Department of Anatomy, School of Medicine, Case Western Reserve University, Cleveland, Ohio 44106-4938
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ABSTRACT |
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Hypoxia-inducible factor-1 (HIF-1), a
heterodimeric transcription factor consisting of HIF-1 and HIF-1
subunits, controls the expression of a large number of genes involved
in the regulation of cellular responses to reduced oxygen availability.
The oxygen-regulated subunit, HIF-1
, is stabilized in cells exposed
to hypoxia. The regulation of hypoxic responses by nitric oxide (NO) is
believed to have wide pathophysiological relevance, thus we
investigated whether NO affects HIF-1 activation in hypoxic cells. Here
we show that NO generated from NO donors prevented HIF-1
hypoxic accumulation in Hep 3B and PC-12 cells. Addition of a glutathione analog or peroxynitrite scavengers prevented the NO-induced inhibition of HIF-1
accumulation in both cell lines. Exposure to NO was associated with inhibition of mitochondrial electron transport and
compensatory glycolysis, which maintained normal cellular ATP content.
Succinate, a Krebs cycle intermediate and respiratory chain substrate,
restored HIF-1
hypoxic induction in the cells, suggesting
involvement of mitochondria in regulation of HIF-1
accumulation
during hypoxia. Regulation of HIF-1
by NO is an additional important
mechanism by which NO might modulate cellular responses to hypoxia in
mammalian cells.
hypoxia-inducible factor-1; mitochondria; oxygen sensing
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INTRODUCTION |
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HYPOXIA-INDUCIBLE
FACTOR-1 (HIF-1), a general hypoxia-inducible
transcription factor, plays a central role in mediating cellular responses to hypoxia (reviewed in Refs. 41,
42, 51). HIF-1 was initially discovered
through its ability to activate the erythropoietin gene during hypoxia
via binding to the enhancer element located in the 3'-flanking region
of the gene (43). Cloning of HIF-1 revealed a
heterodimeric protein consisting of two subunits, HIF-1 and HIF-1
(48, 50). HIF-1
, identical to the aryl
hydrocarbon receptor nuclear translocator, is a common binding subunit
of many basic helix-loop-helix heterodimer transcription factors with Per-Arnt-Sim domains, besides HIF-1
(21).
HIF-1
is constitutively expressed, whereas HIF-1
is the
oxygen-regulated subunit, rapidly accumulating in cells exposed to
hypoxia (22). Under well-oxygenated conditions, the
HIF-1
protein undergoes rapid ubiquitination and degradation by the
proteasome system (23, 28, 38). HIF-1
is ubiquitinated
by the von Hippel-Lindau protein (pVHL) (34), which binds
directly to the oxygen-dependent degradation domain of HIF-1
and
targets it for proteasome degradation (34, 46). VHL is
associated with elongins B and C, cullin-2, and likely other factors
that constitute part of a multiprotein complex (35, 46).
Interaction between pVHL and a specific domain of the HIF-1
subunit was reported to be regulated through hydroxylation of a
proline residue by an enzyme prolyl hydroxylase (15, 25, 26). Prolyl hydroxylases have a requirement for dioxygen, iron, and 2-oxoglutarate (25, 26). In hypoxic conditions
prolyl hydroxylation of the HIF-1
subunit is suppressed,
leading to stabilization of the protein. Models based on a putative
hemeprotein with oxygen-sensing properties (18), NADPH
oxidase activity, and reactive oxygen species formation
(16) have been proposed earlier to explain the ability of
cells to sense changes in the oxygen concentration. We (1)
and others (10, 11) have recently suggested that signaling
from mitochondria might contribute to the activation of HIF-1
during hypoxia.
HIF-1 activation can be modulated by various factors including
nitric oxide (NO; reviewed in Ref. 42). The regulation of hypoxic responses by NO is believed to have wider pathophysiological relevance. NO has been implicated in developmental and physiological responses to hypoxia in Drosophila melanogaster
(52). Physiological concentrations of NO inhibit
respiratory complex IV (cytochrome-c oxidase) in a manner
that is competitive with oxygen, and this has potential to
modulate cellular respiration (reviewed in Ref. 9).
Importantly, high concentrations of NO, which cause damage to mitochondria, are encountered in various pathologies
associated with hypoxia (such as brain ischemia-reperfusion
injury) (7); thus NO may directly affect the outcome in
these diseases (reviewed in Ref. 6).
Regulation of HIF-1 by NO is an additional important mechanism by
which NO might modulate cellular responses to hypoxia. Recent studies
addressing regulation of HIF-1 by NO reveal a complex picture. NO was
found to inhibit HIF-1 DNA-binding activity and hypoxia-inducible gene
expression in hypoxic cells (32, 44). Inhibition of
HIF-1
accumulation in hypoxic cells by NO was reported (24), but this was not corroborated by others (32,
44). NO is also reported to induce HIF-1
, HIF-1 DNA-binding
activity, and expression of HIF-1 target genes under nonhypoxic
conditions via mechanisms that differ from NO suppression of HIF-1
activation in hypoxia (29, 30, 37, 40).
HIF-1 protein accumulation is the initial and crucial step in the
activation of HIF-1 during hypoxia. Thus we address here specifically
whether NO prevents HIF-1
protein accumulation in hypoxic cells and
look for the possible involvement of mitochondria in mediating the
effect of NO.
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MATERIALS AND METHODS |
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Cell culture and reagents. A human hepatoblastoma cell line Hep 3B (ATCC HB-8064) was maintained in MEM supplemented with 10% fetal bovine serum and 1% penicillin/streptomycin. PC-12 cells (ATCC CRL-1721) were grown in RPMI 1640 medium, supplemented with 10% fetal bovine serum, 5% horse serum, and 1% penicillin/streptomycin. All media and antibiotics were purchased from Life Technologies. The NO donor 2,2-(hydroxynitrosohydrazino)bis-ethanamine (DETA-NO; NOC-18) and glutathione ethyl ester (GSH-ethyl ester) were obtained from Calbiochem (San Diego, CA). S-nitroso-N-acetylpenicillamine (SNAP), and other reagents were purchased from Sigma (St. Louis, MO). Cells were pretreated with vehicle control or indicated agents for 1.5 h before exposure to hypoxia (1% O2-5% CO2-94% N2) for 4 h in Plexiglas modular chambers (Billups, Rothenburg). When indicated, cells were permeabilized with 4 µg/ml digitonin in a medium containing 4 mM succinate.
NO measurements. Concentrations of NO produced by DETA-NO and SNAP were measured using modified differential pulse voltametry as previously described (14). The electrochemical measurements were performed with BAS (BioAnalytical Systems) 100B/W and CH 660 (CH Instruments) electrochemical analyzers. A platinum wire (99.99%, Goodfellow) with a diameter of 25-75 µm was sealed into a glass capillary. To prepare standard NO stock solutions, NO was generated chemically from KNO2 by an excess amount of ascorbic acid, which reduces nitrite to NO (14). A solution containing (in mM) 30 KNO2, 30 HCl, and 90 NaCl was purged with nitrogen for 15 min in a rubber-sealed glass vial to remove traces of oxygen, followed by the addition of 30 mM crystalline ascorbic acid. Before adding NO donor, the cells were washed and medium was replaced with a balanced buffer (pH 7.0) containing (in mM) 108 NaCl, 5 KCl, 1.5 CaCl2, 0.5 MgCl2, 0.5 KH2PO4, 0.5 Na2HPO4, 25 HEPES, and 5 glucose. Cells were grown to confluency in 35-mm dishes, and the electrode for NO measurements was placed ~1 mm above the attached cells. Data analysis and numerical procedures were carried out with SigmaPlot 3.0 (Jandel Scientific).
Immunoblot analysis.
To determine HIF-1 protein levels in cultured cells, nuclear
extracts were prepared and 20 µg of nuclear protein per lane were
used for electrophoresis. Membranes were blocked with 5% nonfat dry
milk, incubated with monoclonal anti-HIF-1
antibodies either from
Transduction Laboratories (Lexington, KY ) or Novus Biologicals
(Littleton, CO), followed by secondary antibody detection by enhanced
chemiluminescence (Amersham, Piscataway, NJ).
Nuclear extracts and DNA electrophoretic mobility shift assay. After treatment, cells were scraped into cold PBS, centrifuged, and washed in five packed cell volumes of buffer containing (in mM) 10 Tris · HCl (pH 7.5), 1.5 MgCl2, and 10 KCl and freshly supplemented with dithiothreitol (DTT), sodium vanadate, phenylmethylsulfonyl fluoride (PMSF), leupeptin, and aprotinin. This was followed by 10 min of incubation in the same buffer on ice and homogenization in a glass Dounce homogenizer (49). Nuclei were pelleted by centrifugation at 10,000 g for 10 min, the supernatant was discarded, and nuclei were resuspended in a buffer [0.42 M KCl, 20 mM Tris · HCl (pH 7.5), 20% (vol/vol) glycerol, and 1.5 mM MgCl2], freshly supplemented with DTT, sodium vanadate, PMSF, leupeptin, and aprotinin. The suspension was rotated at 4°C for 30 min and centrifuged for 30 min at 14,000 rpm. The supernatant containing the nuclear proteins was collected. Nuclear proteins were used for an electrophoretic mobility shift assay (49), using oligonucleotide probe from the erythropoietin enhancer region, which includes the HIF-1 binding site (5'-GCCCTACGTGCTGTCTCA-3').
Measurement of complex I activity. Mitochondria from cultured cells were prepared by discontinuous Percoll gradient centrifugation (2). Adherent cells were harvested and homogenized in an isolation buffer containing 0.32 M sucrose, 1 mM EDTA (K+ salt), and 10 mM Tris · HCl (pH 7.4). The homogenate was centrifuged at 1,330 g for 3 min and the supernatant recentrifuged at 21,200 g for 10 min. Collected mitochondria were isolated in a Percoll gradient. Complex I activity was measured as described previously (1), by measuring the oxidation of NADH at 340 nm using decylubiquinone as the electron acceptor. A total of 20-40 µg of mitochondrial protein were used, and assays were performed at 37°C in a Beckman DU 640 spectrophotometer. Data were reported as means ± SD. Statistical analysis between multiple groups was performed by one-way ANOVA with Tukey correction, and P < 0.05 was considered significant.
Cell viability analysis. After cells were exposed for 5.5 h to NO donors, cell viability was estimated by trypan blue exclusion (0.4% trypan blue, light microscopy) and lactate dehydrogenase (LDH) release in the medium (5). LDH release into the medium following exposure to 40 µg/ml of digitonin was taken as 100% of total activity.
ATP and lactate assays.
Intracellular concentrations of ATP and lactate released in the media
were assayed. Lactate was measured using a lactate analyzer (YSI 2300, STAT Plus; Yellow Springs Instrument, Yellow Springs, OH). For ATP
measurements, Hep 3B and PC-12 cells were washed with PBS and lysed by
adding 0.1M NaOH. The cells and media were collected, rapidly frozen,
and stored at 80°C. ATP intracellular concentrations were analyzed
using a luciferase bioluminescence procedure previously described
(33). Briefly, 10 µl of cell extract were mixed with 50 µl of ATP reagent [solution mixture: 0.5 ml of imidazole-HCl buffer
(1 M), 0.02 ml of MgCl2 (1 M), and 0.750 ml of KCl (1 M)]
and vortexed. To measure ATP content, 10 µl of cell lysate were mixed
with luciferin-luciferase reagent (250 µl) and the light emission was
measured using a Chrono-log Lumi-Vette luminometer S900 (Chrono-log,
Havertown, PA). All experiments were performed in triplicates,
and the ATP content in nanomoles was normalized for cell protein. The
protein content was measured using a standard Bradford assay (Bio-Rad).
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RESULTS |
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SNAP prevents HIF-1 protein accumulation in Hep 3B and PC-12
cells during hypoxia.
Here we show that incubation of Hep 3B and PC-12 cells with SNAP
prevented hypoxic accumulation of HIF-1
(Fig.
1, A and
B). At the concentration of SNAP used (50 µM), a
concentration of 0.46 ± 0.1 µM NO (means ± SD,
n = 3) was achieved at the end of the experiment (5.5 h
of exposure). NO and its derivative peroxynitrite (ONOO
) can interact with a large number of proteins to
modulate various cellular functions, including cell respiration
(19). Incubation of cells with NO donors is reported to
result in a rapid and reversible inhibition of cytochrome-c
oxidase (complex IV), which can be reversed by removing NO (4, 8,
13). After prolonged incubation, cells show an additional
inhibition of respiration, which appears to be due to inhibition of
complex I (4, 8, 35, 36) and could be prevented or
reversed by thiols (such as GSH-ethyl ester) or exposure to "cold"
light (illumination from a halogen bulb) (4, 8).
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DETA-NO (NOC-18) prevents HIF-1 accumulation in Hep 3B and PC-12
cells during hypoxia.
DETA-NO, a chemically different agent from SNAP, is a slow-releasing NO
donor (half-life of NO release
20 h) characterized by a
controlled and spontaneous release of NO in solution. At the
concentration of DETA-NO used (1 mM), a concentration of 1.48 ± 0.12 µM NO (means ± SD, n = 3) was achieved at
the end of the experiment (5.5 h of exposure). DETA-NO effectively
blocked accumulation of HIF-1
in hypoxic Hep 3B (Fig.
2A) and PC-12 cells (Fig.
2B). Thus the effects of SNAP can be reproduced by using a
different NO donor, further confirming that release of NO is
specifically responsible for suppression of HIF-1
during hypoxia. In
agreement with the results obtained with SNAP, hypoxic Hep 3B and PC-12 cells (Fig. 2, A and B, respectively) treated
with DETA-NO regained the ability to induce HIF-1
in the presence of
GSH-ethyl ester (2 mM). These results confirm that excessive NO
prevents HIF-1
accumulation in hypoxic cells.
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Effects of NO, GSH-ethyl ester, and uric acid on mitochondrial
complex I activity.
The mitochondrial model of oxygen sensing proposed by Schumacker and
colleagues (10, 11) assumes that inhibition of electron flow at complex I prevents hypoxic accumulation of HIF-1. Thus any
significant inhibition of complex I by NO would be expected to prevent
HIF-1
hypoxic accumulation. This prompted us to examine complex I
activity in NO-treated cells.
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Succinate, a Krebs cycle intermediate and complex II respiratory
substrate, overcomes NO suppression of HIF-1 stabilization in
hypoxic cells.
Inhibition of mitochondrial complex I (1, 10, 11), as well
as complex III (at the myxothiazol binding site) (10, 11),
but not inhibition of complex IV (41), was reported to prevent induction of HIF-1
in hypoxic cells. Thus electron flux into
complex III was suggested to be necessary for normal HIF-1
accumulation during hypoxia (10, 11). If NO prevents
HIF-1
hypoxic accumulation by inhibiting complex I activity, then NO suppression should be overcome by bypassing complex I and using complex
II as an alternative route of electron transport into complex III. In
this experiment succinate was used as a complex II substrate in
digitonin-permeabilized cells. In both Hep 3B (Fig.
4A) and PC-12 (Fig.
4B) cells, DETA-NO prevented accumulation of HIF-1
during
hypoxia. However, DETA-NO suppression of HIF-1
was overcome in the
presence of succinate (4 mM) and digitonin (4 µg/ml) in both cell
lines (Fig. 4, A and B). We also assayed HIF-1
DNA binding in nuclear extracts from PC-12 cells. As expected, HIF-1
DNA binding paralleled the increase in the HIF-1
protein levels
after hypoxia, DETA-NO, and succinate treatments (Fig. 4B).
To exclude the possibility that digitonin or succinate may alter
HIF-1
protein levels, we tested the possible effects of these two
agents in the absence of NO donors (Fig. 4C). Digitonin, either alone or combined with succinate, had no effect on normoxic or
hypoxic protein levels of HIF-1
(Fig. 4C). Thus
NO-treated hypoxic Hep 3B and PC-12 cells regained the ability to
induce HIF-1
in the presence of succinate, which maintains electron flow via complex II into complex III. These results show that complex I
is targeted by NO and suggest that maintaining electron transport chain
activity into complex III is necessary for hypoxic induction of
HIF-1
.
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Effects of DETA-NO and SNAP on normoxic induction of HIF-1
protein levels in normoxic Hep 3B and PC-12 cells.
In view of recent reports that NO induces HIF-1
during normoxia
(29, 30, 37, 39), we tested the effects of DETA-NO and
SNAP in normoxic Hep 3B and PC-12 cells (Fig.
7, A and B, respectively). DETA-NO (1 mM) induced only minimal and transient HIF-1
protein expression in normoxic Hep 3B cells after 5.5 and 8 h (Fig. 7A, lanes 3 and 4), but
not in PC-12 cells (Fig. 7B, lanes 3 and
4). No induction of HIF-1
during normoxia could be demonstrated with SNAP (50 µM) in either cell line (Fig. 7,
A and B, lanes 5 and 6).
Therefore, under conditions tested, we observed only a minor induction
of HIF-1
by DETA-NO in normoxic Hep 3B cells.
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DISCUSSION |
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Current attempts to understand the molecular basis of cellular
oxygen sensing are focused on the regulation of HIF-1 and the key
question: How is the decrease in oxygen concentration sensed by the
cells and what signaling pathways lead to HIF-1
protein
stabilization? An important development are the recent reports of
HIF-1
regulation via direct proline hydroxylation by an enzyme
prolyl hydroxylase, which requires molecular oxygen, iron, and
2-oxoglutarate for enzymatic activity (15, 25, 26). The
requirement for molecular oxygen and iron for proline hydroxylase activity could explain the stabilization of HIF-1
under hypoxic conditions or after treatment with agents that eliminate or compete with iron. However, the process of oxygen sensing is likely more complex. Earlier studies suggested that oxygen sensing involves reactive oxygen species, protein phosphorylation, signaling from mitochondria, and activation of multiple signal transduction pathways (reviewed in Refs. 41, 42, 51).
Conceivably, these changes could directly or indirectly affect various
steps of HIF-1
activation, including proline hydroxylation, although
the connections are not clear at present.
Although several recent reports agree that NO inhibits HIF-1 DNA
binding and transcriptional activation of HIF-1 target genes (24,
32, 44), there appears to be no consensus when it comes to the
ability of NO to prevent HIF-1 accumulation during hypoxia. Our
results show that excessive NO abrogates HIF-1
accumulation in
hypoxic cells. To explain the mechanisms of NO suppression of HIF-1
activation, it was suggested that NO might interact with a putative
oxygen sensor (24). Importantly, HIF-1
itself could be
the target of S-nitrosylation; however, an earlier study found no evidence for direct modification of HIF-1
by NO, making this an unlikely explanation (24).
Herein we tested an alternative explanation for the NO suppression of
HIF-1 that involves the primary inhibition of the mitochondrial electron transport by NO. Physiological and reversible inhibition of
complex IV by NO is an early event. However, with time, a pathological inhibition/damage of complex I is reported to occur (4,
8). Our study shows that both SNAP and DETA-NO caused a marked
52 and 70-74% complex I inhibition, respectively. Thus, on the
basis of the mitochondrial model of HIF-1 regulation (10,
11), the potent inhibitory effect of NO on complex I could
explain its ability to block hypoxic accumulation of HIF-1. This
appears to be further indirectly supported by the findings that
1) addition of GSH-ethyl ester prevented the decrease of
complex I activity by NO and restored HIF-1
hypoxic accumulation and
2) uric acid, a scavenger of peroxynitrite, markedly
attenuated the inhibition of complex I activity and restored HIF-1
hypoxic induction in the presence of NO. These findings show a
correlation between complex I inhibition and impaired HIF-1
hypoxic accumulation.
NO suppression of HIF-1 hypoxic accumulation could be overcome by
the complex II substrate succinate, which restores electron flow into
complex III, indicating an isolated inhibition/damage of complex I by
NO. Under these experimental conditions, NO should also inhibit complex
IV (reviewed in Ref. 9), thus succinate does not fully
restore electron transport. However, complex IV inhibition was reported
to have no effect on HIF-1
hypoxic induction (41).
Again, these findings seem to be in agreement with the proposed model
according to which the mitochondrial signal for HIF-1
stabilization
is generated at complex III (10, 11), although a possible
alternative explanation emerges from recently reported studies
(25, 26, 45, 47). The requirement for an active
mitochondrial electron transport chain in hypoxic HIF-1
accumulation
was very recently questioned in two reports, which show that HIF-1
protein stabilization and transcriptional activity were preserved in
cells lacking mitochondrial DNA (
0 cells) (45, 47). In these two studies, the HIF-1
response was tested in cells exposed to 0.1% O2 (nearly anoxia) (47)
and 0.5% O2 (45), rather than 1.5%
O2 (10, 11) or 1% (1, 20). If
prolyl hydroxylases require oxygen as a substrate, then severe oxygen deprivation close to anoxia should cause HIF-1
stabilization, because in the virtual absence of oxygen the proline hydroxylation would always be inhibited. Hence, the fact that anoxia (or near anoxia)
stabilizes HIF-1
in cells does not exclude the possibility that
mitochondrial complex III might be required for a HIF-1
response
toward a less severe oxygen deprivation (hypoxia). In support of the
mitochondrial model, it was previously reported that pharmacological
inhibitors of complex I rotenone (1, 10, 11, 20),
1-methyl-4-phenylpiridinium (1), diphenylene iodonium (10, 11), and complex III inhibitor myxothiazol (10,
11) prevent hypoxic stabilization of HIF-1
. Clearly, further
studies are required to resolve the existing controversies and to
delineate the role of the respiratory chain in HIF-1 regulation. These
results, however, do not rule out the possible involvement of the
mitochondria in regulation of HIF-1 via different mechanisms. For
example, a novel prolyl hydroxylase was shown to regulate HIF-1
degradation/stabilization (25, 26). Prolyl hydroxylases
require the Krebs cycle intermediate 2-oxoglutarate (
-ketoglutarate)
as cosubstrate (25, 26). It was shown that synthesized
structural analogs of 2-oxoglutarate can interfere with HIF-1
protein induction (26). Thus perturbations in Krebs cycle
metabolism may possibly have a potential to interfere with regulation
of HIF-1
. The ability of succinate to restore HIF-1
induction
thus may hint toward an alternative explanation that metabolic effects
of Krebs cycle intermediates, rather than restoration of electron
transport, might impinge on signaling pathways that lead to HIF-1
accumulation under hypoxic conditions. This appears intriguing, because
it could reconcile the role of mitochondria with the model of
oxygen-dependent prolyl hydroxylation of HIF-1
.
In view of the recent intriguing reports that NO induces HIF-1 under
normoxic conditions, we tested the effect of SNAP and DETA-NO in
normoxic Hep 3B and PC-12 cells. We observed a minimal induction of
HIF-1
only in Hep 3B cells after DETA-NO, and no effect of SNAP in
either cell line (Fig. 7, A and B). Although the
use of different NO donors, dose, length of treatment, and cell
type-specific responses may account for some of the observed differences between this and previous studies, the issue may be more
complex (29, 30, 37, 39). Reactive oxygen and nitrogen species might each modulate HIF-1
expression in hypoxic cells via
distinct mechanisms (17). Recently, it was suggested that distinct cis elements in the promotor of some genes (such as
the vascular endothelial factor) bind protein complexes and mediate gene induction by NO under normoxic conditions (30). The
mechanisms by which NO suppresses HIF-1
hypoxic stabilization thus
differ from the activation of HIF-1
in normoxic cells.
In conclusion, we show that NO interferes with the signaling events
that lead to HIF-1 accumulation during hypoxia. This may have
important consequences, because a number of pathologies associated with
oxygen deprivation are also linked to excessive NO production. NO
suppression of HIF-1
hypoxic accumulation was counteracted by the
Krebs cycle intermediate succinate. This suggests that succinate acts
by restoring electron flow into respiratory complex III. Alternatively,
this may indicate an unsuspected metabolic effect of succinate, which
hints toward a link between the Krebs cycle and regulation of HIF-1
under hypoxic conditions.
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ACKNOWLEDGEMENTS |
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We thank Miklos Gratzel and Gautham Shetty for help with nitric oxide measurements and Sue Foss and Max Neal for preparing and editing the manuscript.
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FOOTNOTES |
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This work was supported by National Institute of Neurological Disorders and Stroke Grants NS-41309 (to F. H. Agani), and NS-37111 and NS-38632 (to J. LaManna). F. H. Agani is also supported by National Heart, Lung, and Blood Institute Grant HL-56470.
Address for reprint requests and other correspondence: F. H. Agani, Dept. of Anatomy, School of Medicine, Case Western Reserve Univ., 10900 Euclid Ave., Cleveland, OH 44106 (E-mail: fxa5{at}po.cwru.edu).
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.
First published February 20, 2002;10.1152/ajpcell.00381.2001
Received 7 August 2001; accepted in final form 18 February 2002.
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