HIF-1{alpha} mRNA and protein upregulation involves Rho GTPase expression during hypoxia in renal cell carcinoma

Sandra Turcotte, Richard R. Desrosiers and Richard Béliveau*

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


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 Materials and Methods
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The small G proteins of the Rho family are involved in reorganization of the actin cytoskeleton, cell migration and in the regulation of gene transcription. Hypoxia-induced ATP depletion results in the disruption of actin organization which could affect Rho functions. In solid tumors, regions with low oxygen tension stimulate angiogenesis in order to increase oxygen and nutrient supply. This process is mediated by stabilization of the transcriptional factor hypoxia inducible factor 1 (HIF-1), which increases vascular endothelial growth factor (VEGF) production. In this study, we investigated the activities of Rho proteins, which are key regulators of cytoskeleton organization during hypoxia in renal cell carcinoma. Caki-1 cells were exposed to hypoxia (1% O2) and exhibited increased Cdc42, Rac1 and RhoA protein expression. Immunoprecipitation of metabolically labelled RhoA showed that overexpression was at least due to neo-synthesis. The Rho GTPases overexpressed during hypoxia were mainly located at membranes and pull-down assays demonstrated that they were active since they bound GTP. RT-PCR analysis indicated that the increase in RhoA protein expression was also reflected at the mRNA level. Overexpression and activation of Rho proteins were downstream of, and dependent on, the production of reactive oxygen species (ROS) since, in the presence of an inhibitor, both the rise of ROS and upregulation of Rho proteins were abolished. Importantly, preincubation of cells with the toxin C3, which inhibits RhoA, reduced HIF-1{alpha} protein accumulation by 84% during hypoxia. Together, these results support a model where ROS upregulate Rho protein expression and where active RhoA is required for HIF-1{alpha} accumulation during hypoxia.

Key words: Hypoxia, Hypoxia inductible factor 1, Rho proteins, Carcinoma cells, Reactive oxygen species


    Introduction
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 Introduction
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Decreased levels of cellular O2, a condition known as hypoxia, are seen in pathological states including ischemia, pulmonary diseases and cancer (Semenza, 2000aGo). In the absence of angiogenesis, tumors with a volume greater than 1-2 mm3 display hypoxic areas because of poor oxygen diffusion (Dachs and Tozer, 2000Go). Several cellular responses including erythropoiesis, glycolysis and angiogenesis are activated in hypoxia (Levy et al., 1995Go; Semenza and Wang, 1992Go; Firth et al., 1994Go). The enhanced transcription of some of these genes is mediated by the binding of hypoxia inducible factor 1 (HIF-1) at a specific consensus sequence (5'-RCGTG-3') found in the promoter regions of these genes (Semenza et al., 1996Go). HIF-1 is a transcription factor induced by hypoxia, which possesses two subunits, HIF-1{alpha} and HIF-1ß (Wang and Semenza, 1995Go). HIF-1ß, also called ARNT, is constitutively expressed whereas HIF-1{alpha} expression is regulated by [O2]. Recently, several studies have emphasized the importance of von Hippel-Lindau protein (pVHL) in the degradation of HIF-1{alpha} upon normoxia. Specific hydroxylation of proline residues of HIF-1{alpha} results in the binding of this subunit to pVHL, thereby enabling its polyubiquitination and its rapid degradation by the associated proteasome pathway (Semenza, 2000bGo; Ivan et al., 2001Go; Jaakkola et al., 2001Go). Under hypoxic conditions, the hydroxylases cease to function and HIF-1{alpha} escapes the degradation. In addition to its protection against pVHL-dependent proteolysis, there is an upregulation of HIF-1{alpha} mRNA expression in certain cell types and tissues (Catron et al., 2001Go). Consequently, the synthesis and the stabilization of HIF-1{alpha} promotes its accumulation in the nucleus, where it forms a functional transcription factor when bound to HIF-1ß.

Although several studies have been performed to investigate the role of HIF-1{alpha}, 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, 2000Go). Several studies demonstrated that ROS generation was necessary for the transcriptional response induced by hypoxia (Chandel and Schumacker, 2000Go; Li and Jackson, 2002Go; Duranteau et al., 1998Go; Pearlstein et al., 2002Go; Chandel et al., 1998Go). 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{alpha}, erythropoietin (EPO), glycolytic enzymes and vascular endothelial growth factor (VEGF) (Chandel et al., 1998Go).

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., 1997Go; Xia et al., 2001Go). 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., 2001Go); the transcriptional activation is a consequence of HIF-1{alpha} 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., 1992Go; Houck et al., 1991Go). 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., 2001Go).

The depletion of cellular ATP that occurs during hypoxia disrupts the actin cytoskeleton in many cell types including renal epithelial cells (Molitoris, 1991Go). 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, 1998Go; Takai et al., 2001Go). 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., 2000Go). The cyclic passage between these two forms is tightly controlled by different classes of regulatory proteins (Boivin et al., 1996Go). 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, 1999Go; Gopalakrishnan et al., 1998Go). 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, 2001Go). 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{alpha} mRNA and protein. Our findings demonstrated that Rho proteins are necessary regulators of the cascade leading to HIF-1{alpha} accumulation and permit a best understanding of carcinogenesis during hypoxia.


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 Materials and Methods
 Results
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Cell culture and experimental conditions
Caki-1 cells were obtained from American Type Culture Collection (ATCC, Rockville, MD) and cultured in McCoy's medium supplemented with 50 units/ml penicillin, 50 µg/ml streptomycin and 10% fetal calf serum (FCS) (Gibco-BRL Life Technologies, Burlington, Ontario, Canada). The cells were seeded at 1.5x106 cells/100 mm diameter dish for 3-4 days at 37°C in presence of 10 ml of medium in a humidified atmosphere of 5% CO2. All experiments were carried out with confluent cultures. Hypoxic conditions were attained by incubation of cells in an anaerobic box. The oxygen was maintained at 1% by a compact gas oxygen controller Proox model 110 (Reming Bioinstruments, Redfield, NY) with a residual gas mixture composed of 94% N2 and 5% CO2.

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{alpha} (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., 1999Go; Ren et al., 1999Go). 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{alpha}, VEGF and {alpha}-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{alpha}, 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 {alpha}-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.


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Table 1. PCR Primers used for Cdc42, RhoA, {alpha}-tubulin, HIF-1{alpha} and VEGF

 

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
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 Materials and Methods
 Results
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 References
 
ATP depletion and cell survival during hypoxia
In order to characterize the early cellular events in our RCC model, ATP depletion was measured in Caki-1 cells during 2 hours of hypoxic treatment at 1% O2 (Fig. 1A). ATP levels quickly dropped and reached 50% of control values after only 13 minutes of hypoxia. More than 90% of ATP was depleted following treatments of at least 30 minutes. In spite of this marked energy drop, 6 hours of hypoxia was not cytotoxic for Caki-1 cells (Fig. 1B). ATP depletion has been well documented as causing a disorganization of the actin cytoskeleton (Dagher, 2000Go). Since regulation of the actin cytoskeleton is mediated by Rho proteins, we next investigated the effects of hypoxia on expression and activation of these proteins.



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Fig. 1. ATP depletion and cell survival during hypoxia in renal cell carcinoma. Caki-1 cells were exposed to 1% O2 and ATP content was measured for up to 120 minutes using a luciferase assay (A). The cytotoxicity of hypoxia was measured by cleavage of WST-1 in formazan by spectrophotometry (B). ATP levels and cytotoxicity during hypoxia are represented as the percentage values relative to normoxia. Means ± s.e.m. are given for three separate experiments.

 

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, 2001Go). 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.



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Fig. 2. Upregulation of Rho GTPase protein expression by hypoxia in renal cell carcinoma. Caki-1 cells were exposed to 1% O2 from 0 to 6 hours. Post-nuclear supernatants (20 µg of protein) were separated by SDS-PAGE and submitted to immunodetection using Cdc42, Rac1, RhoA, RhoGDI and RhoB antibodies (A). The expression of these Rho GTPases was quantified by densitometric analysis and expressed as means ± s.e.m. for three separate experiments relative to normoxic values (B). Significant differences (P<0.05) from normoxia are indicated by asterisks (*).

 

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.



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Fig. 3. Subcellular distribution of RhoA and Cdc42 proteins during hypoxia. After hypoxic treatment, renal cell carcinoma were fractionated into soluble and crude membrane fractions by centrifugation at 100,000 g for 1 hour. Each fraction (20 µg of protein) was separated by SDS-PAGE then immunodetected using Cdc42 (A) and RhoA antibodies (B). Densitometric analysis of both GTPases was carried out for soluble ({square}) and membrane ({blacksquare}) fractions. Means ± s.e.m. are shown from three independent experiments relative to samples from normoxic cells. Significant differences (P<0.05) from normoxia are indicated by asterisks (*).

 

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.



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Fig. 4. Metabolic labelling of RhoA during hypoxia is localized to membranes. Caki-1 cells were pre-incubated in cystein and methionine free medium for 1 hour, then incubated with 50 µCi/ml of [35S] Met/Cys. Subsequently, cells under normoxia or hypoxia at 1% O2 for 4 hours were fractionated in soluble fractions (S) and crude membranes (M). RhoA was immunoprecipitated, analyzed by SDS-PAGE, and labelled RhoA detected by autoradiography (A). As a control, Caki-1 cells were exposed to normoxia or hypoxia for 4 hours, fractionated in soluble and membrane fractions, then analyzed by western blot and immunodetected with RhoA antibody (A). Densitometric analysis of immunoprecipitated RhoA from cells under normoxia ({square}) and hypoxia ({blacksquare}) conditions is represented (B). Means ± s.e.m. are shown for two different experiments relative to normoxia. Significant differences (P<0.05) from normoxia are indicated by asterisks (*).

 

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.



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Fig. 5. Cdc42 and RhoA activation in Caki-1 cells exposed to 1% O2. Activation states of Cdc42 and RhoA proteins were measured by affinity precipitation with GST-PBD and GST-RBD, respectively. Cells were exposed to 1% O2 from 0 to 6 hours, lysed and incubated with 20 µg fusion proteins. After pull-down precipitation, the GTP-bound forms of Cdc42 and RhoA proteins were analyzed by SDS-PAGE and western blot. Parallel experiments were performed to assess total amounts of Rho GTPases under these conditions. Cdc42 (A) and RhoA (B) were immunodetected with appropriate antibodies. Means ± s.e.m. from densitometric analysis of active GTPases for three independent experiments are shown. Data are expressed as the percentage of relative activity compared with values found in normoxic cells. Significant differences (P<0.05) from normoxia are indicated by asterisks (*).

 

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 {alpha}-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.



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Fig. 6. Effect of hypoxia on RhoA and Cdc42 mRNA levels in renal cell carcinoma. Total RNA was isolated from Caki-1 cells in normoxia and hypoxia. RT-PCR amplification was performed using primers for RhoA for 15 to 30 cycles in order to define optimal conditions (A). At 25 cycles, RT-PCR analysis for RhoA (183 bp), Cdc42 (400 bp), and {alpha}-tubulin (321 bp) were carried out on total RNA isolated from cells incubated under hypoxia for up to three hours (B).

 

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., 1998Go). 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, 2000Go; Duranteau et al., 1998Go; Grishko et al., 2001Go).



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Fig. 7. Hypoxia increases ROS production in renal cell carcinoma. Caki-1 cells were incubated with 10 µM DCFH in hypoxic conditions for 0 to 240 minutes. Production of ROS was measured by fluorescence emitted from oxidized DCF. After hypoxia, cells were lyzed and fluorescence was measured at 530 nm (A). To inhibit ROS production, cells were preincubated with 10 uM DPI then exposed to hypoxic or normoxic conditions for 1 hour (B). Fluorescence data are represented relative to normoxic values. Means ± s.e.m. were calculated from three independent experiments. Significant differences (P<0.1) are indicated by asterisks (*).

 

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).



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Fig. 8. Effect of DPI on overexpression of Rho GTPases in hypoxic conditions. Caki-1 cells were pre-treated with 10 µM DPI in normoxia. Cells were then incubated in hypoxic conditions from 0 to 4 hours. Cell lysates (20 µg of protein) were separated by SDS-PAGE, followed by immunodetection. Densitometric analysis of Cdc42 (A) and RhoA (B) expression are presented for cells in hypoxia ({blacksquare}) and cells in hypoxia plus DPI ({square}). Means ± s.e.m. for three independent experiments are indicated. Data are expressed relative to normoxic values. Significant differences (P<0.05) from normoxia are indicated by asterisks (*).

 

HIF-1{alpha} and VEGF mRNA levels are increased in renal cell carcinoma exposed to hypoxia
Several studies have demonstrated stabilization of HIF-1{alpha} and upregulation of VEGF in hypoxic conditions. To examine the role of Rho protein induction during hypoxia, we studied their involvement in HIF-1{alpha} and VEGF expression. Amplification conditions for the primer sets were determined by RT-PCR analysis. Fig. 9A shows that an amplification of HIF-1{alpha} 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{alpha} 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., 2000Go). As negative control, {alpha}-tubulin was amplified at same time (Fig. 9B). Immunoprecipitation followed by immunodetection with HIF-1{alpha} 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{alpha} mRNA and protein induction.



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Fig. 9. HIF-1{alpha} and VEGF mRNA expression during hypoxia. Amplification of HIF-1{alpha} (487 bp) and VEGF for 15 to 30 cycles was carried out by RT-PCR to determine the linear portion of the amplification reaction (A). Two isoforms of VEGF mRNA were amplified, VEGF165 (627 bp) and VEGF189 (699 bp). Caki-1 cells were exposed to 1% O2 and total RNA was isolated. RT-PCR analysis of HIF-1{alpha} at 25 cycles and VEGF at 30 cycles using total RNA from Caki-1 cells under kinetic of hypoxia is shown (B). mRNA {alpha}-tubulin (321 bp) was used as a control. HIF1-a protein expression was analyzed by its immunoprecipitation from nuclei and immunodectection (C). Two independent experiments were performed.

 

Overexpressed RhoA stimulates HIF-1{alpha} and VEGF mRNA levels in normoxia
We determined whether Rho GTPase overexpression could enhance HIF-1{alpha} 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{alpha} (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 {alpha}-tubulin was examined (Fig. 10C). These results suggest that the level of activated RhoA contributes to stimulate HIF-1{alpha} and VEGF mRNA levels.



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Fig. 10. RhoA activates HIF-1{alpha} and VEGF mRNA levels in normoxia. Caki-1 cells were transfected or not with vector alone (pcDNA3) or with dominant-active RhoA (pcDNA3-RhoAV14-Myc) for 24 hours using lipofectamine. Total RNA was isolated then HIF-1{alpha} (A), VEGF (B) and {alpha}-tubulin (C) mRNA expressions were evaluated by RT-PCR. Two isoforms of VEGF were amplified, VEGF165 and VEGF189. Densitometric analysis was used to assess relative mRNA expression levels. Means ± s.e.m. from two experiments and values are plotted relative to values from normoxic cells from the sum of both isoforms is presented on the histograms. Significant differences (P<0.05) from normoxia are indicated by asterisks (*).

 

Toxin C3 blocks HIF-1{alpha} 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).



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Fig. 11. Exotoxin C3 blocks HIF-1{alpha} mRNA and protein overexpression and VEGF mRNA stimulation. Cells were pre-treated with 50 µg/ml of C3 toxin for 24 hours in normoxia. After this, cells were placed in hypoxia (H) or kept in normoxia (N) for 4 hours. Cells were fractionated into soluble and membrane fractions and proteins were separated by SDS-PAGE. Efficiency of ADP-ribosylation by C3 toxin was verified by immunodetection of RhoA, as the ADP-ribosylation reduces protein mobility (A). Alternatively, following incubation with the toxin, cells were exposed to hypoxia or in normoxia for 4 hours and total RNA isolated. The effects of the toxin were evaluated on mRNA levels (B) of HIF-1{alpha}, VEGF and {alpha}-tubulin as the negative control, and on protein expression (C) of HIF-1{alpha} and {alpha}-tubulin by RT-PCR or western blot analysis. These experiments were carried out at least twice. Significant differences (P<0.05) from normoxia are indicated by asterisks (*).

 

We next studied whether RhoA inhibition by C3 toxin might affect HIF-1{alpha} and VEGF mRNA levels. Incubation with C3 had no effect on HIF-1{alpha} and VEGF mRNA levels under normoxia conditions (Fig. 11B). More notably, C3 toxin significantly abolished the induction of HIF-1{alpha} and VEGF mRNA observed during hypoxia (Fig. 11B). Again, the level of {alpha}-tubulin mRNA was analyzed as negative control and found unaffected by hypoxia (Fig. 11B). To further validate the RhoA contribution on HIF-1{alpha} 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{alpha} expression (twofold), which indicates an accumulation of the protein either due to an increase synthesis or resulting from of HIF-1{alpha} stabilization. A pre-treatment with toxin C3 has no effect on HIF-1{alpha} protein expression in normoxic conditions. As expected, toxin C3 prevented the accumulation of HIF-1{alpha} protein by 84% under hypoxia, which suggests that new synthesis was necessary to increase HIF-1{alpha} level (Fig. 11C). Hypoxia and treatment with this toxin unaffected {alpha}-tubulin protein expression. These results clearly demonstrate that RhoA upregulation and activation occurring during hypoxia are upstream and contribute to HIF-1{alpha} mRNA and protein accumulation in RCC.


    Discussion
 Top
 Summary
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Hypoxic conditions regulate several metabolic enzymes and transcription factors that are involved in cancer, ischemia and pulmonary diseases (Semenza, 2000aGo). One cellular event that follows low oxygen concentrations is ATP depletion, which is reflected in disruptions of cell polarity and intercellular junctions through changes in actin cytoskeleton dynamics (Dagher, 2000Go). Since this organization of actin structures is mediated by Rho proteins such as Cdc42, Rac1 and RhoA (Raman and Atkinson, 1999Go; Bokoch, 2000Go), we decided to investigate the role of these key regulators of cytoskeletal rearrangement during hypoxic conditions in a renal carcinoma model using Caki-1 cells.

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, 2001Go). However, this study did not observe a role of RhoA for HIF-1{alpha} 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, 1995Go; Ridley and Hall, 1992Go). 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{alpha} 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{alpha} (Chandel and Schumacker, 2000Go; Li and Jackson, 2002Go; Duranteau et al., 1998Go; Pearlstein et al., 2002Go; Chandel et al., 1998Go). 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, 2000Go; Duranteau et al., 1998Go; Grishko et al., 2001Go). 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, 2000Go). 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., 1996Go). 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, 2001Go; Jiang et al., 2001Go). Other studies have also demonstrated that PI-3K may activate Rho proteins such as RhoA and Rac1 in human fibrosarcoma cells (Gupta et al., 2001Go). 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, 1998Go). 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{alpha} (Maxwell et al., 1997Go). 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., 1996Go). 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., 1998Go). To better understand the function of early Rho expression and activation in hypoxic conditions, we thus evaluated the kinetics of HIF-1{alpha} 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{alpha} and VEGF expression, we firstly transfected cells with a dominant-active RhoA cDNA. In RhoAV14-transfected RCC, HIF-1{alpha} and VEGF mRNA levels are induced but less than by hypoxia (Fig. 10). Interestingly, these data suggest that mRNA expression of HIF-1{alpha} 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{alpha} and VEGF during hypoxia. Secondly, supporting this conclusion for a key role of RhoA in hypoxia-induced HIF-1{alpha} 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{alpha} protein during hypoxia by 84% (Fig. 11). From the results showing that dominant-active RhoA induces HIF-1{alpha} and VEGF mRNA in normoxia and that toxin C3 blocks the induction of HIF-1{alpha} mRNA and protein and VEGF mRNA stimulation by hypoxia, we conclude that RhoA is necessary for HIF-1{alpha} upregulation at low O2 tension in RCC.

This new pathway involving RhoA in the upregulation of HIF-1{alpha} is likely a complementary process to the stabilization of the protein during hypoxia. Under normoxic conditions, HIF-1{alpha} is recognized by the tumour-suppressor protein pVHL. This interaction promotes a rapid degradation of HIF-1{alpha} by the proteasome (Ivan et al., 2001Go). pVHL binds the oxygen degradation domain of HIF-1{alpha} 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., 2002Go). When the oxygen concentration diminishes, the hydroxylation of proline and asparagine residues is stopped and pVHL-HIF-1{alpha} interaction is lost allowing an accumulation of HIF-1{alpha}. As expected, the basal level of HIF-1{alpha} in RCC is barely detectable in normoxia (Fig. 9). Thus, if the efficiency of HIF-1{alpha} mRNA translation is unaffected by hypoxia and that the protein is protected against proteolysis, the accumulation of HIF-1{alpha} could be a gradual process. However, the upregulation of HIF-1{alpha} 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{alpha} protein expression in hepatocarcinomas cells (Hep3B) (Hirota and Semenza, 2001Go), 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{alpha} and VEGF production in renal cell carcinoma since they are inhibited by C3 toxin.



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Fig. 12. Model for Rho-dependent HIF-1{alpha} and angiogenesis activation during hypoxia. This schema summarizes the main steps that we have identified to play crucial roles in the induction of HIF-1{alpha} mediated by Rho GTPases. The involvement of ROS production was abolished by DPI and prevented Cdc42 and RhoA induction. Also, the C3 toxin that abolished RhoA expression prevented HIF-1{alpha} induction under hypoxic conditions. The kinetics demonstrated 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 GTPase cascades (dotted arrows).

 


    Acknowledgments
 
We thank Allan Hall, Gary M. Bokoch and Martin Schwartz for their gifts of expression plasmids. This work was supported by grants from the Canadian Institutes of Health Research to R.B. and from the Natural Sciences and Engineering Research Council of Canada (NSERC) to R.R.D.


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 Discussion
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