Oxygen is of vital importance for the metabolism and function of all cells in the human body. Hypoxia, the reduction of oxygen supply, results in adaptationally appropriate alterations in gene expression through the activation of hypoxia-inducible factor 1 (HIF-1) to overcome any shortage of oxygen. Thyroid hormones are required for normal function of nearly all tissues, with major effects on oxygen consumption and metabolic rate. Thyroid hormones have been found to augment the oxygen capacity of the blood by increasing the production of erythropoietin (EPO) and to improve perfusion by vasodilation through the augmented expression of adrenomedullin (ADM). Because the hypoxic expression of both genes depends on HIF-1, we studied the influence of thyroid hormone on HIF-1 activation in the human hepatoma cell line HepG2 under normoxic and hypoxic conditions. We found that thyroid hormones increased HIF-1α protein accumulation by increasing HIF-1α protein synthesis rather than attenuating its proteasomal degradation. HIF-1α expression directly correlated with augmented HIF-1 DNA binding and transcriptional activity of luciferase reporter plasmids, whereas HIF-1β levels remained unaffected. Knocking down HIF-1α by short interfering RNA (siRNA) clearly demonstrated that thyroid hormone-induced target gene expression required the presence of HIF-1. Although an increased association of the two known coactivators of HIF-1, p300 and SRC-1, was found, thyroid hormone did not affect the activity of the isolated COOH-terminal transactivating domain of HIF-1α. Increased synthesis of HIF-1α may contribute to the adaptive response of increased oxygen demand under hyperthyroid conditions.
- hypoxic gene expression
- oxygen sensing
erythropoietin (EPO), a 30.4-kDa glycoprotein hormone, is the major physiological stimulator of red blood cell formation in mammals (18). The main EPO production sites are the kidney in adults and the liver in fetuses (6). EPO production is induced by hypoxia, a state when oxygen supply does not cover the demand of the tissue. EPO mRNA levels are induced 50- to 100-fold in vitro by physiologically relevant levels of hypoxia (9). In vivo under severe hypoxia, production of EPO can be increased up to 1,000-fold (18, 29).
Oxygen-dependent EPO expression is regulated by hypoxia-inducible factor 1 (HIF-1), a heterodimer of the O2-labile 120-kDa α-subunit, and the constitutive 91- to 94-kDa β-subunit (37). The cellular levels of HIF-1α are adjusted by the ubiquitin-proteasome-dependent degradation of HIF-1α under normoxic conditions, allowing to tightly couple the protein appearance to the ambient oxygen tension (31). At high Po2, HIF-1α is posttranslationally hydroxylated at proline residues 402 and 564 by O2-sensitive prolyl hydroxylases, termed PHD1, PHD2, and PHD3, which are recognized today as the most likely cellular O2 sensors (7, 17). Hydroxylated HIF-1α is recognized by the tumor suppressor von Hippel-Lindau (pVHL) protein for ubiquitination by the E3 ubiquitin-protein ligase (23). Under hypoxia, HIF-1α evades proteasomal degradation because of the lack of proline hydroxylation and concomitantly accumulates and is translocated into the nucleus to form the HIF-1 complex by dimerization with constitutively expressed HIF-1β (identical to aryl hydrocarbon receptor nuclear translocator 1, ARNT1) (14, 37). Binding of HIF-1 to DNA at HIF-binding site (HBS) within the hypoxia response element (HRE) in the 3′-flanking EPO enhancer increases expression of the gene under hypoxic conditions (33). In addition to hypoxic accumulation, the trans-activity of HIF-1α is regulated by the O2-sensitive asparagyl-hydroxylase FIH-1 (factor inhibiting HIF-1) (22). Under hypoxia the lack of hydroxylation of Asp803 allows binding of the adapter protein p300 to recruit further coactivator proteins such as steroid receptor coactivator-1 (SRC-1) (3, 20).
Like EPO, the expression of adrenomedullin (ADM), a hypotensive peptide originally isolated from a pheochromocytoma, is hypoxia inducible (4). Promoter analysis revealed that this induction was primarily mediated by HIF-1 bound to regulatory DNA sequences within the promoter of rodent (4, 25) and human cells (12, 16).
Thyroid hormone is required in nearly all tissues, with major effects on oxygen consumption and metabolic rate. Adaptation to this increased metabolic demand is partly achieved by potent effects of thyroid hormone on erythropoiesis and thus blood oxygen capacity. Thyroid hormones directly increase the proliferation of erythroid progenitors (5, 13) and thyroid hormone receptors were identified on nucleated erythroid cells isolated from hypoxic hamsters (1). Apart from the direct effect in erythroid precursors, thyroid hormone directly enhanced hypoxia-inducible EPO formation both in the isolated perfused rat kidney and HepG2 cells (10). In addition, tissue perfusion may be increased through the stimulated expression of the potent vasodilator ADM by thyroid hormones (15, 25).
The molecular mechanisms of thyroid hormone-inducible EPO and ADM expression have not yet been elucidated. Thyroid hormone binds to the intracellular thyroid hormone receptor (TR), a member of the nuclear hormone receptor family, which acts as a transcription factor and regulates gene expression (8). Moreover, it has been reported that steroid receptor coactivator-1 (SRC-1) functions as a positive regulator of the TR-mediated transactivation pathway (19). Thus herein we studied whether thyroid hormones impinge on the HIF-1 activation pathway and by that means affect hypoxia-inducible expression of EPO and ADM.
MATERIALS AND METHODS
Cell culture and in vitro stimulation.
The human hepatoma cell line, HepG2, obtained from the American Type Culture Collection (ATCC HB 8065) was grown in RPMI 1640 supplemented with 10% fetal calf serum (FCS), penicillin (100 U/ml), and streptomycin (100 μg/ml) in a humidified atmosphere (5% CO2 in air) at 37°C. Care was taken to avoid formation of cell clumps that would result in a heterogeneous distribution of the pericellular oxygen tension. To examine the effects of thyroid hormone, cells were regularly cultured serum free in the RPMI 1640 medium containing 1% serum supplement (SS; transferrin, insulin, selenium, albumin; Sigma, Munich, Germany) for 24 h before the experiment to remove the effects of serum-derived hormones. Transiently transfected HepG2 cells were kept in RPMI 1640 supplemented with 1% SS for 8 h before the experiments.
At the beginning of an experiment, HepG2 cells received fresh medium (RPMI 1640 supplemented with 1% SS) containing the respective thyroid hormone concentration. Triiodothyronine (T3) and thyroxine (T4; Sigma, Munich, Germany) stock solutions (14.86 mM in 1:4 1 N HCl-ethanol for T3 and 56.2 mM in 4 N ammonium hydroxide in methanol for T4) were stored at −80°C. To achieve hypoxic conditions, culture dishes were placed in a Heraeus incubator (Hanau, Germany) with 5% CO2, and nitrogen (N2) to balance for different O2 concentrations. Hypoxia was defined as 3% O2 if not indicated otherwise. Control normoxic cells were placed in an incubator (5% CO2 in air) for equivalent time periods. For reoxygenation experiments, cells were exposed to hypoxia for 4 h and then transferred to 21% O2 for different times. Nuclear extracts were prepared using the method of Schreiber et al. (32) and subjected to Western blot analysis and electrophoretic mobility shift assays (EMSAs).
Quantitative real-time RT-PCR analysis.
Total RNA was extracted by guanidinium isothiocyanate extraction as described (34). Total RNA (1 μg) was reverse transcribed with oligo(dT) and Moloney murine leukemia virus reverse transcriptase (Promega). Gene expression of human EPO was quantitated using the qPCR Mastermix for SYBR Green I (Eurogentec, Belgium) and the GeneAmp 5700 sequence Detection System (PE Biosystems). The PCR reactions were set up in a final volume of 25 μl with 0.5 μl cDNA, 1× reaction buffer containing SYBR Green I, 10 pmol forward (F), and 10 pmol reverse primer (R). Primer sets used for EPO: (F) 5′-CTCCGAACAATCACTGCT-3′ and (R) 5′-GGTCATCTGTCCCCTGTCCT-3′; ADM: (F) 5′-GGATGCCGCCCGCATCCGAG-3′ and (R) 5′-GACACCAGAGTCCGACCCGG-3′; β-actin: (F) 5′-TCACCCACACTGTGCCCATCTA CGA-3′ and (R) 5′-CAGCGGAACCGCTCATTGCCAATGG-3′; HIF-1α: (F) 5′-GCTGGC CCCAGCCGCTGGAG-3′ and (R) 5′-GAGTGCAGGGTCAGCACTAC-3′. Oligonucleotides were purchased from Invitrogen.
Agarose gel electrophoresis confirmed the specificity of the amplification product. The resulting PCR fragments were visualized on ethidium bromide-stained 1.5% agarose gels. Tenfold dilutions of purified PCR products starting at 1 pg to 0.1 fg were used as standards. Amplification conditions were set to 10 min at 95°C followed by 45 PCR cycles (15 s at 95°C, 1 min at 60°C). The quantity of cDNA used in each reaction was normalized to the β-actin cDNA and expressed as cDNA per microgram total RNA.
ELISA for EPO.
EPO protein in the culture supernatant was measured by ELISA (Quantikine IVD EPO; R&D System, Wiesbaden-Nordenstadt, Germany).
Protein extract preparation and Western immunoblotting.
Nuclear extracts were prepared using the methods of Schreiber et al. (32). All procedures were performed at 4°C. Briefly, HepG2 cells (70–80% density of the cells in dishes with 10-cm diameter) were washed with cold PBS, drained, and scraped from plates with 150 μl cold nuclear extract buffer A (10 mM HEPES pH 7.9, 1.5 mM MgCl2, 10 mM KCl, 0.5 mM PMSF, 0.5 mM DTT, 0.4% NP-40, 1× protease-inhibitor-cocktail, Roche), transferred into Eppendorf tubes, and incubated on ice for 20 min. The cell lysate was centrifuged at 5,000 g for 5 min at 4°C and the supernatant was discarded. The pellet was resolved in 80 μl cold nuclear extract buffer B (20 mM HEPES pH 7.9, 420 mM NaCl, 1.5 mM MgCl2, 0.2 mM EDTA, 0.5 mM PMSF, 0.5 mM DTT, 1× protease-inhibitor-cocktail) and homogenized with a magnetic stirrer on ice for 30 min. Cellular debris was removed by centrifugation at 13,000 g for 15 min at 4°C. The supernatant was used as a nuclear extract, and 5 μl of supernatant was taken out for protein determination. The supernatant was stored at −80°C. For whole cell lysates, cells were lysed on the plate with 100 μl extract buffer (0.1% NP-40, 300 mM NaCl, 10 nM Tris pH 7.9, 1 mM EDTA, 1:7 dilution protein-inhibitor-cocktail) for 20 min on ice. Extracts were spun down in a microfuge, 3,600 g for 5 min at 4°C, quantitated using the Bio-Rad protein assay reagent and stored at −80°C. After addition of volume of 4× sample buffer (50 mM Tris pH 6.8, 2% SDS, 5% β-mercaptoethanol, 0.0125% bromphenol blue, 1% glycerin), 75 μg of total cell lysate or 20 μg nuclear extract per lane were subjected to 7.5% SDS-PAGE, separated, and transferred to a nitrocellulose membrane (0.2 μm pore size; Schleicher and Schuell). Blots were stained with Ponceau S solution to ensure equal protein loading and transfer. The membranes were blocked with 5% nonfat dry milk powder in TBS-T, incubated with anti-human HIF-1α mouse monoclonal (Transduction Laboratories, Heidelberg, Germany; 1:250 dilution in TBS-T containing 5% nonfat milk), anti-HIF-2α (Novus Biologicals), or anti-nuclear factor (NF)-κB (Cell Signalling, Hamburg, Germany). Horseradish peroxidase-conjugated anti-mouse IgG or anti-rabbit antibodies were used as a secondary antibody at a 1:10,000 dilution in TBS-T containing 5% nonfat milk. Anti-α-tubulin antibody (at a 1:500 dilution, Santa Cruz) was used as a loading control. Immunoreactive proteins were visualized using the enhanced chemiluminescence plus (ECL) detection system followed by exposure to X-ray film (Agfa, Mortsel, Belgium).
Electrophoretic mobility shift assay.
Double-stranded oligonucleotides containing the HIF binding site from the HRE (5′-GCC CTA CGT GCT GTC TCA-3′) of the EPO enhancer were used as probes. The double-strand fragments were end labeled by filling in 5′ overhangs with 32P-labeled adenosine triphosphate ([32P]dATP) using T4 polynucleotide kinase. DNA-protein binding reactions were carried out in a total volume of 20 μl containing 5 μg nuclear extract, 30 fmol 32P-labeled oligonucleotides, and a nonspecific competitor (50 ng calf thymus DNA) in a binding buffer with a final concentration of 12 mM HEPES, 4 mM Tris (pH 7.9), 60 mM potassium chloride, 1 mM EDTA, and 1 mM dithiothreitol. After incubation for 30 min at room temperature, 1 μg anti-HIF-1α antibody was added for supershift detection of the HIF-1 complex and incubated overnight at 4°C. The products were analyzed by electrophoresis in 5% non-denaturing polyacrylamide gels. Electrophoresis was performed at 80 V in 0.25× TBE buffer at 4°C for 4 h. The dried gels were exposed to X-ray films and PhosphoImage sheets overnight.
One milligram of cell extract was incubated 2 h at 4°C with 2.5 μg of anti-Src-1 antibody (sc-6098, Santa Cruz) or anti-p300 (sc-584, Santa Cruz), followed by overnight incubation with 20 μl of protein A-Sepharose beads (Santa Cruz) on a rotator at 4°C. Beads were washed five times with 1× TBS denatured samples and 4× sample buffer at 95°C for 5 min. Beads were removed by centrifugation, and supernatants were loaded on 7.5% SDS-PAGE followed by Western blotting and detection with the anti-HIF-1α antibody as described above.
Transient transfection assays.
For reporter gene assays, HepG2 cells were transfected by electroporation. The luciferase reporter gene plasmid pH3SVL containing an SV40 promoter-luciferase unit downstream of six HIF binding sites from the transferrin enhancer was a kind gift of R. Wenger, Zurich (30). To measure the activation of HIF-1α by its COOH terminus, the Gal4 chimeric activator/reporter system was used in which the HIF-1α 775–826 COOH-terminal transactivating domain (C-TAD) was fused to Gal4 DNA binding domain. The CMV promoter ensures expression of the fusion protein. The reporter plasmid contains the luciferase gene under the control of Gal4 binding sites. Both plasmids of the chimeric activator/reporter system were generously provided by Peter Ratcliffe, Oxford, UK (28). HepG2 (1 × 107) cells were electroporated with 10 μg plasmid at 975 μF, 250 V in 0.4-mm-thick cuvettes in 400 μl of RPMI 1640 serum-free medium using a Gene Pulser and Capacitance Extender apparatus (Bio-rad). After overnight incubation, cells were preincubated in 1% SS medium for 8 h, and then the medium was replaced by a thin layer of fresh 1% SS medium containing 50 nM T3 or carrier alone. The plates were incubated for another 18–24 h in 21% or 3% O2. Cells were lysed with 100 μl 1× reporter lysis buffer. Luciferase activity was measured with the luciferase assay system (Promega, Heidelberg, Germany). Luciferase activity was expressed in relative light units (RLU) and normalized to total cellular protein measured by using the Bio-Rad protein assay kit.
Cells, starved for 1 h in serum- and methionine-free medium (PromoCell, Heidelberg, Germany), were replaced with methionine-free medium containing 1% SS and 100 μCi/ml [35S]methionine (ICN Biomedicals) for 6 h in the absence or presence of 50 nM T3. Cells were then washed with PBS, resuspended in lysis buffer (50 mM Tris, 150 mM NaCl, 5 mM EDTA, 0.5% NP-40, 5% glycerol, 1 mM PMSF, protease inhibitor cocktail, pH 7.5) followed by immediate vortexing (3 × 15 s). After centrifugation (15,000 g for 30 min) supernatants were transferred into new tubes. The supernatant containing 1 mg total protein was supplied with 1 μg anti-HIF-1α antibody and incubated at 4°C for 1 h. Thereafter, 50 μl protein G microbeads (Miltenyi Biotec, Bergisch Gladbach, Germany) was added and incubations continued at 4°C overnight. Beads were magnetically collected following the manufacturer's manual and washed three times with 100 μl lysis buffer. Coprecipitated proteins were finally eluted by adding 95°C preheated SDS-PAGE sample buffer according to the manufacturer's manual. Protein samples were separated on 7.5% SDS-PAGE. The gel was dried for 2 h and exposed to X-ray films for 32 h.
Statistical significance was calculated using the GraphPad Instat software applying the one-way ANOVA followed by the Bonferroni multiple comparison post test.
Thyroid hormones enhance expression of the HIF-1 target genes EPO and ADM.
Expression of EPO and ADM is under control of the HIF-1 complex and was thus stimulated by hypoxia (3% O2) in HepG2 cells. Quantitation of mRNA levels by quantitative real-time PCR revealed that thyroid hormones significantly stimulated normoxic expression and augmented hypoxic induction of EPO and ADM expression after 3 h of treatment (Fig. 1).
HepG2 cells, maintained in six-well culture plates for 24 h, produced 15.8 ± 0.4 mIU EPO/ml supernatant in the absence of thyroid hormone under normoxia. T3 provoked formation of 17.6 ± 0.6 mIU EPO/ml (P < 0.05 vs. untreated controls; n = 4), whereas T4 led to the production of 18.7 ± 0.7 mIU EPO/ml (P < 0.05 vs. untreated controls; n = 4). Under hypoxia, HepG2 cells secreted 26.1 ± 0.7 mIU EPO/ml EPO protein in 24 h, which was further increased to 39.1 ± 4.1 mIU EPO/ml by T3 or 30.6 ± 1.4 mIU EPO/ml by T4 (P < 0.05 for both thyroid treated groups vs. untreated controls; each group n = 4). Thus HepG2 cells showed an oxygen-dependent EPO and ADM gene expression. Thyroid hormones significantly stimulate expression of both genes under normoxia and hypoxia.
HIF-1α protein accumulation in HepG2 cells is stimulated by thyroid hormone.
To investigate whether stimulation of HIF-1-dependent target gene expression by thyroid hormones requires HIF-1 signaling, we determined HIF-1α protein levels by Western analysis performed with whole cell lysate. HepG2 cells were treated with T3 or T4 under normoxia or hypoxia for 6 h. T3, and to a lesser extent T4, induced HIF-1α protein levels in HepG2 cells incubated under hypoxic (3% O2) conditions for 6 h (Fig. 2A). Effective concentrations of T3 on HIF-1α protein expression ranged from 2 to 500 nM (data not shown). With the use of very long exposure times (L) it was revealed that T3 also enhanced HIF-1α protein levels under normoxic conditions (Fig. 2A). In contrast, HIF-1β was constitutively present and expression remained unchanged by thyroid hormone treatment (Fig. 2A). In addition, 50 nM T3 had no effect on NF-κB accumulation, whereas HIF-1α levels were increased (Fig. 2B). Interestingly, HepG2 cells displayed high levels of HIF-2α under normoxia, which were only marginally increased by hypoxia. In the same cells, HIF-1α showed a strong hypoxic response. Nevertheless, 50 nM T3 increased HIF-1α accumulation but did not affect HIF-2α levels (Fig. 2B).
To test whether thyroid hormones affect transcription of the hif-1α gene, the effect of thyroid hormones on HIF-1α mRNA levels was determined by RT-PCR. As shown in Fig. 2C, HIF-1α mRNA levels remained constant, suggesting that thyroid hormone most likely induced HIF-1α protein expression via posttranscriptional regulation.
Thyroid hormones do not stabilize HIF-1α but increase its synthesis.
To elucidate posttranscriptional mechanisms, we examined whether T3 would stabilize HIF-1α. Therefore, HepG2 cells were exposed to hypoxia for 4 h and then returned to normoxia. Although T3 increased HIF-1α protein levels under hypoxia it did not affect HIF-1α degradation on reoxygenation (Fig. 3A). On the transition from hypoxia to reoxygenation, HIF-1α completely disappeared within 30 min in control and T3-treated cells. When the destruction of HIF-1α was inhibited by the addition of MG132, an inhibitor of proteasomal degradation, T3 was still able to increase HIF-1α protein levels (Fig. 3B). The addition of the translational inhibitor cycloheximide (CHX), however, prevented the T3-dependent increase of HIF-1α (Fig. 3C). It is interesting to note that under conditions of 3% O2 in the incubation gas and a corresponding pericellular Po2 of 5–7 mmHg, HIF-1α was cleared from the cells when new synthesis is prevented by CHX. Again, as shown above, T3 did not increase the half-life of HIF-1α, but CHX prevented the T3-induced increase in HIF-1α levels. Because these data indicated that T3 might elevate HIF-1α levels by increasing the rate of translation, 35S-radioisotopic labeling of nascent HIF-1α protein was performed. In these experiments, T3 clearly increased HIF-1α protein synthesis (Fig. 3D).
Thyroid hormone increases the formation of an HBS bound complex containing HIF-1α.
To test whether T3 also increased HIF-1 DNA binding, nuclear extracts from HepG2 cells treated with or without T3 in 3% O2 for 6 h were isolated for EMSA using a double-stranded oligonucleotide containing the HBS from the EPO 3′-enhancer as a probe (Fig. 4).
Induction of HIF-1 DNA-binding activity was detected in the nuclear extracts from hypoxic controls (lane 3). T3 increased HIF-1 DNA-binding activity under hypoxic (lane 4) conditions. To confirm the identity of the HIF-1 complex, supershift experiments were performed with the monoclonal anti-HIF-1α antibody, which shifted the complete DNA/protein complex to lower mobility. This indicates that in HepG2 cells, T3 induced HIF-1α is part of the HIF-1 complex.
Thyroid hormone stimulates the transcriptional activity of HIF-1.
To investigate whether the T3-induced HIF-1 complex resulted in an increased transcriptional activity of HIF-1, luciferase reporter gene assays were performed with the plasmid pH3SVL containing an SV40 promoter-luciferase unit downstream of six HIF binding sites from the transferrin enhancer. Twenty-four hours after transient transfection, cells were incubated under normoxic and hypoxic conditions in the presence or absence of 50 nM T3 for another 18 h. T3 significantly increased the HIF-1-dependent activation of the reporter gene (Fig. 5A).
In contrast, the biological inactive analog, reverse T3, was without any effect (data not shown). To further prove the requirement of HIF-1α for the T3 effect, HIF-1α was knocked down by siRNA for HIF-1α. HepG2 cells were efficiently cleared from HIF-1α protein (Fig. 5B) and HIF-1α mRNA (data not shown) although a faint band was still visible in siRNA and T3-treated cells (Fig. 5B, lane 12). A subsequently performed luciferase reporter gene assay revealed the complete loss of hypoxic and T3-dependent induction (Fig. 5A).
Thyroid hormone does not stimulate the activity of the C-TAD of HIF-1α despite the increased formation of HIF-1α-p300/SRC-1 complexes.
To study whether T3, in addition to increasing HIF-1α protein levels, also induces the activity of the C-TAD, coimmunoprecipitation for the known coactivators of HIF-1 p300 and SRC-1 was performed. For both coactivators, increased amounts of complexes with HIF-1α were found (Fig. 6A). However, when the activity of the C-TAD was determined, T3 was without any effect although hypoxia increased the activity of the C-TAD significantly (Fig. 6B). Thus increased coimmunoprecipitation of HIF-1α with p300 and SRC-1 under T3 treatment probably reflects more input, i.e., HIF-1α loading, rather than supporting the notion that the interaction between HIF-1α and p300 or SRC-1 was increased.
Thyroid hormones exert a calorigenic effect on the tissue that increases the demand for oxygen. Therefore erythropoiesis, which provides the necessary oxygen capacity of the blood and the control of erythropoiesis by EPO have always been closely linked with the effects of thyroid hormones (2). Most investigators understood increased serum levels of EPO in hyperthyroidism to be caused by a greater demand for oxygen in the tissue under the action of thyroid hormones. Increased oxygen consumption was thought to cause a sufficiently low Po2 in the tissue to trigger the production of EPO (26). However, there is no experimental evidence that the action of thyroid hormone on EPO production exclusively depends on an increase in oxygen consumption. In fact, some groups found a noncalorigenic effect of thyroid hormones on erythropoiesis when oxygen consumption was not closely correlated with the increase in radio-iron incorporation into red blood cells after the application of thyroid hormone (5, 27). A direct, noncalorigenic stimulation of EPO production in vitro was revealed by direct measurement of oxygen consumption in HepG2 cells (10) and further supported by the experiment in which HepG2 cells were maintained in hypoxia due to diffusion-limited oxygen supply (24). In this setting, thyroid hormones exerted their full stimulatory effect on EPO production, although the culture conditions prevented oxygen consumption-dependent changes in the Po2 (10).
The biological meaning of the effect of thyroid hormones on EPO production and erythropoiesis appears obvious due to the tight coupling of blood oxygen capacity and oxygen demand of the tissue. In contrast, the finding that T3 stimulates the expression of the hypotensive hormone ADM in vitro and in vivo (25) is less clear, but it has been hypothesized that a better perfusion caused by vasodilation may be advantageous for the tissue under the influence of thyroid hormones (25). Both genes, EPO and ADM, have in common that they are hypoxia inducible and that the hypoxic induction depends on activation of HIF-1 (4). Herein, hypoxia-induced expression of both genes was significantly augmented by thyroid hormones (Fig. 1) and we therefore aimed at studying potential effects of T3 on HIF-1 signaling. Indeed, T3 increased HIF-1α protein accumulation, HIF-1 DNA-binding activity, and HIF-1-mediated reporter gene transcription, whereas the biologically inactive rT3 was without effect (data not shown). T3 stimulated HIF-1 under normoxia but also augmented the hypoxic activation of HIF-1. This is in support of earlier findings where T3 stimulated EPO protein synthesis in HepG2 cells and isolated perfused rat kidneys under hypoxic conditions (10).
One hypothesis to explain the increased HIF-1 activation by thyroid hormones might be that an increased O2 consumption could cause a decrease of the Po2 if O2 diffusion becomes limiting. However, our setup for hypoxic exposure makes this hypothesis very unlikely. First, direct measurement of the O2 consumption of HepG2 cells under the influence of T3 revealed no increase (10). Second, a Po2 in the gas phase of ∼20 mmHg (corresponding to 3% O2) results in a pericellular Po2 of <1 mmHg (24), even with cells of an ∼80% confluent cell monolayer as used in this study. Finally, if aggravated hypoxia accounted for the increase in HIF-1α accumulation and activation, T3 treatment should have delayed the degradation and increase the stability of HIF-1α, which was not the case (Fig. 3A). Considering that thyroid hormone might increase HIF-1α gene expression was rejected because HIF-1α mRNA levels were not elevated by T3 treatment (Fig. 2C). This finding indicated that the T3-inducible accumulation of HIF-1α protein might be due to an increase of HIF-1α protein synthesis. To test this hypothesis we used MG132, a specific inhibitor of the ubiquitin proteasome complex, to maintain ubiquitinated HIF-1α and block HIF-1α degradation. By MG132 treatment the rate of HIF-1α accumulation is at large a function of the rate of HIF-1α synthesis. T3 significantly enhanced HIF-1α protein levels in the presence of MG132 under both normoxic and hypoxic conditions (Fig. 3B). The effect of T3 was inhibited by cycloheximide, an inhibitor of translation (Fig. 3C), and direct labeling of nascent HIF-1α with [35S]methionine confirmed the stimulation of translation of HIF-1α by T3 (Fig. 3D). Moreover, HIF-1α was indispensable for T3-induced stimulation of HIF-1-dependent reporter and target gene expression as shown by knocking down HIF-1α by siRNA. Thus increased synthesis by T3 cooperates with hypoxic stabilization and accumulation of HIF-1α that is due to inhibition of prolyl hydroxylase activity (7) and ensures enhanced expression of the HIF-1 target genes EPO and ADM.
It appeared important to us to test whether thyroid hormones would also specifically increase the activity of HIF-1α. However, although T3 caused an increased coimmunoprecipitation of the known HIF-1α coactivator p300 and SRC-1, the activity of the C-TAD of HIF-1α was not stimulated by T3, although a significant hypoxic induction was observed (Fig. 6). Thus increased capture of the HIF-1α binding protein p300 and SRC-1 is most likely due to the higher input of HIF-1α in T3-treated cells, but does not indicate a specific increase of HIF-1 activity by T3. Therefore the stimulation of HIF-1-dependent gene expression by T3 appears to be solely mediated through increased HIF-1α synthesis and thus accumulation.
It is interesting that stimulation of target gene expression was first observed under hypoxic conditions when increased levels of HIF-1 are also activated by concomitant hypoxia (10). Thus the effect of thyroid hormone is similar to that observed with insulin and a series of growth factors, including insulin-like growth factor (IGF)-1, IGF-2, EGF, basic fibroblast growth factor-2 (FGF-2), HGF, and HER2 (neu) receptor tyrosine kinase (11, 21, 35, 36, 38). All of above hormones and growth factors do not stabilize HIF-1α but increase the rate of HIF-1α synthesis and would therefore act in concert with hypoxic accumulation and activation of HIF-1α.
This study was supported by a grant from the Deutsche Forschungsgemeinschaft (DFG Fa 225/18–1).
We are grateful to P. J. Ratcliffe and R. Wenger for plasmids and T. Kietzmann and K. Rutkowski for contribution in the very early phase of this study.
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked “advertisement” in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
- Copyright © 2004 the American Physiological Society