Laboratoire de médecine moléculaire, Hôpital Sainte-Justine, Université du Québec à Montréal, CP 8888, Succursale centre-ville, Montréal, Québec, Canada H3C 3P8
* Author for correspondence (e-mail: oncomol{at}nobel.si.uqam.ca)
Accepted 12 February 2003
![]() |
Summary |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Key words: Hypoxia, Hypoxia inductible factor 1, Rho proteins, Carcinoma cells, Reactive oxygen species
![]() |
Introduction |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Although several studies have been performed to investigate the role of
HIF-1, little is known about the sensor mechanisms by which cells
detect [O2] modulation. Among mechanisms thought to regulate oxygen
sensing, one postulates increased ROS generation by mitochondria
(Chandel and Schumacker, 2000
).
Several studies demonstrated that ROS generation was necessary for the
transcriptional response induced by hypoxia
(Chandel and Schumacker, 2000
;
Li and Jackson, 2002
;
Duranteau et al., 1998
;
Pearlstein et al., 2002
;
Chandel et al., 1998
). The
addition of diphenylene iodonium (DPI), an inhibitor of ROS, or the use of
cells depleted in mitochondria were able to abolish the hypoxic induction of
HIF-1
, erythropoietin (EPO), glycolytic enzymes and vascular
endothelial growth factor (VEGF) (Chandel
et al., 1998
).
The expression of VEGF is increased in renal cell carcinoma (RCC) at both
the mRNA and protein levels and contributes to neovascularization and tumour
progression (Nicol et al.,
1997; Xia et al.,
2001
). Under hypoxic conditions, the induction of VEGF occurs by
activation of gene transcription and also by an increase in mRNA stability
(von Marschall et al., 2001
);
the transcriptional activation is a consequence of HIF-1
stabilization.
One member of the VEGF family, VEGF-A, contains five isoforms generated by
alternative splicing (121, 145, 165, 189 and 206 amino acids)
(Ferrara et al., 1992
;
Houck et al., 1991
). A recent
study showed that VEGF-A accounts for most of the VEGF overexpression during
hypoxia and that VEGF-B and VEGF-C do not seem to be involved
(Gunningham et al., 2001
).
The depletion of cellular ATP that occurs during hypoxia disrupts the actin
cytoskeleton in many cell types including renal epithelial cells
(Molitoris, 1991). Regulation
of the cytoskeletal architecture has been shown to be mediated by Rho
proteins, which also participates in cell adhesion, migration, invasion and in
gene transcription (Hall,
1998
; Takai et al.,
2001
). The Rho GTPase family includes several members, but RhoA,
Rac1 and Cdc42 are the best characterized. Each of these proteins is active
when bound to GTP at the membrane and maintained inactive in the cytosol when
complexed with GDP-dissociation inhibitor (GDI)
(Matozaki et al., 2000
). The
cyclic passage between these two forms is tightly controlled by different
classes of regulatory proteins (Boivin et
al., 1996
). Previous studies have demonstrated that constitutively
activated Rho proteins protect against the disruption of stress fibers,
cortical F-actin and tight junctions caused by chemical depletion of ATP by
antimycin A (Raman and Atkinson,
1999
; Gopalakrishnan et al.,
1998
). This decrease in the ATP level also occurs during hypoxia
but few studies have investigated the role of Rho proteins in this process
(Hirota and Semenza, 2001
).
Since ATP depletion and cytoskeleton disruption are early events in hypoxia,
we characterized Rho GTPase expression and investigated their potential roles
in pathways involved in hypoxia responses.
In this report, we analyzed the effects of hypoxia on Rho proteins in
Caki-1 cells, a renal carcinoma model. We observed an increase in the Cdc42
and RhoA proteins and mRNA under hypoxic conditions. This upregulation
occurred after the generation of ROS; incubation with DPI abolished both ROS
production and Rho upregulation. In addition, incubation of cells with C3
exotoxin which ADP-ribosylates and inactives RhoA, RhoB and RhoC prevented
HIF-1 mRNA and protein. Our findings demonstrated that Rho proteins are
necessary regulators of the cascade leading to HIF-1
accumulation and
permit a best understanding of carcinogenesis during hypoxia.
![]() |
Materials and Methods |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Measurement of ATP levels
After hypoxic periods of 0 to 120 minutes, Caki-1 cells were washed twice
with phosphate-buffered saline (PBS) then lysed in 6% perchloric acid (v/v).
Insoluble material was removed by centrifugation at 12,000 g
for 5 minutes, supernatants were diluted with H2O and pH was
adjusted to 7.0 with 5 M potassium carbonate. ATP levels were measured by
luminescence at 542 nm on a SpectraMAX Gemini (Molecular Devices, Sunnyvale,
CA) luminometer using a Sigma bioluminescent somatic cell assay kit following
the manufacturer's protocol (Sigma, St Louis, MO). Protein was quantified by a
micro bicinchoninic acid (micro BCA) method (Pierce, Rockford, IL).
Cytotoxicity
Cytotoxicity caused by hypoxia was measured by cleavage of
4-[3-(4-iodophenyl)-2-(4-nitrophenyl)-2H-5-tetrazolio]-1,3-benzene disulfonate
(WST-1) in formazan and followed on a spectrophotometer (Molecular Devices,
Sunnyvale, CA) at 450 nm. Following hypoxic intervals of 0 to 6 hours, 10,000
cells were placed overnight in a microplate in a final volume of 100 µl
culture medium per well at 37°C and 5% CO2. Assays were started
by the addition of 10 µl of WST-1 to the well. Some cells were further
incubated at 37°C with 21% O2 and the survival rates were
measured after 1-2 hours (Roche Molecular Biochemicals, Quebec, Canada).
Rho protein subcellular distribution
Following hypoxic incubation, cell monolayers were lyzed on ice for 10
minutes with buffer A (10 mM Hepes-Tris pH 7.4, 10 mM KCl, 1.5 mM
MgCl2, 1 mM dithiothreitol and protease inhibitors). Cells were
further disrupted with a Polytron (3x15 seconds) and centrifuged for 10
minutes at 1000 g at 4°C. Aliquots of clarified
post-nuclear supernatants (PNS) were conserved at 80°C while the
rest of the supernatants were centrifuged at 100,000 g for 1
hour at 4°C. Pellets (crude membranes) were resuspended with a minimum
volume of buffer A and supernatants (cytosol) were conserved at
80°C. Proteins of PNS, cytosol and membranes were quantified by the
micro BCA method. Identical amounts of protein were solubilized in Laemmli
buffer (62.5 mM Tris/HCl pH 6.8, 10% glycerol, 2% SDS, 5%
ß-mercaptoethanol and 0.00625% bromophenol blue), boiled for 4 minutes
and then analyzed by SDS-PAGE as described below.
Western blot analysis
Proteins were separated by 12.5% polyacrylamide gel electrophoresis in the
presence of SDS (SDS-PAGE) followed by semi-dry transfer onto polyvinylidene
difluoride membranes (PVDF) (Roche Molecular Biochemicals, Quebec, Canada)
using standard procedures. The membranes were blocked overnight at 4°C in
5% powdered nonfat milk in Tris-buffered saline (TBS) (50 mM Tris pH 7.4, 150
mM NaCl) containing 0.1% Tween 20 (TBS-T). Membranes were washed three times
for 15 minutes in TBS-T. The PVDF membranes were incubated with primary
antibodies diluted 1:1000 for RhoA, Cdc42, RhoGDI, RhoB (Santa Cruz
Biotechnology, Santa Cruz, CA), Rac1 (Transduction Laboratories, Lexington,
KY) and HIF-1 (Novus Biologicals, Littleton, CO) in TBS-T, 1% BSA and
0.03% NaN3 for 1 hour at 37°C. Membranes were washed three
times for 15 minutes each and incubated for 1 hour at room temperature with
horseradish peroxidase-conjugated anti-rabbit or anti-mouse antibodies
(Jackson Immunoresearch Laboratories, West Grove, PA) diluted 1:5000 in TBS-T
containing 5% milk powder. PVDF membranes were washed and antigens detected
using the western blot chemiluminescence reagent plus (NENTM Life Science
Products, Boston, MA). Blots were exposed to Fuji films and the autoradiograms
were scanned with a Personal Densitometer (Molecular Dynamics, Sunnyvale,
CA).
Metabolic labelling and immunoprecipitation
Prior to hypoxia, cells were washed twice with PBS and incubated for 1 hour
at 37°C in a 5% CO2 atmosphere in a methionine-cysteine-free
RPMI 1640 medium. Fifty µCi/ml of 35S-Met/Cys (0.043 µM)
(ICN, Costa Mesa, CA) were added to the medium and cells were kept in hypoxia
or in normoxia for 4 hours. After labelling, cells were lyzed in buffer A and
PNS were fractionated into cytosol and membranes as described above. Aliquots
of each fraction (100 µg of protein) were solubilized in 1 ml of buffer B
(0.1% SDS, 1% NP-40, 0.5% deoxycholate, 50 mM Tris-HCl pH 7.5, 150 mM NaCl),
then were precleared by incubation for 1 hour at 4°C with 20 µl of
protein G-Sepharose beads (50% in PBS) (Amersham Pharmacia Biotech, Uppsala,
Sweden). After centrifugation at 1000 g for 3 minutes at
4°C, supernatants were immunoprecipitated by overnight incubation with 1
µg anti-RhoA at 4°C with agitation. Twenty µl of protein G-Sepharose
beads were added to the immune complexes for 2 hours at 4°C with
agitation. Immunoprecipitated RhoA was pelleted by centrifugation at 1000
g for 3 minutes at 4°C. Following three washings of the
beads with buffer B, proteins were solubilized with Laemmli buffer, boiled for
4 minutes, and centrifuged at 1000 g for 2 minutes.
Immunoprecipitated proteins were analyzed by SDS-PAGE as described above and
detected by autoradiography to identify the cellular distribution of newly
synthesized RhoA.
Purification of recombinant fusion proteins
Recombinant plasmids of the expression vector pGEX-2T containing cDNAs
encoding the fusion proteins (i) glutathione S-transferase-toxin C3
transferase from Clostridium botulinum (GST-C3) (gift of Dr A. Hall,
University College London, London, UK); (ii) glutathione S-transferase-Rho
binding domain of PAK1 (GST-PBD) (gift of Dr G. M. Bokoch, The Scripps
Research Institute, CA); and (iii) glutathione S-transferase-Rho binding
domain of rhotekin (GST-RBD) (gift of Dr M. Schwartz, The Scripps Research
Institute) were expressed in Escherichia coli. Fusion proteins were
purified from isopropyl-ß-D-thiogalactopyranoside-induced
exponential-phase bacterial cultures by standard procedures. For C3, the GST
moiety of the fusion protein was removed by incubating GST-C3, while bound to
glutathione-Sepharose beads, for 4 hours at room temperature with thrombin
protease (Pharmacia, Uppsala, Sweden). Contaminating thrombin was removed by
incubation with p-aminobenzamidine linked to agarose beads (Sigma, St Louis,
MO). Protein concentration was measured using the Bradford assay (Pierce,
Rockford, IL). To verify the purity of recombinant C3 toxin, aliquots of each
fraction were analyzed by SDS-PAGE and stained with Coomassie blue. For
GST-PBD and GST-RBD, the fusion proteins were collected by incubation with
glutathione-Sepharose 4B beads (Amersham Pharmacia Biotech, Uppsala, Sweden)
for 1 hour at 4°C following procedures previously described
(Benard et al., 1999;
Ren et al., 1999
). Fusion
proteins still bound to glutathione-Sepharose 4B beads were then washed,
resuspended in buffer C (50 mM Tris pH 7.4, 1% Triton X-100, 0.5% sodium
deoxycholate, 0.1% SDS, 150 mM NaCl, 10 mM MgCl2 and protease
inhibitors), aliquoted and conserved at 80°C. To determine the
amount of purified protein, 20 µl of bead suspension were analyzed by
SDS-PAGE and Coomassie Blue staining.
Transfection with RhoA mutants
Caki-1 cells (60% confluent/100 mm) were transfected with vector (pcDNA3)
or pcDNA3-RhoAV14-Myc (dominant-active RhoA mutant with a Gly14Val mutation)
(gift from W. Moolenaar, The Netherlands Cancer Institute, Amsterdam, The
Netherlands). Transfection was carried out with Lipofectamine (Gibco-BRL Life
Technologies) as a carrier using cells that had been serum-starved for 1 hour
then incubated with vectors for 5 hours at 37°C in a humidified atmosphere
of 5% CO2. To permit cell recuperation, the mixture was replaced by
complete McCoy's medium containing 10% FCS and incubated overnight. The levels
of RhoA mutant in the transfected cells were determined in parallel
experiments by immunodection of cell lysates with an anti-Myc epitope
monoclonal antibody (SantaCruz Biotechnology, Santa Cruz, CA).
Pull-down assays
Caki-1 cells experienced hypoxia for 6 hours. Cells were then washed twice
with PBS and protein was extracted in buffer C for 10 minutes on ice. Lysates
were centrifuged at 1000 g for 10 minutes at 4°C. Soluble
lysates were incubated with 20-30 µg of GST-RBD or GST-PBD bound to
glutathione-Sepharose 4B beads for 45-60 minutes at 4°C. Samples were
centrifuged at 800 g for 3 minutes at 4°C. The pelleted
proteins were solubilized in Laemmli sample buffer and heated at 95°C for
4 minutes. Proteins bound to GST-RBD and GST-PBD were separated by SDS-PAGE
and transferred onto PVDF membranes for immunodetection with Cdc42 and RhoA
antibodies. Data are expressed as the percentage of relative activity compared
with values found in normoxic cells.
Measurement of ROS and treatment with DPI
Intracellular ROS production was assessed using
2',7'-dichlorofluorescein (DCFH) (Molecular Probes, Eugene, OR). A
stock solution (10 mM) was prepared in 100% ethanol, aliquoted and kept at
80°C. A DPI (Sigma) solution (10 mM) was freshly prepared in
dimethyl sulfoxide (DMSO) for use in each experiment. Cells experienced
normoxia or hypoxia in the presence of 10 µM DCFH with or without 10 µM
DPI. Cells were lyzed in 50 mM Tris pH 7.4 containing 0.1% Triton X-100.
Fluorescence was measured using excitation at 485 nm and emission at 530 nm.
Following treatments in the absence or presence of DPI, cells were lyzed and
proteins separated on 12.5% SDS-PAGE and transferred onto PVDF membanes.
Immunodetection with Cdc42 and RhoA antibodies was carried out to determine
the effect of ROS inhibition on Rho GTPase expression.
Treatments with C3 exotoxin
To permit entrance of the toxin into cells, 50 µg/ml of C3 toxin was
added to confluent cells for 24 hours incubation. Afterwards, cells were
placed under hypoxic conditions for 4 hours. The efficiency of
ADP-ribosylation caused by C3 on RhoA protein was observed on 12.5% SDS-PAGE
by shift mobility of the RhoA protein after immunodetection with RhoA
antibody.
RNA isolation and RT-PCR
After hypoxia, monolayers of cells were lyzed with TRIzol reagent
(Gibco-BRL Life Technologies) using the manufacturer's directions for total
cellular RNA extraction. RNA was quantitated by absorbance at 260 nm. Using a
MasterAmp kit (Epicentre Technologies, Madison Technologies, Madison, WI), 1
µg RNA was amplified by reverse transcription polymerase chain reaction
(RT-PCR). cDNAs were amplified in a 50 µl reaction mixture containing 3 mM
MgCl2, 0.5 mM MnSO4, RT-PCR buffer 1x, MasterAmp
PCR Enhancer 1x, 400 µM dNTP mix, 0.25 µM of each primer, and 2.5
units of RetroAmp RT DNA Polymerase. The PCR primers used for RhoA, Cdc42,
HIF-1, VEGF and
-tubulin cDNA amplification are listed in
Table 1. The reverse
transcription was performed at 60°C for 20 minutes. The PCR conditions for
RhoA and Cdc42 were 25 cycles at a denaturation temperature of 94°C for 30
seconds, annealing at 58°C for 1 minute and extension at 72°C for 1
minute. For HIF-1
, 25 cycles were carried out with 30 seconds at
95°C, 1 minute at 55°C and 2.5 minutes at 72°C. VEGF was amplified
for 30 cycles at 94°C for 1 minute, 55°C for 2 minutes and 72°C
for 3 minutes. Finally, primers for
-tubulin were designed for this
study using MacVector 7.0 software (Oxford Molecular, Madisson WI) and
amplified for 25 cycles at 94°C for 30 seconds, 58°C for 30 seconds
and 72°C for 30 seconds after a denaturation initial at 94°C for 2
minutes. A final extension of 7 minutes at 72°C was carried out. PCR
fragments were analyzed on 1.8% agarose gels stained with ethidium
bromide.
|
Statistical analysis
Data obtained from the densitometric analysis were expressed as the ratios
of immunodetected proteins by western blot under hypoxic conditions to those
detected under normoxic conditions. RT-PCR products stained with ethidium
bromide were also quantified by densitometric analysis. All data were
expressed as means ± s.e.m. for at least three separate experiments and
analyzed with the Student's t-test. The only significant differences
(P<0.05 or P<0.1) are indicated in the figures by an
asterisk.
![]() |
Results |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
|
Hypoxia increases Rho protein expression in renal cell carcinoma
Cdc42, Rac1 and RhoA protein expressions were all upregulated in RCC under
hypoxia (Fig. 2A).
Densitometric analysis of immunodetected Rho GTPases showed that their
expression reached a maximum level then gradually returned to control values
after 6 hours (Fig. 2B). One
hour of hypoxia was sufficient to significantly upregulate Cdc42 protein
expression by twofold. Maximum increase of Rac1 protein expression (threefold)
was observed after 2 hours of hypoxia while RhoA expression displayed the
greatest stimulation (fourfold) after 4 hours of hypoxia. These kinetics
showed that the individual Rho protein expressions are sequentially
stimulated; Cdc42, Rac1 then RhoA. To evaluate the specificity of hypoxia to
Rho protein expression, the effect of this cellular stress on related proteins
was examined. We found by Western blot analysis that levels of an inhibitor of
Rho activity, the GDP-dissociation inhibitor of Rho proteins (RhoGDI), and
RhoB protein expression were unaffected by the drop of O2
concentration (Fig. 2). This
upregulation of Rho protein expression during hypoxia did not seem to involve
Ras pathways because N-Ras and K-Ras expressions remained stable in Caki-1
cells (data not shown). Thus, hypoxia upregulated the expression of a subset
of Rho GTPases. This observation is supported by a recent report showing that
Rac1 is activated in hypoxia (Hirota and
Semenza, 2001). Consequently, Cdc42 and RhoA were the focus of
next experiments to investigate their expression and to study the molecular
mechanisms used by Rho proteins to regulate hypoxia responses.
|
Effect of hypoxia on subcellular distribution of Cdc42 and RhoA
proteins
To gain a better understanding of the Rho functions overexpressed during
hypoxia, cells were fractionated into soluble (cytosol) and crude membrane
fractions since active Rho GTPases are membrane-bound. Quantification by
densitometry indicated that Cdc42 expression significantly increased by
1.8-fold at membranes between 1 and 2 hours of hypoxia
(Fig. 3A). For RhoA,
significantly enhanced expression at membranes was observed between 2 and 6
hours but was maximal at 4 hours for a 3.9-fold increase
(Fig. 3B). A lesser increase in
Cdc42 and RhoA expression was noted in the soluble fraction at the time of
their overexpression at membranes. In contrast, we did not observe the effect
of this cellular stress on RhoB protein expression into membrane fraction or
on RhoGDI expression in soluble fraction (data not shown). This preferential
localization at membranes suggested that Cdc42 and RhoA were active and that
this may enable them to bind to the effectors that mediate their action.
|
RhoA synthesized during hypoxia rapidly translocates to
membranes
The increased expression of Rho proteins during hypoxia could be explained
either by stimulation of synthesis or decreased protein turnover. To
distinguish between these possibilities, metabolic labelling was performed to
visualize newly synthesized protein during hypoxia. The contribution of
synthesis to RhoA upregulation during hypoxia was analyzed by
immunoprecipitation of the GTPase from the soluble and membrane fractions.
Interestingly, autoradiography and their quantification by densitometry showed
that synthesized RhoA increased significantly by 2.2-fold in membranes but
only 1.4-fold in the soluble fraction during hypoxia when compared with
normoxia values (Fig. 4).
However, we observed by western blot analysis that the majority of RhoA was
present in soluble fractions (Fig.
4A). Although the same amount of proteins from each fraction were
used for [35S]-labelled RhoA immunoprecipitation, only a weak
signal for RhoA was present in soluble fractions compared with that in
membranes. This could be explained by the inability of RhoA antibody to
immunoprecipitate the protein due to its interaction with RhoGDI in soluble
fractions. These data clearly indicate that hypoxia induces the synthesis of
RhoA and that this newly synthesized GTPase translocated to membranes. From
these findings, we can postulate that upregulated Rho GTPases arise, at least
in part, from new synthesis under hypoxia.
|
Effect of hypoxia on activation of Cdc42 and RhoA
To efficiently interact with effectors, active Rho proteins must bind GTP.
To assess the activation states of Cdc42 and RhoA during hypoxia, pull-down
assays were performed which used GST-fusion proteins conjugated to the
Rho-binding domains of Rho effectors in order to selectively precipitate
active forms of Rho proteins. We used GST-RBD containing the Rho-binding
domain of serine/threonine protein kinase PAK, which is a specific effector of
Cdc42 and Rac1, as well as GST-RBD carrying the Rho-binding domain of the
effector rhotekin, used for RhoA. The results displayed in
Fig. 5 show that hypoxia
significantly increased the activation of Rho GTPases in RCC. The peak of
Cdc42 total expression at 1 hour of hypoxia was also accompanied by an
increase in the GTP-bound form of this protein
(Fig. 5A). This activation
diminished after 2 hours, and had dropped to control values after 6 hours of
hypoxia. Similarly, levels of activated RhoA gradually increased during
hypoxia to reach 2.4-fold after 4 hours, corresponding to the time of its
highest level of total expression, then diminished to 1.6-fold after 6 hours
of hypoxia (Fig. 5B). However,
the increase in the levels of GTP-bound forms of Rho GTPases were lower than
the increase in protein expression seen in cell lysates
(Fig. 2). The gradual increase
in RhoA and Cdc42 activation under hypoxia agrees with the observation that
upregulated Rho GTPases are also found at the membranes
(Fig. 3). These results suggest
that hypoxia induces new synthesis of Rho GTPases which translocate to
membranes where they are activated.
|
Hypoxia increases RhoA mRNA expression in renal cell carcinoma
To further characterize RhoA GTPase during hypoxia, we next studied the
effect of O2 tension on its mRNA expression. Optimal PCR
amplification conditions were determined by varying cycle number. We found
that 25 cycles were necessary to obtain maximal difference of the
amplification reactions for RhoA and Cdc42
(Fig. 6A). Under these
conditions, 2 hours of hypoxia induced significantly the highest RhoA mRNA
expression (Fig. 6B). Here, the
peak of mRNA expression occurred before that of protein expression at 4 hours
(Fig. 2). Cdc42 mRNA was also
slightly induced by 1 hour of hypoxia but not significantly
(Fig. 6B). By contrast, as a
negative control, the mRNA level of -tubulin was examined and found
unaffected by the reduced O2 tension. These results suggest that an
increased RhoA mRNA level enhances protein synthesis upon hypoxia.
|
Hypoxia stimulates ROS production in renal cell carcinoma
After having characterized the upregulation of Rho proteins during hypoxia,
we were interested in identifying upstream events triggering Rho
overexpression. Production of ROS had been proposed as a possible sensor
mechanism by which cells could detect the decrease in [O2]
(Chandel et al., 1998). We thus
monitored ROS levels in Caki-1 cells as a consequence of hypoxia. ROS
production as measured by conversion of DCFH into oxidized DCF demonstrated a
rapid and significantly increase (2.2-fold) following 1 hour of hypoxia
(Fig. 7A). Even after 30
minutes of hypoxia, ROS production had increased 1.4-fold, whereas ROS levels
returned to control values of normoxia after 2 hours of hypoxia. Furthermore,
the fluorescence of oxidized DCF was measured in the presence of DPI, an
inhibitor of ROS production which interferes with electron transport in
flavin-containing systems including NADPH oxidase and mitochondrial complex I.
In normoxia, this inhibitor did not affect ROS production, whereas DPI
significantly inhibited the increase in DCF fluorescence obtained after 1 hour
of hypoxia (Fig. 7B). Our
results are in agreement with previous studies also examining ROS production
during hypoxia (Chandel and Schumacker,
2000
; Duranteau et al.,
1998
; Grishko et al.,
2001
).
|
Inhibition of ROS production prevents Rho upregulation during
hypoxia
To determine whether ROS are involved in the upregulation of Rho GTPases
during hypoxia, Cdc42 and RhoA protein levels were studied in the presence of
the ROS inhibitor DPI. As expected, Cdc42
(Fig. 8A) and RhoA
(Fig. 8B) expression was
stimulated after 1 to 4 hours of hypoxic treatments but in the presence of the
ROS inhibitor this upregulation was prevented. These results support the
hypothesis that hypoxia stimulates ROS production, which is required to
increase Rho expression. Thus, our kinetic analysis shows that hypoxia-induced
ATP depletion (0.5 hour) is followed by ROS production (1 hour), which
ultimately triggers Rho expression (1-4 hours).
|
HIF-1 and VEGF mRNA levels are increased in renal cell
carcinoma exposed to hypoxia
Several studies have demonstrated stabilization of HIF-1 and
upregulation of VEGF in hypoxic conditions. To examine the role of Rho protein
induction during hypoxia, we studied their involvement in HIF-1
and
VEGF expression. Amplification conditions for the primer sets were determined
by RT-PCR analysis. Fig. 9A
shows that an amplification of HIF-1
for at least 25 cycles was
necessary to observe a difference between normoxia and hypoxia. The primer
pair used for VEGF is known to generate three isoforms (VEGF145,
VEGF165, VEGF189). In Caki-1 cells, 30 cycles were
necessary to obtain 2 isoforms at 165 and 189 amino acids and, under normoxia,
little VEGF was observed (Fig.
9A). In Caki-1 cells under hypoxia, mRNA levels of HIF-1
and VEGF have already occurred by 2 hours but were maximal at 4 hours
(Fig. 9B). Expression of
VEGF165 mRNA seemed to increase more than VEGF189 and
has been shown to be secreted in large quantities in the kidney
(Robert et al., 2000
). As
negative control,
-tubulin was amplified at same time
(Fig. 9B). Immunoprecipitation
followed by immunodetection with HIF-1
antibody in nuclear fractions
showed that the level of this transcription factor became increased and
remained stable between 4 and 8 hours of hypoxia
(Fig. 9C). Secreted and
cellular VEGF was undetectable in Caki-1 cells by western blot analysis (data
not shown). These results suggested that the upregulation of Rho GTPases
expressions during hypoxia, since they occurred earlier, could be upstream of
HIF-1
mRNA and protein induction.
|
Overexpressed RhoA stimulates HIF-1 and VEGF mRNA levels in
normoxia
We determined whether Rho GTPase overexpression could enhance HIF-1
and VEGF mRNA levels. RCC cells were transfected with dominant-active RhoA
(RhoAV14). This particular protein was tagged with Myc and the expression of
mutated RhoA was confirmed with a Myc antibody (data not shown). Cells
expressing dominant-active RhoA showed significantly enhanced mRNA levels of
HIF-1
(2.2-fold) and VEGF (1.8-fold) in normoxic conditions
(Fig. 10A,B). No significant
difference was observed between control cells and cells incubated with vector.
As a negative control, the level of
-tubulin was examined
(Fig. 10C). These results
suggest that the level of activated RhoA contributes to stimulate HIF-1
and VEGF mRNA levels.
|
Toxin C3 blocks HIF-1 and VEGF overexpression during
hypoxia
To understand the role of Rho protein upregulation during hypoxia, we used
toxin C3, which selectively inhibits RhoA, RhoB and RhoC activation by
ADP-ribosylation. This toxin has little effect on Cdc42 and Rac1 activity.
Following hypoxia, cells were fractionated into soluble and crude membrane
fractions. To evaluate toxin activity, ADP-ribosylation efficiency was
examined by western blot analysis and shift in mobility of RhoA protein. We
observed a shift mobility of all RhoA due to ADP-ribosylation on
Asn41 in soluble and membranes fractions
(Fig. 11A). The weak signal of
RhoA immunodetected when cells were treated with toxin may be explained by a
lower recognition of ADP-ribosylated protein by RhoA antibody. Hypoxia
increased RhoA level mainly in the membranes, as shown above
(Fig. 3).
|
We next studied whether RhoA inhibition by C3 toxin might affect
HIF-1 and VEGF mRNA levels. Incubation with C3 had no effect on
HIF-1
and VEGF mRNA levels under normoxia conditions
(Fig. 11B). More notably, C3
toxin significantly abolished the induction of HIF-1
and VEGF mRNA
observed during hypoxia (Fig.
11B). Again, the level of
-tubulin mRNA was analyzed as
negative control and found unaffected by hypoxia
(Fig. 11B). To further
validate the RhoA contribution on HIF-1
expression, we next studied the
effect of C3 on HIF-a protein amount in cell lysates
(Fig. 11C). We observed a
significant difference upon hypoxic stress on HIF-1
expression
(twofold), which indicates an accumulation of the protein either due to an
increase synthesis or resulting from of HIF-1
stabilization. A
pre-treatment with toxin C3 has no effect on HIF-1
protein expression
in normoxic conditions. As expected, toxin C3 prevented the accumulation of
HIF-1
protein by 84% under hypoxia, which suggests that new synthesis
was necessary to increase HIF-1
level
(Fig. 11C). Hypoxia and
treatment with this toxin unaffected
-tubulin protein expression. These
results clearly demonstrate that RhoA upregulation and activation occurring
during hypoxia are upstream and contribute to HIF-1
mRNA and protein
accumulation in RCC.
![]() |
Discussion |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
RCC were exposed to 1% O2, a concentration frequently found
during ischemia/hypoxia events in vivo, and appeared to tolerate up to 6 hours
of hypoxia, although intracellular ATP levels became diminished by 90%. This
tolerance to ATP depletion might be attributable to molecular compensation,
allowing cells to adapt to hypoxia by use of proteins known to play a pivotal
role in actin polymerization. Towards this end, we have found that hypoxia
increases expression and activity of such key proteins as Cdc42, Rac1 and RhoA
in RCC Caki-1. In addition, expression of these proteins was also increased in
other cell lines tested: in two renal cell lines (OK and Caki-2) and in a
human microvascular endothelial cell line (HMEC-1) (data not shown). Recently,
another study also demonstrated a rise in Rac1 activity after 2 hours of
hypoxia, supporting our findings (Hirota
and Semenza, 2001). However, this study did not observe a role of
RhoA for HIF-1
induction. The difference in results could be explained
by the cell lines used and probably by the methodological approach. The study
by Hirota and Semenza used a reporter gene assay in cells co-transfected with
the luciferase gene containing an HIF-1-dependent HRE in its promoter and with
the dominant-negative Rho in hepatocarcinoma cells whereas we have observed
the rise of RhoA expression in Caki-1 cell lysates directly without
transfection by western blot. In our study, upregulation of Rho protein
expression during hypoxic conditions occurred between 1 and 4 hours and was
found to be sequential, starting with Cdc42, then Rac1, and finally RhoA
(Fig. 2). Others studies have
reported that activation of Cdc42 by growth factors or other stimuli led to
Rac1 activation, which in turn activated RhoA in Swiss 3T3 fibroblasts
(Nobes and Hall, 1995
;
Ridley and Hall, 1992
). At
this point, however, it remains to be established whether this cascade
reflects a linear activation pathway of Rho GTPases in RCC or whether Cdc42,
Rac1 and RhoA are stimulated by separate pathways during hypoxia.
To gain a better understanding of the mechanisms involved in RhoA upregulation, we investigated its expression at different levels during hypoxia. Our results demonstrate an induction of RhoA transcript level by hypoxia (Fig. 6). Immunoprecipitation experiments show that protein synthesis of RhoA is induced in RCC under hypoxia (Fig. 4). Subcellular fractionation indicated that the increased amount of Rho proteins is mainly located at membranes (Fig. 3). Moreover, determination of the activation states of Cdc42 and RhoA under hypoxia confirmed that both proteins were present as GTP-bound forms (Fig. 5). The levels of protein expression and activation status of Rho GTPases returned to control levels after 6 hours of hypoxia (Figs 2 and 5). It is thus concluded that Rho GTPases are transiently expressed during hypoxia and that they are subsequently targeted for rapid proteolysis.
Analysis of events upstream of HIF-1 suggests a crucial role for ROS
generation upon hypoxia via mitochondria as a sensor mechanism of the oxygen
level, and that ROS may be involved in stabilizing HIF-1
(Chandel and Schumacker, 2000
;
Li and Jackson, 2002
;
Duranteau et al., 1998
;
Pearlstein et al., 2002
;
Chandel et al., 1998
). In cells
depleted of mitochondrial DNA or in the presence of antioxidants such as
ebselen or pyrrolidinedithio-carbamate, this response is lost
(Chandel and Schumacker, 2000
;
Duranteau et al., 1998
;
Grishko et al., 2001
). During
mitochondrial respiration, O2 consumption generates energy in the
form of ATP, and it has been suggested that the diminution of O2
concentration in hypoxic conditions may result in a drop in ATP synthesis that
will favour ROS production (Chandel and
Schumacker, 2000
). This agrees with our data that the peak of ROS
production occurs after 60 minutes of hypoxia in RCC when the ATP amount has
already reached its lowest level (Figs
1,
7). Since ROS generation is
concomitant or earlier to Rho GTPase upregulation in hypoxia, we thus studied
the effect of DPI on Rho protein expression and demonstrate that ROS
generation occurs upstream of and is essential to Rho upregulation
(Fig. 8).
The signal transduction mechanisms by which ROS stimulate Rho expression in
response to hypoxia remain largely unknown. Although a large number of
signaling pathways appear to be regulated by ROS, most studies address this
question using exogenous oxidants rather than the cellular ROS that would be
generated by growth factors, cytokines or hypoxia. There is growing evidence,
however, that intracellular redox regulation may occur at multiple levels. For
example, Ras is a direct target of ROS and thus may be responsible for sensing
the intracellular redox status (Lander et
al., 1996). This effect appears to involve PI-3K activity, which
is known to be downstream of Ras, since its inhibition by wortmannin prevents
Rac1 and HIF-1 transcriptional induction in response to hypoxia
(Hirota and Semenza, 2001
;
Jiang et al., 2001
). Other
studies have also demonstrated that PI-3K may activate Rho proteins such as
RhoA and Rac1 in human fibrosarcoma cells
(Gupta et al., 2001
).
Together, these data suggest that PI-3K activity is an upstream event
mediating Rho GTPases activation by ROS during hypoxia in RCC.
VEGF is expressed at high levels in several cancers, including RCC, and has
been identified as one of the most potent inducers of tumour-associated
angiogenesis (Brenchley, 1998).
The hypoxic conditions found in tumors have been shown as upregulating VEGF in
vitro and in vivo. The control of VEGF gene expression is complex but it is
thought that, in hypoxic regions within solid tumours, the expression of VEGF
is regulated in part by HIF-1
(Maxwell et al., 1997
). In
addition to binding sites for HIF-1 within the promoter region of VEGF genes,
studies have also indicated that increased VEGF mRNA stability occurred during
hypoxia and was mediated via specific sequences found in the 3'
untranslated region (Levy et al.,
1996
). Furthermore, an active internal ribosomal entry site (IRES)
encoding an alternative initiation site was recently reported to ensure
efficient translation of VEGF mRNA during hypoxia
(Stein et al., 1998
). To
better understand the function of early Rho expression and activation in
hypoxic conditions, we thus evaluated the kinetics of HIF-1
and VEGF
mRNA induction and showed that their upregulation occurred after that of RhoA
mRNA in RCC (Figs 6,
9).
To validate this finding, and to determine whether Rho protein upregulation
modulates HIF-1 and VEGF expression, we firstly transfected cells with
a dominant-active RhoA cDNA. In RhoAV14-transfected RCC, HIF-1
and VEGF
mRNA levels are induced but less than by hypoxia
(Fig. 10). Interestingly,
these data suggest that mRNA expression of HIF-1
and VEGF are dependent
on RhoA level. This lower induction of HIF-1 and VEGF in RhoAV14-transfected
cells could be partially explained by the efficiency of transfection, or more
likely by a contribution of Cdc42 and Rac1 to the induction of HIF-1
and VEGF during hypoxia. Secondly, supporting this conclusion for a key role
of RhoA in hypoxia-induced HIF-1
and VEGF mRNA upregulation, these
inductions are abolished when cells are incubated with toxin C3
(Fig. 11). Exotoxin C3
irreversibly inhibits RhoA by ADP-ribosylation at Asn41 and C3
toxin also acts on RhoB and RhoC, but the former is not modulated during
hypoxia while RhoC is not immunodetected in Caki-1 cells (data not shown).
More important, the toxin also prevents the accumulation of HIF-1
protein during hypoxia by 84% (Fig.
11). From the results showing that dominant-active RhoA induces
HIF-1
and VEGF mRNA in normoxia and that toxin C3 blocks the induction
of HIF-1
mRNA and protein and VEGF mRNA stimulation by hypoxia, we
conclude that RhoA is necessary for HIF-1
upregulation at low
O2 tension in RCC.
This new pathway involving RhoA in the upregulation of HIF-1 is
likely a complementary process to the stabilization of the protein during
hypoxia. Under normoxic conditions, HIF-1
is recognized by the
tumour-suppressor protein pVHL. This interaction promotes a rapid degradation
of HIF-1
by the proteasome (Ivan et
al., 2001
). pVHL binds the oxygen degradation domain of
HIF-1
through conserved proline residues. In the presence of iron and
oxygen, proline 402 and proline 564 are hydroxylated. Similarly, hydroxylation
of asparagine residues in the transactivation domain C-TAD occurs in nucleus
under normoxia (Lando et al.,
2002
). When the oxygen concentration diminishes, the hydroxylation
of proline and asparagine residues is stopped and pVHL-HIF-1
interaction is lost allowing an accumulation of HIF-1
. As expected, the
basal level of HIF-1
in RCC is barely detectable in normoxia
(Fig. 9). Thus, if the
efficiency of HIF-1
mRNA translation is unaffected by hypoxia and that
the protein is protected against proteolysis, the accumulation of HIF-1
could be a gradual process. However, the upregulation of HIF-1
mRNA and
protein by RhoA accelerates the accumulation of the transcriptional factor
facilitating cell adaptation to the hypoxic environment. Since RhoA was
induced in others renal (OK, Caki-2) and endothelial (HMEC-1) cell lines (data
not shown) and that it has been reported that Rac1 is required to induce
HIF-1
protein expression in hepatocarcinomas cells (Hep3B)
(Hirota and Semenza, 2001
), it
could be postulated that Rho participitates in HIF-1 induction in several cell
types.
The sequence of events presented in
Fig. 12 summarizes the major
findings of our study. Hypoxia induces a drop in cellular ATP that is followed
by ROS production. Activation of Rho GTPases is dependent on and occurs
downstream of ROS production. Our results also demonstrate that Cdc42, Rac1
and RhoA are upregulated sequentially by hypoxia but it remains to be
established whether there is a hierarchy of activation in Rho GTPases.
Finally, RhoA activation is necessary for HIF-1 and VEGF production in
renal cell carcinoma since they are inhibited by C3 toxin.
|
![]() |
Acknowledgments |
---|
![]() |
References |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Benard, V., Bohl, B. P. and Bokoch, G. (1999).
Characterization of rac and cdc42 activation in chemoattractant-stimulated
human neutrophils using a novel assay for active GTPases. J. Biol.
Chem. 274,13198
-13204.
Boivin, D. Bilodeau, D. and Béliveau, R. (1996). Regulation of cytoskeletal functions by Rho small GTP-binding proteins in normal and cancer cells. Can. J. Physiol. Pharmacol. 74,801 -810.[CrossRef][Medline]
Bokoch, G. M. (2000). Regulation of cell function by Rho family GTPases. Immunologic. Res. 21,139 -148.[Medline]
Boussat, S., Eddahibi, S., Coste, A., Fataccioli, V., Gouge, M., Housset, B., Adnot, S. and Maitre, B. (2000). Expression and regulation of vascular endothelial growth factor in human pulmonary epithelial cells. Am. J. Physiol. 279,L371 -L378.
Brenchley, P. E. C. (1998). Antagonising the expression of VEGF in pathological angiogenesis. Exp. Opin. Ther. Patents 8,1695 -1706.
Catron, T., Mendiola, M. A., Smith, S. M., Born, J. and Walker,
M. K. (2001). Hypoxia regulates avian cardiac Arnt and
HIF-1lpha mRNA expression. Biochem. Biophys. Res.
Commun. 282,602
-607.[CrossRef][Medline]
Chandel, N. S. and Schumacker, P. T. (2000).
Cellular oxygen sensing by mitochondria: old questions, new insight.
J. Appl. Physiol. 88,1880
-1889.
Chandel, N. S., Maltepe, E., Goldwasser, E., Mathieu, C. E.,
Simon, M. C. and Schumacker, P. T. (1998). Mitochondrial
reactive oxygen species trigger hypoxia-induced transcription.
Proc. Natl. Acad. Sci. USA
95,11715
-11720.
Dachs, G. U. and Tozer, G. M. (2000). Hypoxia modulated gene expression: angiogenesis, metastasis and therapeutic exploitation. Eur. J. Cancer 36,1649 -1660.[CrossRef][Medline]
Dagher, P. C. (2000). Modeling ischemia in vitro: selective depletion of adenine and guanine nucleotide pools. Am. J. Physiol. 279,C1270 -C1277.
Duranteau, J., Chandel, N. S., Kulisz, A., Shao, Z. and
Schumacker, P. T. (1998). Intracellular signaling by reactive
oxygen species during hypoxia in cardiomyocytes. J. Biol.
Chem. 273,11619
-11624.
Ferrara, N., Houck, K., Jakeman, K. and Leung, D. (1992). Molecular and biological properties of the vascular endothelial growth factor family of proteins. Endocr. Rev. 13,18 -32.[Medline]
Firth, J. D., Ebert, B. L. and Ratcliffe, P. J. (1994). Oxygen-regulated control elements in the phosphoglycerate kinase 1 and lactate dehydrogenase A genes: similarities with the erythropoietin 3' enhancer. Proc. Natl. Acad. Sci. USA 91,6496 -6500.[Abstract]
Gopalakrishnan, S., Raman, N., Atkinson, S. J. and Marrs, J. A. (1998). Rho GTPase signaling regulates tight junction assembly and protects tight junctions during ATP depletion. Am. J. Physiol. 275,C798 -C809.[Medline]
Gothié, E., Richard, D. E., Berra, E., Pagès, G.
and Pouysségur, J. (2000). Identification of
alternative spliced variants of human hypoxia-inducible factor-1alpha.
J. Biol. Chem. 275,6922
-6927.
Grishko, V., Solomon, M., Breit, J. F., Killilea, D. W., Ledoux,
S. P., Wilson, G. L. and Gillespie, M. N. (2001). Hypoxia
promotes oxidative base modifications in the pulmonary artery endothelial cell
VEGF gene. FASEB J. 15,1267
-1269.
Gunningham, S. P., Currie, M. J., Han, C., Turner, C., Scott, P.
A. E., Robinson, B. A., Harris, A. L. and Fox, S. B. (2001).
Vascular endothelial growth factor-B and vascular endothelial growth factor-C
expression in renal cell carcinomas: regulation by the von Hippel-Lindau gene
and hypoxia. Cancer Res.
61,3206
-3211.
Gupta, S., Stuffrein, S., Plattner, R., Tencati, M., Gray, C.,
Whang, Y. E. and Stanbridge, E. J. (2001). Role of
phosphoinositide 3-kinase in the aggressive tumor growth of HT1080 human
fibrosarcoma cells. Mol. Cell. Biol.
21,5846
-5856.
Hall, A. (1998). Rho GTPases and the actin
cytoskeleton. Science
279,509
-514.
Hirota, K. and Semenza, G. L. (2001). Rac1
activity is required for the activation of hypoxia-inducible factor 1.
J. Biol. Chem. 276,21166
-21172.
Houck, K. A., Ferrara, N., Winer, J., Cachianes, G., Li, B. and Leung, D. W. (1991). The vascular endothelial growth factor family: identification of a fourth molecular species and characterization of alternative splicing of RNA. Mol. Endocrinol 5,1806 -1814.[Abstract]
Ivan, M., Kondo, K., Yang, H., Kim, W., Valiando, J., Ohh, M.,
Salic, A., Asara, J. M., Lane, W. S. and Kaelin, W. G., Jr
(2001). HIFalpha targeted for VHL-mediated destruction by proline
hydroxylation: implications for O2 sensing.
Science 292,464
-468.
Jaakkola, P., Mole, D. R., Tian, Y. M., Wilson, M. I., Gielbert,
J., Gaskell, S. J., Kriegsheim, Av., Hebestreit, H. F., Mukherji, M.,
Schofiels, C. J., Maxwell, P. H., Pugh, C. W. and Ratcliffe, P. J.
(2001). Targeting of HIF-alpha to the von Hippel-Lindau
ubiquitylation complex by O2-regulated prolyl hydroxylation.
Science 292,468
-472.
Jiang, B.-H., Jiang, G., Zheng, J. Z., Lu, Z., Hunter, T. and
Vogt, P. K. (2001). Phosphatidylinositol 3-kinase signaling
controls levels of hypoxia-inducible factor 1. Cell Growth
Differ. 12,363
-369.
Lander, H. M., Milbank, A. J., Tauras, J. M., Hajjar, D. P., Hempstead, B. L., Schwartz, G. D., Kraemer, R. T., Mirza, U. A., Chait, B. T., Burk, S. C. and Quilliam, L. A. (1996). Redox regulation of cell signalling. Nature 381,380 -381.[CrossRef][Medline]
Lando, D., Peet, D. J., Whelan, D. A., Gorman, J. J. and
Whitelaw, M. L. (2002). Asparagine hydroxylation of the HIF
transactivation domain a hypoxic switch. Science
295,858
-861.
Levy, A. P., Levy, N. S., Wegner, S. and Goldberg, M. A.
(1995). Transcriptional regulation of the rat vascular
endothelial growth factor gene by hypoxia. J. Biol.
Chem. 270,13333
-13340.
Levy, A. P., Levy, N. S. and Goldberg, M. A.
(1996). Hypoxia-inducible protein binding to vascular endothelial
growth factor mRNA and its modulation by the von Hippel-Lindau protein.
J. Biol. Chem. 271,25492
-25497.
Li, C. and Jackson, R. M. (2002). Reactive
species mechanisms of cellular hypoxia-reoxygenation injury. Am. J.
Physiol. Cell Physiol. 282,C227
-C241.
Matozaki, T., Nakanishi, H. and Takai, Y. (2000). Small G-protein networks: their crosstalk and signal cascades. (2000). Cell Signaling 12,515 -524.[CrossRef][Medline]
Maxwell, P. H., Dachs, G. U., Gleadle, J. M., Nicholls, L. G.,
Harris, A. L., Stratford, I. J., Hankinson, O., Pugh, C. W. and Ratcliffe, P.
J. (1997). Hypoxia-inducible factor-1 modulates gene
expression in solid tumors and influences both angiogenesis and tumor growth.
Proc. Natl. Acad. Sci. USA
94,8104
-8109.
Molitoris, B. A. (1991). Ischemia-induced loss of epithelial polarity: potential role of the actin cytoskeleton. Am. J. Physiol. 260,F769 -F778.[Medline]
Nicol, D., Hii, S. I., Walsh, M., Teh, B., Thompson, L., Kennett, C. and Gotley, D. (1997). Vascular endothelial growth factor expression is increased in renal cell carcinoma. J. Urol. 157,1482 -1486.[Medline]
Nobes, C. D. and Hall, A. (1995). Rho, rac, and cdc42 GTPases regulate the assembly of multimolecular focal complexes associated with actin stress fibers, lamellipodia, and filopodia. Cell 81,53 -62.[Medline]
Pearlstein, D. P., Ali, M. H., Mungai, P. T., Hynes, K. L.,
Gewertz, B. L. and Schumacker, P. T. (2002). Role of
mitochondrial oxidant generation in endothelial cell responses to hypoxia.
Arterioscler. Thromb. Vasc. Biol.
22,566
-576.
Raman, N. and Atkinson, S. J. (1999). Rho controls actin cytoskeletal assembly in renal epithelial cells during ATP depletion and recovery. Am. J. Physiol. 276, C1312, C1324.[Medline]
Ren, X. D., Kiosses, W. B. and Schwartz, A.
(1999). Regulation of the small GTP-binding protein Rho by cell
adhesion and the cytoskeleton. EMBO J.
18,578
-585.
Ridley, A. J. and Hall, A. (1992). The small GTP-binding protein rac regulates growth factor-induced membrane ruffling. Cell 70,389 -399.[Medline]
Robert, B., Zhao, X. and Abrahamson, D. R.
(2000). Coexpression of neuropilin-1, Flk1, and VEGF(164) in
developing and mature mouse kidney glomeruli. Am. J. Phys. Renal
Physiol. 279,F275
-F282.
Semenza, G. L. (2000a). HIF-1 and human
disease: one highly involved factor. Genes Dev.
14,1983
-1991.
Semenza, G. L. (2000b). HIF-1: mediator of
physiological and pathophysiological responses to hypoxia. J. Appl.
Physiol. 88,1474
-1480.
Semenza, G. L. and Wang, G. L. (1992). A nuclear factor induced by hypoxia via de novo protein synthesis binds to the human erythropoietin gene enhancer at a site required for transcriptional activation. Mol. Cell. Biol. 12,5447 -5454.[Abstract]
Semenza, G. L., Jiang, B. H., Leung, S. W., Passantino, R.,
Concordet, J.-P., Maire, P. and Giallongo, A. (1996). Hypoxia
response elements in the aldolase A, enolase 1, and lactate dehydrogenase A
gene promoters contain essential binding sites for hypoxia-inducible factor
1. J. Biol. Chem. 271,32529
-32537.
Stein, I., Itin, A., Einat, P., Skaliter, R., Grossman, Z. and
Keshet, E. (1998). Translation of vascular endothelial growth
factor mRNA by internal ribosome entry: implications for translation under
hypoxia. Mol. Cell. Biol.
18,3112
-3119.
Suwa, H., Ohshio, G., Imamura, T., Watanabe, G., Arii, S., Imamura, M., Narumiya, S., Hiai, H. and Fukumoto, M. (1998). Overexpression of the rhoC gene correlates with progression of ductal adenocarcinoma of the pancreas. Br. J. Cancer. 77,147 -152.[Medline]
Takai, Y., Sasaki, T. and Matozaki, T. (2001).
Small GTP-binding proteins. Physiol. Rev.
81,153
-208.
von Marschall, Z., Cramer, T., Höcker, M., Finkenzeller,
G., Wiedenmann, B. and Rosewicz, S. (2001). Dual mechanism of
vascular endothelial growth factor upregulation by hypoxia in human
hepatocellular carcinoma. Gut
48, 87-96.
Wang, G. L. and Semenza, G. L. (1995).
Purification and characterization of hypoxia-inducible factor 1. J.
Biol. Chem. 270,1230
-1237.
Xia, G., Kageyama, Y., Hayashi, T., Kawakami, S., Yoshida, M. and Kihara, K. (2001). Regulation of vascular endothelial growth factor transcription by endothelial PAS domain protein 1 (EPAS1) and possible involvement of EPAS1 in the angiogenesis of renal cell carcinoma. Cancer 91,1429 -1436.[CrossRef][Medline]