Mitochondrial Electron Transport Complex I Is a Potential
Source of Oxygen Free Radicals in the Failing Myocardium
Tomomi Ide, Hiroyuki Tsutsui, Shintaro Kinugawa, Hideo Utsumi, Dongchon Kang, Nobutaka Hattori,
Koji Uchida, Ken-ichi Arimura, Kensuke Egashira, Akira Takeshita
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Abstract—Oxidative stress in the myocardium may play an important role in the pathogenesis of congestive heart failure
(HF). However, the cellular sources and mechanisms for the enhanced generation of reactive oxygen species (ROS) in
the failing myocardium remain unknown. The amount of thiobarbituric acid reactive substances increased in the canine
HF hearts subjected to rapid ventricular pacing for 4 weeks, and immunohistochemical staining of 4-hydroxy-2-nonenal
ROS-induced lipid peroxides was detected in cardiac myocytes but not in interstitial cells of HF animals. The generation
of superoxide anion was directly assessed in the submitochondrial fractions by use of electron spin resonance
spectroscopy with spin trapping agent, 5,59-dimethyl-1-pyrroline-N-oxide, in the presence of NADH and succinate as
a substrate for NADH– ubiquinone oxidoreductase (complex I) and succinate– ubiquinone oxidoreductase (complex II),
respectively. Superoxide production was increased 2.8-fold (P,0.01) in HF, which was due to the functional block of
electron transport at complex I. The enzymatic activity of complex I decreased in HF (274613 versus 13669 nmol z
min21 z mg21 protein, P,0.01), which may thus have caused the functional uncoupling of the respiratory chain and the
deleterious ROS production in HF mitochondria. The present study provided direct evidence for the involvement of ROS
in the mitochondrial origin of HF myocytes, which might be responsible for both contractile dysfunction and structural
damage to the myocardium. (Circ Res. 1999;85:357-363.)
Key Words: antioxidant n free radical n heart failure n myocardial contraction n reactive oxygen species
C
ongestive heart failure (HF) is an important cause of
morbidity and mortality in patients with various heart
diseases. Despite extensive studies, the fundamental mechanisms responsible for the development and progression of left
ventricular (LV) failure have not yet been fully elucidated.
Reactive oxygen species (ROS) such as superoxide anions
(zO22) and hydroxy radicals (zOH) cause the oxidation of
membrane phospholipids, proteins, and DNAs, and they have
been implicated in a wide range of pathological conditions
including ischemia-reperfusion injury, neurodegenerative diseases, and aging. Under physiological conditions, their toxic
effects are prevented by such scavenging enzymes as superoxide dismutase (SOD), glutathione peroxidase, and catalase
as well as by other nonenzymatic antioxidants. However,
when the production of ROS becomes excessive, oxidative
stress might have a harmful effect on the functional integrity
of biological tissue. Oxygen free radicals have been shown to
cause contractile failure and structural damage in the myocardium.1,2 However, their significance has been demonstrated to limited subsets of cardiac diseases, which include
ischemic heart disease3 and adriamycin-induced cardiac
toxicity.4
Recent investigations have suggested the generation of
ROS to increase in chronic HF. Lipid peroxides and 8-isoprostaglandin F2a, which are the major biochemical consequences of ROS generation, have been shown to be elevated
in plasma and pericardial fluid of patients with HF and also
positively correlated to the severity of HF.5–7 In addition, a
decrease of myocardial antioxidant reserve has been shown in
animal models of HF.8 –10 However, these experimental and
clinical findings have provided only indirect evidence of ROS
generation in failing myocardial tissue. Direct measurements
of ROS using electron spin resonance (ESR) spectroscopy11
are therefore needed for direct quantitation within biological
tissues.
The cellular sources and mechanisms for the increased
ROS in HF also remain to be elucidated. Within the heart,
possible cellular sources of ROS generation include cardiac
myocytes, endothelial cells, and neutrophils. The contribution
of neutrophils to ROS generation was found to be minor in an
animal model of HF owing to rapid ventricular pacing,
because the infiltration of inflammatory cells within the
myocardium is not prominent. Within cardiac myocytes,
Received February 15, 1999; accepted June 25, 1999.
From the Research Institute of Angiocardiology and Cardiovascular Clinic (T.I., H.T., S.K., K.A., K.E., A.T.) and Department of Clinical Chemistry
and Laboratory Medicine (H.U.), Kyushu University School of Medicine, Fukuoka, Japan; Department of Biophysics (D.K.), Kyushu University,
Fukuoka, Japan; Department of Neurology (N.H.), Juntendo University School of Medicine, Tokyo, Japan; and Laboratory of Food and Biodynamics
(K.U.), Nagoya University School of Bioagricultural Sciences, Nagoya, Japan.
Correspondence to Hiroyuki Tsutsui, MD, PhD, Research Institute of Angiocardiology and Cardiovascular Clinic, Kyushu University School of
Medicine, 3-1-1 Maidashi, Higashi-ku, Fukuoka, 812-8582, Japan. E-mail [email protected]
© 1999 American Heart Association, Inc.
Circulation Research is available at http://www.circresaha.org
357
358
Circulation Research
August 20, 1999
oxygen free radicals can be produced by several mechanisms
including the mitochondrial electron transport and xanthine
dehydrogenase/xanthine oxidase.3 Because of the very low
xanthine oxidase activity in some species12 and an extreme
abundance of mitochondria in cardiac myocytes, mitochondrial electron transport could thus be a major subcellular
source of ROS in the failing myocardium.13,14 Therefore, the
ROS production was directly monitored in the mitochondria
isolated from canine failing hearts induced by rapid ventricular pacing by means of ESR spectroscopy in the presence of
a spin trap.11 The magnitude and mechanisms of ROS
generation in the failing hearts were also compared with the
findings in similar preparations from normal control hearts.
Materials and Methods
Animal Model of HF
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HF was produced in adult mongrel dogs (15 to 25 kg body weight)
by rapid ventricular pacing at 240 bpm for 4 weeks, in accordance
with methods described previously.15 All operative procedures were
carried out under full surgical anesthesia with sodium pentobarbital
(25 mg/kg IV). Two-dimensional and M-mode echocardiograms
were recorded by ultrasonography. LV short-axis (cross-sectional)
views were recorded at the papillary muscle level, and the internal
LV dimensions were measured. The LV ejection fraction (%) was
calculated using the formula [(LV End-Diastolic Dimension)32(LV
End-Systolic Dimension)3]/[(LV End-Diastolic Dimension)3]3100.
After making echocardiographic recordings, the animals were generally anesthetized, intubated, and ventilated with a respirator on a
heating pad to maintain body temperature at 37°C. A catheter was
inserted into the aortic arch via the left carotid artery to measure the
systemic arterial pressure. After a thoracotomy was performed, an
externally calibrated 7F catheter-tipped pressure transducer was
inserted into the LV through the left atrium for the measurement of
LV pressure. At the time of the terminal study, the animals were
killed with a lethal dose of a-chloralose. All LV myocardial samples
were immediately excised, freeze-clamped, and stored in liquid
nitrogen for the subsequent ESR measurements.
All procedures and animal care were approved by the Committee
on Ethics of Animal Experiments, Faculty of Medicine, Kyushu
University, and were conducted according to the Guidelines for
Animal Experiments of Faculty of Medicine, Kyushu University.
Measurement of Thiobarbituric Acid Reactive
Substances (TBARS)
Lipid peroxidation is a major biochemical consequence of ROS
attack on biological tissue. We therefore determined the degree of
lipid peroxidation in the myocardial tissue through biochemical
assay of TBARS.16 LV myocardial tissue was homogenized (10%
wt/vol) in 1.15% KCl solution (pH 7.4). The homogenate was mixed
with 0.4% sodium dodecyl sulfate, 7.5% acetic acid adjusted to pH
3.5 with NaOH, and 0.3% thiobarbituric acid. Butylated hydroxytoluene (0.01%) was added to the assay mixture to prevent autoxidation of the sample. The mixture was kept at 5°C for 60 minutes and
was heated at 100°C for 60 minutes. After cooling, the mixture was
extracted with distilled water and n-butanol:pyridine (15:1, vol/vol)
and centrifuged at 1600g for 10 minutes. The absorbance of the
organic phase was measured at 532 nm. The amount of TBARS was
determined by the absorbance with the molecular extinction coefficient of 156 000 and expressed as mmol/g wet weight.
Immunohistochemistry of 4-Hydroxy-2-Nonenal
(HNE)–Modified Protein
To assess the cellular localization of lipid peroxidation by histochemical analysis, sections of LV myocardium obtained from the
free wall at the papillary muscle level were immunolabeled with an
antibody raised against HNE-modified protein, an aldehydic byprod-
uct of lipid peroxidation.17,18 Paraffin-embedded tissue sections
(3 mm thick) were deparaffinized with xylene and refixed with
Bouin’s solution for 20 minutes and immersed in 70%, 90%, and
100% ethanol to remove picric acid. To inhibit endogenous peroxidase, the sections were incubated with 0.3% H2O2 in methanol for 30
minutes. After rinsing in 0.01 mol/L PBS, the sections were
incubated with normal goat serum (diluted to 1:10) to inhibit
nonspecific binding of antibodies. The sections were further incubated with polyclonal antiserum raised against an HNE-modified
histidyl peptide (Gly3-His-Gly3) conjugated with keyhole-limpet
hemocyanin, which had been confirmed to be specific in our
previous studies.17,18 After rinsing with 0.01 mol/L PBS, the sections
were incubated with biotin-labeled goat anti-rabbit IgG antiserum
(diluted 1:100; DAKO A/S) for 60 minutes and then with avidinbiotin complex (Vectastain ABC kit; 1:100) for 60 minutes. After
rinsing, the sections were finally incubated with 0.02% 3,39diaminobenzidine and 0.03% hydrogen peroxide in deionized water
for 6 to 9 minutes. As a negative control, the sections were also
incubated with normal rabbit serum.
A morphometric analysis of HNE-positive myocardial area was
performed with tissue sections stained with HNE. Briefly, each
section was photographed under a microscope and magnified (final
magnification, 3200). Three to four fields were randomly selected
from one or two coronal sections in each animal. As a result, the
HNE-positive areas were measured at approximately five to seven
fields for each animal. Within each field, myocardial segments that
stained positively with anti-HNE antibody were identified and were
manually traced by using a digitizing pad with a computer to
calculate the traced area.
Preparation of Cardiac Subcellular Fractions
The frozen LV tissues were homogenized at 4°C for 1 minute in 6
volumes of buffer consisting of 10 mmol/L HEPES-NaOH (pH 7.4)
and 250 mmol/L sucrose with a Polytron homogenizer. The homogenate was centrifuged at 4°C and 700g for 10 minutes to remove any
nuclear and myofibrillar debris, and the resultant supernatant was
centrifuged at 7000g for 10 minutes to separate any cardiac subcellular fractions. To isolate the mitochondria, the pellet was resuspended at 4°C in a buffer consisting of 10 mmol/L HEPES-NaOH
(pH 7.4), 1 mmol/L EDTA, and 250 mmol/L sucrose (HES) and was
washed three times with HES buffer. The frozen and thawed
mitochondrial fraction was suspended in a buffer consisting of
10 mmol/L HEPES-NaOH (pH 7.4) and 1 mmol/L EDTA and kept
at 4°C for 30 minutes. Submitochondrial particles were prepared by
sonicating the mitochondria. The sonicated mitochondrial suspension was centrifuged at 21 000g for 10 minutes. The submitochondrial particle pellet was washed three times with HES buffer and
stored at 280°C. The postmitochondrial supernatant was centrifuged
for 1 hour at 170 000g at 4°C, and the final supernatant was used as
the cytosolic fraction. The resultant pellet was resuspended at 4°C in
HES buffer and was washed three times to obtain the microsomal
fraction. These submitochondrial particle, cytosolic, and microsomal
fractions were used for the ESR measurement of zO22.
The cardiac SOD levels were determined in the submitochondrial
particle, microsomal, and cytosolic fractions using the xanthine:xanthine oxidase:cytochrome c assay according to methods described
previously.19 The protein concentration of each fraction was adjusted
to obtain approximately a 50% inhibition of the rate of cytochrome
c reduction produced by the xanthine:xanthine oxidase system. No
SOD activity could be detected in the submitochondrial particle
fractions from either control or HF hearts. There was no significant
difference in the SOD activity of each fraction between the control
and HF.
Special care was taken to minimize any artifactual generation of
radical signals, in accordance with methods described by Zweier et
al.20
ESR Measurement of ROS
To demonstrate ROS in the mitochondria obtained from control and
HF hearts, the submitochondrial particle fractions were reacted with
the substrate and a spin trapping agent, 5,59-dimethyl-1-pyrroline-
Ide et al
TABLE 1.
Figure 1. Components of the respiratory chain in mitochondria.
FAD indicates flavin adenine nucleotide; FMN, flavin mononucleotide; Fe-S, iron-sulfur protein; Q, ubiquinone, and Cyt,
cytochrome.
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N-oxide (DMPO; LABOTEC, Ltd), and processed for ESR spectroscopy. Immediately after starting the reaction (, 45 seconds), ESR
spectra were recorded at an ambient temperature (25°C) with an ESR
spectrometer (JES-RE-1X; JEOL) operating at X-band (4.95 GHz).
The microwave power was 10 mW, the field modulation width was
0.063 mT, and the magnetic field range was swept at a scan rate of
5 mT/min. The quantitation of the DMPO signal intensity was
performed by comparing the amplitude of the observed signal to the
standard Mn21/MgO marker.
Two segments of the respiratory chain are primarily responsible
for ROS generation in the mitochondria, the NADH– ubiquinone
reductase in complex I and the ubiquinol– cytochrome c reductase in
complex III (Figure 1).21–23 NADH (200 mmol/L) was used as a
substrate for complex I and succinate (10 mmol/L) for complex II
(succinate– ubiquinone oxidoreductase).
To determine the amount of zO22 production in the microsomal
and cytosolic fraction, each fraction obtained from control and HF
hearts was reacted with NADH (200 mmol/L), and ESR spectra of
DMPO were recorded in accordance with the same methods as those
performed in the submitochondrial fractions.
Oxidative Stress in Heart Failure
359
Characteristics of Rapid Pacing-Induced HF Model
Control (n57)
HF (n57)
Body weight, kg
17.661.2
21.061.5
LV weight, g
81.468.5
92.0610.9
LV end-diastolic dimension, mm
33.762.4
49.461.4*
LV end-systolic dimension, mm
19.461.8
43.761.0†
LV ejection fraction, %
81.061.6
32.062.6†
LV peak1dP/dt, mm Hg/s
22716173
1600697.6*
6.060.8
30.163.8†
LV end-diastolic pressure, mm Hg
Values are expressed as mean6SEM. Statistical comparisons are by
unpaired t test.
*P,0.05 and †P,0.01 indicate a statistically significant difference from
control.
modified protein revealed the lipid peroxides to be positively
stained in myocytes from HF dogs, whereas no labeling was
observed in the control myocardium (Figure 2B). The myocardial area stained positively with HNE was 1866% in HF
hearts, whereas it was 260.2% in control (P,0.01).
Mitochondria as the Source of ROS Production
At baseline conditions, only small ESR signals appeared for
the submitochondrial particle fractions obtained from the
normal heart in the presence of NADH or succinate as a
substrate (Figure 3). When submitochondrial particle fractions were treated with rotenone (200 mmol/L), which selectively blocks the electron transport at the distal site of
complex I, in the presence of NADH, prominent ESR signals
Assay of Mitochondrial Complex I Activity
To delineate the basis for the block of electron transfer at complex I,
its enzymatic activity was measured. To measure the complex I
activity, the submitochondrial particle fractions were assayed for a
reduction of ubiquinone analog, decylubiquinone, using a spectrophotometer, in accordance with the method of Trounce et al24 with
some modifications. Enzyme activity was expressed in nmol z min21 z
mg21 protein.
Statistical Analysis
All data are expressed as mean6SEM. To compare the data between
control and HF, Student’s unpaired t test was used. For multiple
comparisons, one-way ANOVA was used in conjunction with the
post hoc test using Scheffé’s correction. All tests were considered to
be statistically significant at P,0.05.
Results
Contractile Dysfunction in HF Model
Rapid pacing caused an approximately 125% increase in
end-systolic LV dimension and a 60% decrease in LV
ejection fraction (Table 1). For HF dogs, LV peak positive
dP/dt was depressed, and LV end-diastolic pressure was
elevated in comparison to the control values. As a result, HF
dogs showed similar hemodynamic characteristics to those of
humans with dilated cardiomyopathy.15,25
Lipid Peroxidation of Failing Myocytes
TBARS significantly increased in HF animals (Figure 2A).
Most importantly, an immunohistochemical analysis of HNE-
Figure 2. A, Lipid peroxidation as indicated by TBARS in the
myocardial tissue from control (open bar, n57) and HF (filled
bar, n57) hearts. Statistical comparisons are by unpaired t test.
Each assay was performed in triplicate. *P,0.01 for difference
from control value. B, Immunohistochemical micrograph analysis of an HNE-modified histidine peptide in the myocardial tissue sections from control (left) and HF (right) dogs. Bar550 mm.
360
Circulation Research
August 20, 1999
TABLE 2. Magnitude of DMPO-OOH Signals in Normal
and HF Mitochondria in the Presence of NADH or Succinate
as a Substrate
Reaction System (Substrate1Drugs)
n
Magnitude of
DMPO-OOH Signals
Normal mitochondria
NADH (200 mmol/L)
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Figure 3. ESR analysis of DMPO adduct formation in normal
mitochondria in the presence of NADH as a substrate for complex I (A), succinate as a substrate for complex II (B), and in the
presence of pure zO22 generating system (C). A, Reaction mixture consisted of the submitochondrial particles obtained from
the normal heart (2 mg protein/mL), NADH (200 mmol/L) with
alcohol dehydrogenase (9 U/mL) to regenerate NADH, and
DMPO in PBS (pH 7.4). DMPO-OOH signals were amplified by
rotenone (200 mmol/L) or antimycin A (200 mmol/L). B, When
succinate was added in place of NADH, DMPO-OOH signals
were similarly enhanced by antimycin A. C, zO22 was generated
by the interaction of hypoxanthine (500 mmol/L) and xanthine
oxidase (20 mU) in vitro. DMPO revealed zO22 formation, as indicated by the DMPO-OOH spectra (•). Instrumental conditions
were as follows: X-band (9.43 GHz) ESR; microwave power 10
mW; field modulation width 0.063 mT; and sweep time 5
mT/min.
of DMPO–superoxide adduct, DMPO-OOH, were observed
with characteristic hyperfine splittings that yielded 12 resolved peaks (Figure 3A). A preliminary dose-response assay
of rotenone concentration versus DMPO-OOH signal magnitude showed a maximal effect of rotenone to be achieved at
a concentration of 200 mmol/L. We verified that the observed
DMPO signals reflected the presence of zO22 by a comparison
of the signals with the identical hyperfine splittings elicited in
the presence of pure zO22 generated from the reaction of
hypoxanthine (500 mmol/L) and xanthine oxidase (100 mU)
in vitro (Figure 3C). In addition, the DMPO signals were
completely attenuated both in the presence of SOD (100
U/mL) and by the combination of SOD plus catalase (500
U/mL) (Table 2). These results indicate that the functional
block of complex I is capable of producing zO22, which is
consistent with the findings of previous studies.21,22 In addition, antimycin A (200 mmol/L), an inhibitor of complex III,
further increased the DMPO-OOH signals in normal mitochondria, which thus indicated the additive nature of the two
sites (complex I and complex III) in mitochondrial zO22 production (Figure 3A and Table 2). DMPO-OOH signals
elicited by NADH and antimycin A were also attenuated in
the presence of SOD or SOD plus catalase (Table 2), which
indicated that the observed signals originated from zO22.
To further confirm the generation of zO22 at complex III,
normal submitochondrial particles were treated with antimycin A (200 mmol/L) in the presence of succinate (10 mmol/L).
Antimycin A also increased the DMPO-OOH signals (Figure 3B and Table 2), thus indicating that zO22 can also be
5
2.5860.84
1Rotenone (200 mmol/L)
5
8.9560.76*
1Rotenone1SOD (100 U/mL)
5
0.7360.36†
1Rotenone1SOD1catalase (500 U/mL)
5
0.0860.09†
1Antimycin A (200 mmol/L)
5
13.7363.28*
1Antimycin A1SOD (100 U/mL)
5
0.4460.09‡
1Antimycin A1SOD1catalase (500 U/mL)
5
0.1260.09‡
Succinate (10 mmol/L)
1Antimycin A (200 mmol/L)
5
0.7160.39
5
11.6461.46*
1Antimycin A1SOD (100 U/mL)
5
0.4260.25§
1Antimycin A1SOD1catalase (500 U/mL)
5
0.5260.35§
HF mitochondria
NADH (200 mmol/L)
5
6.6661.08
1Rotenone (200 mmol/L)
5
8.1760.96
1Antimycin A (200 mmol/L)
5
15.9861.52*
Succinate (10 mmol/L)
1Antimycin A (200 mmol/L)
5
0.3360.26
5
14.6661.11*
Values are expressed as mean6SEM. n indicates number of preparations.
Effects of mitochondrial electron transport inhibitors (rotenone and antimycin A)
and radical scavengers were examined in 5 preparations in each normal and
HF group. Statistical comparisons are by one-way ANOVA with a post hoc test
using Scheffé’s correction.
*P,0.05, statistically different from the substrate (NADH or succinate);
†P,0.05, statistically different from NADH1rotenone; ‡P,0.05, statistically
different from NADH1antimycin A; and §P,0.05, statistically different from
succinate1antimycin A.
produced from the cytochrome b-c1 segment of complex III.
The DMPO signals elicited by succinate and antimycin A
were abolished in the presence of SOD or SOD plus catalase
(Table 2). This finding is consistent with a previous study that
showed zO22 to be produced by a reduced component of
complex III when antimycin A is present.23 Taken together,
these results suggest that both complex I and complex III are
important sources of ROS in cardiac myocytes.
ROS Production in HF Mitochondria
To determine whether zO22 production is enhanced in mitochondria isolated from HF and to identify its production site,
submitochondrial particles obtained from HF hearts were
reacted with NADH (200 mmol/L), and the ESR spectra of
DMPO were recorded. Only small signals appeared for
normal control heart, whereas HF exerted a prominent ESR
signal of DMPO-OOH, thus indicating the generation of
zO22 (Figure 4A). The magnitude of zO22 was 2.8-fold
(P,0.01) higher in HF than in control mitochondria (Figure
5). For the HF mitochondria studied, each sample demonstrated DMPO-OOH spectra. Repeated experiments on the
same heart exerted the same spectra (repeated measurements
of DMPO-OOH magnitude: 10065% of initial value,
P5NS). These ESR signals were attenuated in the presence
Ide et al
Oxidative Stress in Heart Failure
361
TABLE 3. Magnitude of DMPO-OOH Signals in the Subcellular
Fractions Isolated From Control and HF in the Presence of
NADH as a Substrate
Subcellular Fraction
Microsome
Cytosol
Control (n55)
HF (n55)
2.1760.69
3.7461.18
ND
ND
Values are expressed as mean6SEM. Statistical comparisons are made by
Student’s unpaired t test. ND indicates not detectable.
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Figure 4. ESR analysis of DMPO adduct formation in HF mitochondria in the presence of NADH (A) and succinate (B). A,
Reaction mixture consisted of submitochondrial particles
obtained from the HF heart (2 mg protein/mL), NADH
(200 mmol/L) with alcohol dehydrogenase (9 U/mL) to regenerate NADH or succinate (10 mmol/L), and DMPO in PBS (pH
7.4). DMPO-OOH signals were amplified in the presence of antimycin A (200 mmol/L) but not in the presence of rotenone
(200 mmol/L). B, Similar to NADH, when succinate was reacted
with the mitochondria, the DMPO-OOH signals were enhanced
in the presence of antimycin A. The instrumental conditions are
the same as those in Figure 3.
of SOD (100 U/mL) alone or SOD (100 U/mL) plus catalase
(500 U/mL), thus suggesting that most ESR signals derived
from zO2 (Figure 5).
To examine the production of zO22 at complex III, the
submitochondrial particles were incubated with succinate
(10 mmol/L). In contrast to NADH as a substrate, ESR
demonstrated very few DMPO signals in the HF heart (Figure
4B). The production of zO22 in normal mitochondria treated
with complex I inhibitor, rotenone, in the presence of NADH
(Figure 3A) was comparable to the baseline zO22 production
in the HF mitochondria (Figure 4A). An important finding
was that zO22 production was initiated by the addition of
NADH but not of succinate, thus indicating that the electron
transfer function at complex I was primarily responsible for
such production in HF.
zO22 production by complex I in the submitochondrial
particles from HF hearts was maximally enhanced and there
was no significant “rotenone”-recruitable reserve (Figure 4
and Table 2). We next examined whether the inhibition of
complex III in HF mitochondria could further increase the
zO22 production as shown in normal mitochondria. Antimycin
A further increased the DMPO-OOH signals in the HF
mitochondria in the presence of either NADH or succinate as
a substrate (Figure 4 and Table 2), thus indicating that the
additive nature of complex I and complex III in the zO22 production is also present in HF. Most importantly, the combination of Figure 3 and Figure 4 suggests that zO22 is produced
via a functional block at complex I in HF.
The magnitude of DMPO-OOH signals obtained from the
microsomal fractions was found to be comparable between
control and HF (Table 3). However, the signals could not be
detected in the cytosolic fractions.
Complex I Enzymatic Activity
Complex I activity significantly decreased in HF myocardium
in comparison to control values (274613 versus 13669
nmol z min21 z mg21 protein, P,0.01).
Discussion
Figure 5. Summary of data for ESR analysis of the amplitude of
DMPO-OOH signals in the submitochondrial particles obtained
from control and HF hearts in the presence of NADH or succinate as a substrate. The magnitude of the DMPO-OOH signals
was prominent in HF in the presence of NADH but not in succinate, which was attenuated by either SOD (100 U/mL) or SOD
(100 U/mL) plus catalase (500 U/mL). Statistical comparisons
were done by one-way ANOVA and a means-comparison contrast, in which n indicates the number of preparations in each
group. *P,0.05 for difference from the control value; †P,0.05
for difference from the HF value in the presence of NADH.
The present study using ESR spectroscopy provided the first
direct evidence for the increased generation of zO22 in the
mitochondria isolated from the failing heart. The decrease in
complex I enzymatic activity and the resultant impairment of
electron transfer lead to the deleterious production of ROS in
the intracellular space, which could result in the myocyte
injury in HF.
Mitochondria produce ROS through one electron carrier in
the respiratory chain. Under physiological conditions, small
quantities of ROS are formed during mitochondrial respiration, which, however, can be detoxified by the endogenous
scavenging mechanisms of myocytes. In line with the findings of previous studies,26 the inhibition of electron transport
at the sites of complex I and complex III in the normal
submitochondrial particles resulted in a significant production of zO22 (Figure 3). The most important findings of the
present study were that HF mitochondria produce more
zO22 than normal mitochondria in the presence of NADH but
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Circulation Research
August 20, 1999
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not succinate (Figure 4) and that complex I is the predominant source of such zO22 production (Figures 3 and 4, Table
2). Furthermore, HF mitochondria were also found to be
associated with a decrease in the complex I activity. Although
the present study could not prove the mechanisms for this
decreased activity, it has provided compelling evidence that
the defects in electron transfer function lead to mitochondrial
ROS production. In aging and neurodegenerative diseases
such as Parkinson disease, the mitochondria are the predominant source of ROS.27 In this respect, our results demonstrated a similar pathophysiological link between mitochondrial dysfunction and oxidative stress in failing hearts.
Cardiac myocytes are the likely targets of ROS attack in
the failing heart (Figure 2B). It is conceivable that free
radicals cause damage at or near the site of their formation,
given that they are a highly reactive and short-lived species.
Therefore, as major sources of ROS production, mitochondria
also could be major targets of ROS attack and thus be
particularly susceptible to its attack,28 which further impairs
the function of the respiratory chain (T.I., unpublished data,
1998) and accelerates zO22 production within the mitochondria. Mitochondrial ROS production may underlie the mutations and/or deletions of mitochondrial DNA, which subsequently should lead to a further impairment of the
mitochondrial function.29 The high susceptibility of mitochondrial DNA to mutation and oxidative damage is likely a
reflection of this localized production of ROS by electron
transport oxidation.30
To determine whether the decreased antioxidant capacity
may further aggravate the ROS accumulation in HF, SOD
was quantified in the myocardial tissues. However, there was
no significant difference in the SOD content between control
and HF (H.T., unpublished data, 1998), which thus indicates
that oxidative stress in HF is primarily due to the enhancement of mitochondrial prooxidant generation rather than the
decline in antioxidant defenses. It should be acknowledged
that other potential sources of ROS generation within the
heart including vascular endothelial cells (via xanthine oxidase and/or NADPH oxidase) and activated leukocytes (via
NADPH oxidase)3 could not be completely ruled out in the
present study. Although mitochondrial electron transport
plays an important role in the ROS production in HF, other
mechanisms also might be involved. The activation of neurohumoral factors commonly seen in HF, including catecholamines and cardiac sympathetic tone, renin-angiotensin system, and nitric oxide, can contribute to the generation of
ROS.31–34
The present study demonstrated that the production of ROS
and ROS-induced lipid peroxidation are enhanced in HF
heart. Oxygen radicals zOH and zO22 have been demonstrated
to impair myocardial contractile function in vivo and in
vitro.2,35 Further, lipid peroxides, byproducts of ROS generation, could also cause myocardial contractile defects and
ultimately lead to structural damage.36 Although the pathophysiology of intense cellular damage resulting from the
radical exposure commonly seen under ischemia and reperfusion could not be equated to the cardiac damage in HF,
ROS production exceeding cellular antioxidant defense capabilities can have potentially lethal consequences for cardiac
myocytes over longer periods. Recently, oxygen radicals
have been suggested to be involved in apoptotic cell
death.37,38 As a result, apoptosis might play an important role
in the pathogenesis of HF.39
Although the ESR technique is a direct method to detect
ROS within biological tissue, several crucial steps must be
performed carefully. First, given the highly transient nature of
ROS, the ESR measurement procedures themselves may
modify the status of ROS by destroying the antioxidant
defenses and/or activating the ROS production system.
Therefore, we took special care to minimize the artifactual
generation of ROS and to preserve the in situ status of ROS
by using freeze-clamped myocardial tissues, rigorously maintaining the sample under anaerobic conditions in the liquid
nitrogen.20 More importantly, a differential effect of the
experimental procedures on HF tissue seems unlikely. Second, although DMPO has been the most versatile and commonly used spin trap for measuring zO22, the half-life of
DMPO–superoxide adduct, DMPO-OOH, is as short as 50
seconds in aqueous media, and it spontaneously decomposes
to form the DMPO–OH adduct,40 which was identical to that
formed from trapping the zOH radical. To overcome these
limitations, ESR spectra were recorded immediately after
(, 45 seconds) the addition of DMPO to the sample. We
confirmed that the DMPO signals detected in HF mitochondria are specific for zO22 by use of SOD (Figure 5), and
identical DMPO-OOH signals were observed in the presence
of pure zO22 generated by the interaction of hypoxanthine and
xanthine oxidase in vitro (Figure 3). Furthermore, we obtained the same results by using 5-(diethoxyphosphoryl)-5methyl-1-pyrroline-N-oxide,40 a more sensitive and stable
spin trap than DMPO (T.I., unpublished data, 1998).
Our findings do not prove a causal relationship. Indeed,
given the importance of ROS in the pathophysiology of HF,
the effects of radical scavengers on the reversibility or
prevention of HF and the establishment of a cause-and-effect
relationship will need to be determined in future studies.
HF is associated with alterations of various signal molecules in the myocardium, including the renin-angiotensin and
sympathetic nervous systems, peptide growth factors, cytokines, nitric oxide, and ROS. All of these have the potential
to exert profound effects on the phenotype of the myocardium. There are probably multiple levels of autocrine and
paracrine interactions among these mediators resulting from
both positive and negative feedback loops; therefore, studies
in isolated tissues may not replicate the in vivo environment.41 It must be stated that a purely reductionistic approach
might overlook the critical interactions among the various
signal molecules in the intact heart. Although a more focused
experimental approach is essential for determining the cellular and subcellular mechanisms stimulating these systems in
HF, it will be important to search for a better understanding
of the integrated regulation of various mediators in HF in
future studies.
The present study provides, for the first time, direct
evidence for the increased production of zO22 at the complex
I site in the mitochondria isolated from failing cardiac
myocytes. Although the focus of this study was the heart,
these mechanisms of ROS production probably have a common foundation with other pathological conditions in other
Ide et al
organs such as the brain, liver, and kidney. These results provide
a direct link among ROS generation, mitochondrial dysfunction,
and contractile defects in HF and suggest that oxidative stress
plays an important role in the pathogenesis of HF.
21.
Acknowledgments
22.
This work was supported in part by grants from the Ministry of
Education, Science, and Culture (Nos. 07266220, 08258221,
and 09670724).
23.
References
24.
Downloaded from http://circres.ahajournals.org/ by guest on July 31, 2017
1. Bolli R. Mechanism of myocardial “stunning.” Circulation. 1990;82:
723–738.
2. Josephson RA, Silverman HS, Lakatta EG, Stern MD, Zweier JL. Study
of the mechanisms of hydrogen peroxide and hydroxyl free radicalinduced cellular injury and calcium overload in cardiac myocytes. J Biol
Chem. 1991;266:2354 –2361.
3. McCord JM. Oxygen-derived free radicals in postischemic tissue injury.
N Engl J Med. 1985;312:159 –163.
4. Doroshow JH. Effect of anthracycline antibiotics on oxygen radical formation in rat heart. Cancer Res. 1983;43:460 – 472.
5. Belch JJ, Bridges AB, Scott N, Chopra M. Oxygen free radicals and
congestive heart failure. Br Heart J. 1991;65:245–248.
6. McMurray J, Chopra M, Abdullah I, Smith WE, Dargie HJ. Evidence of
oxidative stress in chronic heart failure in humans. Eur Heart J. 1993;
14:1493–1498.
7. Mallat Z, Philip I, Lebret M, Chatel D, Maclouf J, Tedgui A. Elevated
levels of 8-iso-prostaglandin F2a in pericardial fluid of patients with heart
failure: a potential role for in vivo oxidant stress in ventricular dilatation
progression to heart failure. Circulation. 1998;97:1536 –1539.
8. Dhalla AK, Singal PK. Antioxidant changes in hypertrophied and failing
guinea pig hearts. Am J Physiol. 1994;266:H1280 –H1285.
9. Hill MF, Singal PK. Antioxidant and oxidative stress changes during
heart failure subsequent to myocardial infarction in rats. Am J Pathol.
1996;148:291–300.
10. Hill MF, Singal PK. Right and left myocardial antioxidant responses
during heart failure subsequent to myocardial infarction. Circulation.
1997;96:2414 –2420.
11. Miura Y, Utsumi H, Hamada A. Antioxidant activity of nitroxide radicals
in lipid peroxidation of rat liver microsomes. Arch Biochem Biophys.
1993;300:148 –156.
12. de Jong JW, van der Meer P, Nieukoop AS, Huizer T, Stroeve RJ, Bos E.
Xanthine oxidoreductase activity in perfused hearts of various species,
including humans. Circ Res. 1990;67:770 –773.
13. Das DK, George A, Liu X, Rao PS. Detection of hydroxy radical in the
mitochondria of ischemic-reperfused myocardium by trapping with
salicylate. Biochem Biophys Res Commun. 1989;165:1004 –1009.
14. Ambrosio G, Zweier JL, Duilio C, Kuppusamy P, Santoro G, Elia PP,
Tritto I, Cirillo P, Condorelli M, Chiariello M, Flaherty JT. Evidence that
mitochondrial respiration is a source of potentially toxic oxygen free
radicals in intact rabbit hearts subjected to ischemia and reflow. J Biol
Chem. 1993;268:18532–18541.
15. Yamamoto S, Tsutsui H, Tagawa H, Saito K, Takahashi M, Tada H,
Yamamoto M, Katoh M, Egashira K, Takeshita A. Role of myocyte nitric
oxide in b-adrenergic hyporesponsiveness in heart failure. Circulation.
1997;95:1111–1114.
16. Kikugawa K, Kojima T, Yamaki S, Kosugi H. Interpretation of the
thiobarbituric acid reactivity of rat liver and brain homogenates in the
presence of ferric ion and ethylenediaminetetraacetic acid. Anal Biochem.
1992;202:249 –255.
17. Yoritaka A, Hattori N, Uchida K, Tanaka M, Stadtman ER, Mizuno Y.
Immunohistochemical detection of 4-hydroxynonenal protein adducts in
Parkinson disease. Proc Natl Acad Sci U S A. 1996;93:2696 –2701.
18. Uchida K, Itakura K, Kawakishi S, Hiai H, Toyokuni S, Stadtman ER.
Characterization of epitopes recognized by 4-hydroxy-2-nonenal specific
antibodies. Arch Biochem Biophys. 1995;324:241–248.
19. Doroshow JH, Locker GY, Ifrim I, Myers CE. Prevention of doxorubicin
cardiac toxicity in the mouse by N-acetylcysteine. J Clin Invest. 1981;
68:1053–1064.
20. Zweier JL, Kuppusamy P, Williams R, Rayburn BK, Smith D, Weisfeldt
ML, Flaherty JT. Measurement and characterization of postischemic free
25.
26.
27.
28.
29.
30.
31.
32.
33.
34.
35.
36.
37.
38.
39.
40.
41.
Oxidative Stress in Heart Failure
363
radical generation in the isolated perfused heart. J Biol Chem. 1989;264:
18890 –18895.
Takeshige K, Minakami S. NADH- and NADPH-dependent formation of
superoxide anions by bovine heart submitochondrial particles and
NADH-ubiquinone reductase preparation. Biochem J. 1979;180:
129 –135.
Turrens JF, Boveris A. Generation of superoxide anion by the NADH
dehydrogenase of bovine heart mitochondria. Biochem J. 1980;191:
421– 427.
Turrens JF, Alexandre A, Lehninger AL. Ubisemiquinone is the electron
donor for superoxide formation by complex III of heart mitochondria.
Arch Biochem Biophys. 1985;237:408 – 414.
Trounce IA, Kim YL, Jun AS, Wallace DC. Assessment of mitochondrial
oxidative phosphorylation in patient muscle biopsies, lymphoblasts, and
transmitochondrial cell lines. Methods Enzymol. 1996;264:484 –509.
Takahashi M, Tsutsui H, Kinugawa S, Igarashi-Saito K, Yamamoto S,
Yamamoto M, Tagawa H, Imanaka-Yoshida K, Egashira K, Takeshita A.
Role of microtubules in the contractile dysfunction of myocytes from
tachycardia-induced dilated cardiomyopathy. J Mol Cell Cardiol. 1998;
30:1047–1057.
Cadenas E, Boveris A, Ragan CI, Stoppani AO. Production of superoxide
radicals and hydrogen peroxide by NADH-ubiquinone reductase and
ubiquinol-cytochrome c reductase from beef-heart mitochondria. Arch
Biochem Biophys. 1977;180:248 –257.
Schapira AH, Cooper JM, Dexter D, Jenner P, Clark JB, Marsden CD.
Mitochondrial complex I deficiency in Parkinson’s disease. Lancet. 1989;
1:1269. Letter.
Stadtman ER. Protein oxidation and aging. Science. 1992;257:
1220 –1224.
Giulivi C, Boveris A, Cadenas E. Hydroxyl radical generation during
mitochondrial electron transfer and the formation of
8-hydroxydesoxyguanosine in mitochondrial DNA. Arch Biochem
Biophys. 1995;316:909 –916.
Linnane AW, Marzuki S, Ozawa T, Tanaka M. Mitochondrial DNA
mutations as an important contributor to ageing and degenerative
diseases. Lancet. 1989;1:642– 645.
Daly PA, Sole MJ. Myocardial catecholamines and the pathophysiology
of heart failure. Circulation. 1990;82:I35–I43.
Dzau VJ, Packer M, Lilly LS, Swartz SL, Hollenberg NK, Williams GH.
Prostaglandins in severe congestive heart failure. Relation to activation of
the renin-angiotensin system and hyponatremia. N Engl J Med. 1984;310:
347–352.
Cleeter MWJ, Cooper JM, Darley-Usmar VM, Moncada S, Schapira
AHV. Reversible inhibition of cytochrome c oxidase, the terminal
enzyme of the mitochondrial respiratory chain, by nitric oxide. FEBS Lett.
1994;345:50 –54.
Prasad K, Kalra J, Bharadwaj B. Increased chemiluminescence of polymorphonuclear leucocytes in dogs with volume overload heart failure.
Br J Exp Pathol. 1989;70:463– 468.
Blaustein AS, Schine L, Brooks WW, Fanburg BL, Bing OH. Influence
of exogenously generated oxidant species on myocardial function. Am J
Physiol. 1986;250:H595–H599.
Kaneko M, Matsumoto Y, Hayashi H, Kobayashi A, Yamazaki N.
Oxygen free radicals and calcium homeostasis in the heart. Mol Cell
Biochem. 1994;139:91–100.
McGowan AJ, Ruiz-Ruiz MC, Gorman AM, Lopez-Rivas A, Cotter TG.
Reactive oxygen intermediate(s) (ROI): common mediator(s) of
poly(ADP-ribose)polymerase (PARP) cleavage and apoptosis. FEBS Lett.
1996;392:299 –303.
Cheng W, Li B, Kajstura J, Li P, Wolin MS, Sonnenblick EH, Hintze TH,
Olivetti G, Anversa P. Stretch-induced programmed myocyte cell death.
J Clin Invest. 1995;96:2247–2259.
Semigran MJ, Dec GW, Khaw BA. Apoptosis in myocytes in end-stage
heart failure. N Engl J Med. 1996;335:1182–1189.
Roubaud V, Sankarapandi S, Kuppusamy P, Tordo P, Zweier JL. Quantitative measurement of superoxide generation using the spin trap
5-(diethoxyphosphoryl)-5-methyl-1-pyrroline-N-oxide. Anal Biochem.
1997;247:404 – 411.
Cohn JN, Bristow MR, Chien KR, Colucci WS, Frazier OH, Leinwand
LA, Lorell BH, Moss AJ, Sonnenblick EH, Walsh RA, Mockrin SC,
Reinlib L. Report of the National Heart, Lung, and Blood Institute Special
Emphasis Panel on Heart Failure Research. Circulation. 1997;95:
766 –770.
Downloaded from http://circres.ahajournals.org/ by guest on July 31, 2017
Mitochondrial Electron Transport Complex I Is a Potential Source of Oxygen Free
Radicals in the Failing Myocardium
Tomomi Ide, Hiroyuki Tsutsui, Shintaro Kinugawa, Hideo Utsumi, Dongchon Kang, Nobutaka
Hattori, Koji Uchida, Ken-ichi Arimura, Kensuke Egashira and Akira Takeshita
Circ Res. 1999;85:357-363
doi: 10.1161/01.RES.85.4.357
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