDH activity was measured by
means of a classic procedure (26). Neutralized HClO4
extracts of the effluents were used for NAD+ assay. Data
are presented as cumulative values for the entire reperfusion period.
Lactate dehydrogenase, NAD+, and NADH were purchased from
Roche Molecular Biochemicals. All other enzymes and chemicals were purchased from Sigma and were of the highest available grade. CsA and
MeVal-4-Cs were generous gifts of Novartis (Basel, Switzerland).
The initial aim of our study was to determine whether in isolated
mitochondria NAD+ hydrolysis could be related to PTP
opening. In the experiments of Fig. 1,
the addition of 0.2 mM Ca2+ to RHM (0.2 mg of
mitochondrial protein × ml
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
m),
Ca2+, matrix pH, and CsA, a high affinity inhibitor (11).
PTP opening causes a Ca2+-dependent increase of
mitochondrial permeability to ions and solutes with molecular masses of
up to 1500 Da, matrix swelling, and mitochondrial
deenergization. Several studies performed on isolated cardiomyocytes
(12-14) and perfused hearts (15, 16) support the idea that PTP opening
might be pivotal in determining the transition of the ischemic damage
to the irreversible phase. However, the role of PTP is not yet defined
due to the difficulty of assaying its opening in situ.
MATERIALS AND METHODS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
70 °C) was ground and mixed thoroughly with 2 ml of 14% (v/v)
HClO4. After thawing at 4 °C, this mixture was homogenized and centrifuged as previously described (18). In the case
of isolated mitochondria, 0.1 ml of 21% (v/v) HClO4 was added to 1 mg of protein/ml suspensions.
-NAD (25). The assay was carried out by adding
RHM (0.2 mg of protein/ml of RB medium) with 200 µM
-NAD, which was found to saturate the NADase activity. Fluorescence measurements were performed using a PerkinElmer LS5 spectrofluorometer. The excitation wavelength was set to 310 nm. Fluorescence emission was
followed at 410 nm. The concentration of
-NAD was determined by the
conversion of
-NAD to
-NADH using the alcohol dehydrogenase reaction and assuming a molar extinction coefficient for
-NADH of
6.2 × 106 cm2/mol at 340 nm. The
fluorescence changes produced by mitochondria were calibrated by using
a standard curve produced by incubating
-NAD (at concentrations
ranging from 1 to 50 µM) with excess amounts of NADase
from Neurospora crassa (25).
RESULTS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
1) induced a rapid
decrease of absorbance at 540 nm, which is indicative of swelling.
Under these conditions the NAD+ content was reduced to less
than 20% of control values within 20-30 min, whereas FAD and CoASH
were not hydrolyzed (results not shown). Fig. 1 clearly documents that
the fall in mitochondrial NAD+ content began after the
completion of mitochondrial swelling and that both processes were
largely prevented by pretreatment with 0.2 µM CsA,
suggesting that NAD+ disappearance was related to PTP
opening. The decrease in NAD+ content was also prevented by
5 mM nicotinamide, an inhibitor of NAD+
glycohydrolase that did not affect either the rate or the extent of
mitochondrial swelling (Fig. 1). On the other hand, neither NAD+ depletion nor PTP opening was modified by the addition
of AMP, which inhibits nucleotide pyrophosphatase (27). Superimposable results on the relationship between PTP opening and NAD+
decrease could be obtained in rat liver mitochondria (results not
shown).
View larger version (13K):
[in a new window]
Fig. 1.
The decrease in NAD+ content is
caused by the opening of the PTP in isolated rat heart
mitochondria. A, mitochondrial swelling was monitored
as the decrease in light absorbance at 540 nm
(A540). PTP opening was induced by the addition
of 0.25 mM Ca2+ to untreated RHM (0.25 mg of
protein/ml) (trace a) or mitochondria added with the
following compounds 1 min before Ca2+: 1 µM
cyclosporin A (b), 5 mM nicotinamide
(c), and 1 mM AMP (d). B,
1-ml aliquots of mitochondria (1 mg of protein/ml) were withdrawn at
the indicated times, and their NAD+ contents were measured.
Trace letters refer to the same treatments described for the
experiments of A.
Fig. 2 summarizes a series of experiments
performed to characterize the activity and determine the location of
NAD+ glycohydrolase. RHM displayed a NAD+
hydrolytic activity of 0.92 nmol/min/mg of mitochondrial protein. Considering a mitochondrial content of 55 mg of proteins/g of wet heart
tissue (28), the mitochondrial NAD+ glycohydrolase activity
represents 92% of that measured in the whole tissue (53 milliunits/g
wet weight). Because the NAD+ hydrolytic activity
was totally inhibited by nicotinamide and not affected by AMP (data not
shown), we attribute this activity to NAD+ glycohydrolase.
The rate of -NAD hydrolysis was not modified by the addition of
Triton X-100 to well coupled RHM (Fig. 2). Furthermore, the incubation
of RHM with a serine protease Nagarse, frequently used to increase the
yield of mitochondrial extraction from heart tissues (20, 29), produced
a time-dependent reduction of NAD+
glycohydrolase (Fig. 2) without affecting the respiratory control (data
not shown) and, thus, the integrity of the inner mitochondrial membrane. These results demonstrate that the NAD+
glycohydrolase activity is not localized within the mitochondrial matrix and indicate that NAD+ hydrolysis occurs outside
this compartment. Importantly, the NAD+ glycohydrolase
activity was not affected by CsA (Fig. 2). From these results, we
conclude that PTP opening in isolated mitochondria causes the release
of intramitochondrial NAD+, which then becomes a substrate
for the glycohydrolase located outside the matrix space.
|
The possible occurrence of tissue and mitochondrial NAD+
hydrolysis was then investigated in isolated rat hearts subjected to
ischemia and postischemic reperfusion. Figs.
3 and 4
document the changes of the tissue contents of NAD+ in
perfused hearts and isolated mitochondria. After 30 min of ischemia,
the NAD+ content was decreased by 30% and did not show
further changes as the ischemic period was extended (Fig. 3). On the
other hand, a reperfusion period of 20 min resulted in a severe
decrease of tissue NAD+ contents (Fig. 4). Fig. 4 also
shows that the mitochondrial NAD+ was depleted almost
completely, suggesting that these organelles contribute to a large
extent to the overall changes in cellular NAD+. It is
noteworthy that the decrease of NAD+ content was largely
prevented by 0.2 µM of both CsA and its analogue MeVal-4-Cs. A similar loss of tissue and mitochondrial NAD+
was also observed in aerobic hearts exposed to the Ca2+
paradox protocol (readmission of Ca2+ in the perfusion
buffer after 10 min of perfusion in the absence of Ca2+),
which is known to induce a massive intracellular Ca2+
overload (30) (results not shown).
|
|
We next tested whether NAD+ was released from the cells.
The experiments of Fig. 5 show that
NAD+ was detected in the coronary effluent together with
LDH and that both events were largely reduced by both CsA and
MeVal-4-Cs. Indeed, in hearts perfused with these PTP inhibitors, the
preservation of the mitochondrial NAD+ pool was associated
with a significant decrease of LDH release in the coronary effluent
(Fig. 5).
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The present results establish a causal link between the opening of the PTP and hydrolysis of mitochondrial NAD+ both in isolated organelles and intact hearts, and they document the relevance of the PTP in the injury of the heart produced by postischemic reperfusion.
Mechanism of NAD+ Hydrolysis by Mitochondria--
In
our protocols, the disappearance of mitochondrial NAD+ is
clearly the consequence of its release in the intermembrane space where
it becomes the substrate of glycohydrolase. The presence of
NAD+ glycohydrolase within the matrix space could be
excluded because (i) treatment with Nagarse completely abolished
endogenous NAD+ hydrolysis under conditions where the inner
membrane was demonstrably unaffected, and (ii) the addition of Triton
X-100 did not increase the hydrolysis of added -NAD by intact
isolated mitochondria (Fig. 2). In keeping with the results of a
previous study on liver mitochondria (31), these findings indicate that
NAD+ glycohydrolase is located in the outer membrane (24).
Thus, the inner mitochondrial membrane separates the matrix space where NAD+ is accumulated (and possibly synthesized (27)) from
the intermembrane space where NAD+ hydrolysis takes place.
The redistribution of NAD+ between these two compartments
is made possible by PTP opening because mitochondrial swelling precedes
NAD+ hydrolysis, and both processes are inhibited by CsA
(Fig. 1), which demonstrably did not affect the activity of
NAD+ glycohydrolase.
Our results strongly argue against a model where mitochondrial Ca2+ would be released upon the ADP-ribosylation of an intrinsic protein of the inner mitochondrial membrane following pyridine nucleotide hydrolysis by a Ca2+-stimulated matrix glycohydrolase (32-34). Based on the present results, we conclude that PTP opening, which induces the efflux of mitochondrial Ca2+, is the cause rather than the consequence of NAD+ hydrolysis that cannot occur within the matrix space (Fig. 2). We also note that inhibition of Ca2+ efflux by CsA was also observed by Richter et al. (35), who attributed it to the inhibition of glycohydrolase by CsA. Our experiments demonstrate that this is not the case (Fig. 2) and conclusively prove that NAD+ hydrolysis is only possible after PTP opening. Given that in postischemic reperfusion of the heart we found a CsA-sensitive decrease of mitochondrial and total NAD+ (Fig. 4), clarification of this issue was essential and provided a mechanistic clue into the complex mitochondrial changes occurring in this pathological condition.
Role of the Permeability Transition in Postischemic Reperfusion Injury of the Heart-- In principle, mitochondrial dysfunction could be either the cause or the consequence of reperfusion injury of the heart, a complex issue that is far from having been solved. For instance, sarcolemmal rupture results in the exposure of mitochondria to the millimolar [Ca2+] of the extracellular milieu, which would cause the immediate failure of mitochondria even if they had been fully functional before this terminal event. Not surprisingly, the appearance of calcium phosphate precipitates within the matrix represents the most reliable sign of myocyte necrosis (36). The present results are thus extremely relevant because they indicate that PTP opening is a causative event rather than a consequence in the complex sequence of events linking mitochondrial failure to myocyte injury. Indeed, the loss of tissue viability was greatly reduced by CsA, and the effect could be traced to PTP inhibition because hydrolysis of mitochondrial NAD+ was fully prevented in parallel with cell protection (Figs. 4 and 5). It has to be pointed out that CsA can act through mechanisms other than PTP inhibition. In particular, CsA also inhibits calcineurin, and this effect was suggested to contribute to the protection afforded by CsA in both brain and heart ischemia (37-39). To explore the possibility that calcineurin inhibition was involved in the protective effects of CsA in our model, we also tested the cyclosporin derivative MeVal-4-Cs, which binds cyclophilins and inhibits the PTP but does not affect calcineurin activity (40, 41). The fact that MeVal-4-Cs was as protective as CsA strongly suggests that calcineurin is not involved and supports our interpretation that PTP opening plays a pivotal role in ensuing myocyte death during postischemic reperfusion.
In this scenario, the changes produced by reperfusion in viable myocytes would promote PTP opening, and this event would be followed rather than preceded by the loss of sarcolemmal integrity. Thus, mitochondria would act as transducers and amplifiers of an initial trigger provided by reperfusion, and NAD+ hydrolysis could be part of the amplification pathway. The availability of mitochondrial NAD+ could indeed result in the formation of both cyclic ADP-ribose and nicotinic acid adenine dinucleotide phosphate, which are well known promoters of Ca2+ release from the sarcoplasmic reticulum (42). It is tempting to speculate that the mitochondrial hydrolysis of NAD+ even by a fraction of the mitochondria could thus eventually induce an increase of intracellular [Ca2+], promoting the further spreading of the permeability transition to all mitochondria in the cell. This would then cause generalized mitochondrial dysfunction and establish the conditions (low ATP and high Ca2+) that precipitate irreversible contracture and sarcolemmal rupture. We also note that our results provide a mechanism for the decrease of NAD+ contents that had already been reported in the ischemic myocardium (43, 44).
Mitochondria and Pyridine Nucleotide Metabolism-- Our results also highlight the relevance of mitochondria in the cellular utilization and catabolism of pyridine nucleotides, especially in the myocardium. Indeed, we found that mitochondria are the major stores of NAD+ and possess nearly all of the NAD+ glycohydrolase activity of the cell (72 and 92% of the total, respectively). Thus, any condition leading to a major decrease in cellular NAD+ contents is probably contributed by mitochondria. It has been proposed that the activation of PARP results in the depletion of cellular NAD+ and consequently of ATP that is required for NAD+ resynthesis (45, 46). Our data indicate that the activation of PARP must be coupled with the utilization of mitochondrial NAD+, which is only made possible by PTP opening. Given the importance of both processes for cell death our results may provide a further biochemical link between mitochondrial and cellular pathways to cell death. Future studies will determine whether the same stimuli that alter DNA structure stimulating PARP activation might simultaneously promote PTP opening.
Although the conditions of the present study are designed to address
the role of mitochondrial release of NAD+ in a model of
cell death, PTP opening could underlie the bidirectional fluxes of
NAD+ through the inner mitochondrial membrane observed in
digitonin-treated cell lines (47). The idea that the inner membrane is
permeable to pyridine nucleotides contrasts with notions that are,
however, entirely based on studies with isolated mitochondria where
incubation conditions are designed to maximize energy coupling by
minimizing the chances of pore opening. We have documented transient
PTP openings in healthy cells (48), and it may well be that these openings play an unsuspected role in the physiological transport of
pyridine nucleotides.
![]() |
FOOTNOTES |
---|
* This work was supported by grants from the Consiglio Nazionale delle Ricerche and the Ministero per l'Università e la Ricerca Scientifica e Tecnologica "Il mantenimento della vitalità miocardica a discapito della necrosi" (to F. D. L.) and "Bioenergetica e Trasporto di Membrana" (to P. B.).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
§ To whom correspondence should be addressed: Dipartimento di Chimica Biologica, Viale Giuseppe Colombo 3, I-35121 Padova, Italy. Tel.: 39-049-827-6132; Fax: 39-049-807-3310; E-mail: dilisa@civ.bio.unipd.it.
Published, JBC Papers in Press, November 9, 2000, DOI 10.1074/jbc.M006825200
![]() |
ABBREVIATIONS |
---|
The abbreviations used are:
PTP, permeability
transition pore;
CsA, cyclosporin A;
-NAD, 1,N6-etheno-NAD+;
MeVal-4-Cs, N-methylvaline-4-cyclosporin;
PARP, poly(ADP-ribose)
polymerase;
RHM, rat heart mitochondria;
LDH, lactate dehydrogenase;
MOPS, 4-morpholinepropanesulfonic acid.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | Jennings, R. B., and Reimer, K. A. (1991) Annu. Rev. Med. 42, 225-246[CrossRef][Medline] [Order article via Infotrieve] |
2. | Silverman, H. S., and Stern, M. D. (1994) Cardiovasc. Res. 28, 581-597[Medline] [Order article via Infotrieve] |
3. | Di Lisa, F., Menabò, R., Canton, M., and Petronilli, V. (1998) Biochim. Biophys. Acta 1366, 69-78[Medline] [Order article via Infotrieve] |
4. | Di Lisa, F., Blank, P. S., Colonna, R., Gambassi, G., Silverman, H. S., Stern, M. D., and Hansford, R. G. (1995) J. Physiol. (Lond.) 486, 1-13[Abstract] |
5. | Ganote, C. E., Worstell, J., and Kaltenbach, J. P. (1976) Am. J. Pathol. 84, 327-350[Abstract] |
6. | Ganote, C. E., McGarr, J., Liu, S. Y., and Kaltenbach, J. P. (1980) J. Mol. Cell. Cardiol. 12, 387-408[Medline] [Order article via Infotrieve] |
7. | Miyata, H., Lakatta, E. G., Stern, M. D., and Silverman, H. S. (1992) Circ. Res. 71, 605-613[Abstract] |
8. | Ganote, C. E., and Armstrong, S. C. (1993) Cardiovasc. Res. 27, 1387-1403[Medline] [Order article via Infotrieve] |
9. | Jennings, R. B., Murry, C. E., Steenbergen, C., and Reimer, K. A. (1990) Circulation 82, 2-12 |
10. |
Bernardi, P.
(1999)
Physiol. Rev.
79,
1127-1155 |
11. | Bernardi, P., Broekemeier, K. M., and Pfeiffer, D. R. (1994) J. Bioenerg. Biomembr. 26, 509-517[Medline] [Order article via Infotrieve] |
12. | Nazareth, W., Yafei, N., and Crompton, M. (1991) J. Mol. Cell. Cardiol. 23, 1351-1354[Medline] [Order article via Infotrieve] |
13. | Duchen, M. R., McGuinness, O., Brown, L. A., and Crompton, M. (1993) Cardiovasc. Res. 27, 1790-1794[Medline] [Order article via Infotrieve] |
14. | Minezaki, K. K., Suleiman, M. S., and Chapman, R. A. (1994) J. Physiol. (Lond.) 476, 459-471[Abstract] |
15. | Griffiths, E. J., and Halestrap, A. P. (1993) J. Mol. Cell. Cardiol. 25, 1461-1469[CrossRef][Medline] [Order article via Infotrieve] |
16. | Griffiths, E. J., and Halestrap, A. P. (1995) Biochem. J. 307, 93-98[Medline] [Order article via Infotrieve] |
17. | Vinogradov, A., Scarpa, A., and Chance, B. (1972) Arch. Biochem. Biophys. 152, 646-654[Medline] [Order article via Infotrieve] |
18. |
Di Lisa, F.,
Menabò, R.,
Barbato, R.,
and Siliprandi, N.
(1994)
Am. J. Physiol.
267,
H455-H461 |
19. | Lindenmayer, G. E., Sordahl, L. A., and Schwartz, A. (1968) Circ. Res. 23, 439-450[Medline] [Order article via Infotrieve] |
20. |
Di Lisa, F.,
Fan, C. Z.,
Gambassi, G.,
Hogue, B. A.,
Kudryashova, I.,
and Hansford, R. G.
(1993)
Am. J. Physiol.
264,
H2188-H2197 |
21. |
Scorrano, L.,
Petronilli, V.,
Di Lisa, F.,
and Bernardi, P.
(1999)
J. Biol. Chem.
274,
22581-22585 |
22. | Klingenberg, M. (1985) in Methods of Enzymatic Analysis (Bergmeyer, H. U., ed) , pp. 251-271, Verlag Chemie, Weinheim, Germany |
23. | Veloso, D., and Veech, R. L. (1974) Anal. Biochem. 62, 449-450[Medline] [Order article via Infotrieve] |
24. | Barile, M., Brizio, C., De Virgilio, C., Delfine, S., Quagliariello, E., and Passarella, S. (1997) Eur. J. Biochem. 249, 777-785[Abstract] |
25. | Barrio, J. R., Secrist, J. A., and Leonard, N. J. (1972) Proc. Natl. Acad. Sci. U. S. A. 69, 2039-2042[Abstract] |
26. | Bergmeyer, H. U., and Bernt, E. (1974) in Methods of Enzymatic Analysis (Bergmeyer, H. U., ed) , pp. 607-612, Verlag Chemie, Weinheim, Germany |
27. | Barile, M., Passarella, S., Danese, G., and Quagliariello, E. (1996) Biochem. Mol. Biol. Int. 38, 297-306[Medline] [Order article via Infotrieve] |
28. | Idell Wenger, J. A., Grotyohann, L. W., and Neely, J. R. (1978) J. Biol. Chem. 253, 4310-4318[Abstract] |
29. | Palmer, J. W., Tandler, B., and Hoppel, C. L. (1977) J. Biol. Chem. 252, 8731-8739[Abstract] |
30. | Zimmerman, A. N., and Hülsmann, W. C. (1966) Nature 211, 646-647[Medline] [Order article via Infotrieve] |
31. |
Boyer, C. S.,
Moore, G. A.,
and Moldeus, P.
(1993)
J. Biol. Chem.
268,
4016-4020 |
32. |
Lotscher, H. R.,
Winterhalter, K. H.,
Carafoli, E.,
and Richter, C.
(1980)
J. Biol. Chem.
255,
9325-9330 |
33. | Richter, C., Winterhalter, K. H., Baumhuter, S., Lotscher, H. R., and Moser, B. (1983) Proc. Natl. Acad. Sci. U. S. A. 80, 3188-3192[Abstract] |
34. | Frei, B., Winterhalter, K. H., and Richter, C. (1985) Eur. J. Biochem. 149, 633-639[Abstract] |
35. | Richter, C., Theus, M., and Schlegel, J. (1990) Biochem. Pharmacol. 40, 779-782[Medline] [Order article via Infotrieve] |
36. | Shen, A. C., and Jennings, R. B. (1972) Am. J. Pathol. 67, 417-421[Medline] [Order article via Infotrieve] |
37. | Nishinaka, Y., Sugiyama, S., Yokota, M., Saito, H., and Ozawa, T. (1993) J. Cardiovasc. Pharmacol. 21, 448-454[Medline] [Order article via Infotrieve] |
38. | Morioka, M., Hamada, J., Ushio, Y., and Miyamoto, E. (1999) Prog. Neurobiol. (OXF) 58, 1-30[CrossRef][Medline] [Order article via Infotrieve] |
39. | Ruiz, F., Alvarez, G., Ramos, M., Hernandez, M., Bogonez, E., and Satrustegui, J. (2000) Eur. J. Pharmacol. 404, 29-39[CrossRef][Medline] [Order article via Infotrieve] |
40. | Zenke, G., Baumann, G., Wenger, R. M., Hiestand, P., Quesniaux, V., Andersen, E., and Schreier, M. H. (1993) Ann. N. Y. Acad. Sci. 685, 330-335[Abstract] |
41. |
Nicolli, A.,
Basso, E.,
Petronilli, V.,
Wenger, R. M.,
and Bernardi, P.
(1996)
J. Biol. Chem.
271,
2185-2192 |
42. |
Lee, H. C.
(1997)
Physiol. Rev.
77,
1133-1164 |
43. |
Nunez, R.,
Calva, E.,
Marsch, M.,
Briones, E.,
and Lopez Soriano, F.
(1976)
Am. J. Physiol.
231,
1173-1177 |
44. | Schaper, J., and Schaper, W. (1983) J. Am. Coll. Cardiol. 1, 1037-1046[Medline] [Order article via Infotrieve] |
45. |
Thiemermann, C.,
Bowes, J.,
Myint, F. P.,
and Vane, J. R.
(1997)
Proc. Natl. Acad. Sci. U. S. A.
94,
679-683 |
46. | Pieper, A. A., Verma, A., Zhang, J., and Snyder, S. H. (1999) Trends Pharmacol. Sci. 20, 171-181[CrossRef][Medline] [Order article via Infotrieve] |
47. |
Rustin, P.,
Parfait, B.,
Chretien, D.,
Bourgeron, T.,
Djouadi, F.,
Bastin, J.,
Rotig, A.,
and Munnich, A.
(1996)
J. Biol. Chem.
271,
14785-14790 |
48. |
Petronilli, V.,
Miotto, G.,
Canton, M.,
Brini, M.,
Colonna, R.,
Bernardi, P.,
and Di Lisa, F.
(1999)
Biophys. J.
76,
725-734 |