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Reactive oxygen species-driven HIF1α triggers accelerated glycolysis in endothelial cells exposed to low oxygen tension
Nuclear Medicine and Biology 45 (2016) 8–14
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Reactive oxygen species-driven HIF1α triggers accelerated glycolysis in endothelial cells exposed to low oxygen tension☆ Jin-Young Paik a,1, Kyung-Ho Jung a,b,1, Jin-Hee Lee a,b, Jin-Won Park a,b, Kyung-Han Lee a,b,⁎ a b
Department of Nuclear Medicine, Samsung Medical Center, Seoul, Republic of Korea Samsung Advanced Institute for Health Sciences & Technology, Sungkyunkwan University School of Medicine, Seoul, Republic of Korea
a r t i c l e
i n f o
Article history: Received 3 September 2016 Received in revised form 12 October 2016 Accepted 24 October 2016 Available online xxxx Keywords: Endothelial cell Glycolysis 18 F–FDG Hypoxia HIF1α ROS
a b s t r a c t Endothelial cells and their metabolic state regulate glucose transport into underlying tissues. Here, we show that low oxygen tension stimulates human umbilical vein endothelial cell 18F–fluorodeoxyglucose (18F–FDG) uptake and lactate production. This was accompanied by augmented hexokinase activity and membrane Glut-1, and increased accumulation of hypoxia-inducible factor-1α (HIF1α). Restoration of oxygen reversed the metabolic effect, but this was blocked by HIF1α stabilization. Hypoxia-stimulated 18F–FDG uptake was completely abrogated by silencing of HIF1α expression or by a specific inhibitor. There was a rapid and marked increase of reactive oxygen species (ROS) by hypoxia, and ROS scavenging or NADPH oxidase inhibition completely abolished hypoxiastimulated HIF1α and 18F–FDG accumulation, placing ROS production upstream of HIF1α signaling. Hypoxiastimulated HIF1α and 18F–FDG accumulation was blocked by the protein kinase C (PKC) inhibitor, staurosporine. The phosphatidylinositol 3-kinase (PI3K) inhibitor, wortmannin, blocked hypoxia-stimulated 18F–FDG uptake and attenuated hypoxia-responsive element binding of HIF1α without influencing its accumulation. Thus, ROS-driven HIF1α accumulation, along with PKC and PI3K signaling, play a key role in triggering accelerated glycolysis in endothelial cells under hypoxia, thereby contributing to 18F–FDG transport. © 2016 Elsevier Inc. All rights reserved.
1. Introduction Warburg's original observation of a heightened rate of glycolysis in cancer cells [1] forms the basis for 18F–fluorodeoxyglucose ( 18F–FDG) positron emission tomography (PET) imaging of patients with malignancies [2]. In addition to the widespread use of 18F–FDG PET for cancer diagnosis, the level of tumor 18F–FDG uptake serves as an indicator of treatment response and patient prognosis. This makes it imperative to have a full understanding of key factors that regulate tumor 18F–FDG uptake. The vascular endothelium serves as a conduit for supplying nutrients and oxygen to all types of tissues including tumors. Recognition of endothelia as a significant component of total tumor mass has recently prompted interest in understanding how these cells handle 18F–FDG [3–5]. Endothelial cells share with cancer cells a strong preference for glycolysis as their energy source [6,7]. Interestingly, endothelial cells can influence the metabolic state of neighboring cancer cells [8]. Moreover, a recent study revealed that endothelial cell glucose metabolism is ☆ This research was supported by the Basic Science Research Program through the National Research Foundation of Korea (NRF) funded by the Ministry of Science, ICT and Future Planning (NRF-2015R1A2A2A01006419). ⁎ Corresponding author at: Department of Nuclear Medicine, Samsung Medical Center, 50 Ilwondong, Kangnamgu, Seoul, Republic of Korea. Tel.: +82 2 3410 2630; fax: +82 2 3410 2639. E-mail address: [email protected] (K.-H. Lee). 1 Both authors contributed equally to this work. http://dx.doi.org/10.1016/j.nucmedbio.2016.10.006 0969-8051/© 2016 Elsevier Inc. All rights reserved.
a vital mediator of glucose transport from the circulation into underlying tissues [9]. These findings underscore the importance of endothelial cell metabolism on overall tumor 18F–FDG uptake. A major characteristic of tumors is their regional heterogeneity, which includes fluctuation of oxygen concentration that leads to areas with hypoxia, a key regulatory factor in tumor growth [10]. A large amount of evidence indicates that low oxygen state is associated with increased tumor 18F–FDG uptake [11,12]. Because vascular endothelia in hypoxic tissues need to survive and establish new blood vessels [13], they have evolved to thrive under low oxygen environments [14]. An important adaptive response to hypoxia programmed in endothelial cells to help ensure energy and biomass required for angiogenesis is to shift their metabolism toward glycolytic flux [15,16]. In fact, endothelial cell glycolysis could be a phenotypic switch that promotes angiogenesis when tissues are exposed to hypoxia [17]. The best understood mediator of response to oxygen deficiency is hypoxia-inducible factor-1α (HIF1α). Whereas HIF1α protein normally undergoes rapid degradation via prolyl residue hydroxylation, degradation is suppressed when oxygen tension is low. This causes a buildup of HIF1α that orchestrates the expression of genes necessary for hypoxic adaptation [10,18,19]. A recent study showed that endothelial cellspecific deletion of HIF1α significantly hinders tissue glucose uptake [9]. This underscores the importance of understanding the roles of HIF1α and subsequent mediators on 18F–FDG uptake of endothelial cells. However, in contrast to extensive studies on hypoxia-driven
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cancer cell metabolism [11,12], insight on the regulation of endothelial cell metabolism under low oxygen is relatively scarce. In this study, we thus evaluated endothelial cell glycolysis and 18F–FDG uptake under low oxygen tension, and investigated the molecular mechanisms that drive the metabolic response. 2. Materials and methods 2.1. Endothelial cell culture and exposure to hypoxia Human umbilical vein endothelial (HUVE) cells, human microvascular endothelial (HMVE) cells and calf pulmonary artery endothelial (CPAE) cells (American Type Cell Culture) were maintained in a humidified atmosphere of 5% CO2 at 37 °C in endothelial cell basal medium (Lonza, Basel, Switzerland) supplemented with epidermal growth factor, 12% fetal bovine serum, 2 g/L glucose and 100 U/ml penicillin–streptomycin. Culture media was changed every 3 days, and cells from passage 4–5 were used for experiments performed when confluence reached 80%. Exposure to hypoxia was performed by placing cells in an anaerobic chamber (ANAEROBIC SYSTEM-1029, Forma Scientific) containing 1% O2, 5% CO2 and 94% N2. A cell-permeable amidophenolic compound, 3-(2-(4-Adamantan-1yl-phenoxy)-acetylamino)-4-hydroxybenzoic acid methyl ester, that specifically inhibits hypoxia-induced HIF1α [20] was from Calbiochem (Germany). All other reagents were from Sigma-Aldrich (MO) unless otherwise specified. The effects of HIF1α inhibitor, sodium nitroprusside (SNP), N-acetyl cysteine (NAC), apocynin, cycloheximide, staurosporine and wortmannin were tested by addition to culture media 1 h prior to hypoxic exposure. 2.2. Measurement of 18F–FDG uptake 18
F–FDG was produced on a FASTlab synthesizer (GE Healthcare, Loncin, Belgium) and passed all required quality control tests. Its radiochemical purity was higher than 99%, and specific activity was approximately 20–22 Ci/mmol based on the amount of precursor used. Cells seeded at 1 × 10 5 cells per wells of a 12-well culture plate were incubated for 40 min with 370 kBq 18F–FDG added to 1.5 ml of media. Cells were washed with phosphate buffered saline (PBS), lysed in distilled water, and measured for radioactivity on a high-energy gamma counter (Wallac). 18F–FDG uptake of each sample was normalized by protein content as determined by Bradford assays. 2.3. Measurement of total hexokinase activity Total cellular hexokinase activity was measured as previously described [4]. Briefly, cells were homogenized in homogenizing buffer (50 mM triethanolamine and 5 mM MgCl2; pH 7.6) and centrifuged at 1000 g at 4 °C for 5 min. Supernatants were mixed with 20 °C preincubated homogenizing buffer supplemented with 0.5 mM glucose, 5 mM adenosine triphosphate, 0.25 mM reduced nicotineamide adenine dinucleotide phosphate and 6 units of glucose–6-phosphate dehydrogenase. The reaction mixture was repeatedly measured for 340 nm absorbance, and hexokinase activity in mU/mg protein was determined from a standard curve with 1 unit defined as enzyme activity that phosphorylates 1 μmol of glucose per min at 20 °C. 2.4. Measurement of nitric oxide release Nitric oxide release was measured from 100 μl of culture media that was sampled and incubated with and an equal volume of Griess reagent containing 1% sulfanilamide, 0.1% N-(1-naphthyl)ehtylendiamine dihydrochloride, and 2.5% phosphoric acid for 10 min. Spectrophotometric absorbance at 540 nm was measured and results were expressed as concentration relative to controls.
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2.5. Immunoblot of membrane Gkut-1 expression Two 150 mm plates of cells were washed with PBS and solubilized in 500 μl lysis buffer containing 0.0856 g/ml sucrose, 10 mM/ml HEPES, 25 μM/ml EDTA, 10 μg/ml aprotinin, 10 μg/ml leupeptin, and 1 mM phenylmethylsulfonylfluoride. Following centrifugation at 1000 g for 20 min, the supernatant was transferred to an ultracentrifugation tube containing 1.5 ml lysis buffer (0.0856 g/ml sucrose, 10 mM/ml HEPES and 10 mM/ml MgCl2) and incubated at 4 °C for 1 h. Membrane fraction protein was obtained by high-speed centrifugation at 45,000 rpm for 60 min, dissolved in PBS, and separated (20 μg) on a 10% polyacrylamide gel. After transfer to a hydrobond ECL nitrocellulose membrane (Amersham Biosciences, NJ, USA) and overnight reaction at 4 °C with a polyclonal anti-human Glut-1 antibody (Calbiochem, Frankfurter, Germany; 1:1000), immunoreactive protein was visualized by 1 h incubation at room temperature with a horse-radish peroxidase-conjugated anti-rabbit IgG secondary antibody (Caltag, Buckingham, UK; 1:1000) and exposure on a high performance chemiluminescence film. 2.6. HIF-1α immunoblotting HIF-1α immunoblotting was performed with immune-precipitated protein. Briefly, cells were dissolved in ice-cold RIPA buffer containing 0.079 g/ml Tris base, 0.09 g/ml NaCl, 10% NP-40, 10 μM/L EDTA, 10 μg/ ml aprotinin, 10 μg/ml leupeptin, 1 mM phenylmethyl sulfonylfluoride, and centrifugation was performed at 14,000 g for 15 min. The supernatant was reduced of nonspecific binding by 10 min incubation with 1 ml pre-cleared protein-G sepharose at 4 °C, centrifuged at 14,000 g for 10 min, and transferred to a new tube. The sample was diluted in PBS to 1 mg/ml and incubated with anti-HIF-1α antibody (Novus Biological, CO; 1:500) at 4 °C for 2 h on a shaker. The immunocomplex was captured by gentle rocking overnight at 4 °C after adding 50 μl of proteinG sepharose bead slurry, followed by centrifugation. The immunoprecipitate pellet was washed 2–4 times with 1 ml RIPA buffer and resuspended in 50 μl of 2× electrophoresis sample buffer. Protein samples were boiled for 4 min and separated on a 10% polyacrylamide gel and membrane transferred as above. The membrane was reacted at 4 °C for 24 h with anti-HIF-1α antibody (BD, NJ; 1:1000) and anti-HIF-1α antibody (Novus Biologicals, CO; 1:1000). Immune-reactive protein was visualized as above using an anti-mouse IgG secondary antibody (Caltag, Buckingham, UK; 1:1000). 2.7. HIF-1α binding to hypoxia response element (HRE) Cells were washed twice with ice-cold PBS and centrifuged at 250 g for 5 min. The cell pellet was washed in hypotonic buffer containing 10 mM HEPES (pH 7.9), 1.5 mM MgCl2, 10 mM KCl, 2 mM DTT, 2 mM sodium vandate, 4 μg/ml pepstatin, 4 μg/ml leupeptin, and 4 μg/ml aprotitnin, and incubated on ice for 15 min. Cells were homogenized by repeated passage through a 26G needle, and the nuclear fraction was obtained by centrifugation at 8000 g at 4 °C for 20 min. The pellet was suspended in an extraction buffer containing 20 mM HEPES (pH 7.9), 1.5 mM MgCl2, 0.42 M NaCl, 0.2 mM EDTA, 2 mM DTT, 1.0% Igepal CA-630, 25% glycerol, sodium vandate, 4 μg/ml pepstatin, 4 μg/ml leupeptin, and 4 μg/ml aprotitnin. Nuclear debris was removed by centrifugation at 16,000 g, 4 °C for 30 min, and the final nuclear extract was adjusted to 2 mg/ml with cold PBS. HIF-1α binding to HRE was measured with Trans AM™ HIF-1α kit [21] from Active Motif (Carlsbad, CA). Briefly, 5 μg/ml of nuclear extracts were added to a 96-well plate to which oligonucleotides containing an HRE (5´-TACGTGCT-3´) from the erythropoietin gene were immobilized. This oligonucleotide binds HIF-1α as a transcription factor with specific DNA binding activity. The plate was reacted with a primary antibody against HIF-1α followed by a secondary HRP-conjugated antibody, and spectrophotometry was performed at 450 nm using a reference wavelength of 655 nm.
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2.8. Inhibition of HIF-1α by RNA interference HIF-1α silencing was performed with 20–25 nucleotide targetspecific small interfering RNA (siRNA) designed to knock-down human HIF-1α expression (Santa Cruz Biotechnology, CA). Briefly, to a sterile tube was added 100 μl of transfection medium (Opti-MEM, Lonza, Basel, Switzerland) and 6 μl of lipofectamine™ 2000 (invitrogen, CA). HIF-1α inactivating siRNA was added to a final concentration of 100 nM and the mixture was incubated at room temperature for 45 min. The mixture was added to media of 60% confluent cells on 6-well plates that was seeded 24 h earlier. After vigorous agitation (for even siRNA dispersion), cells were incubated in a humidified atmosphere of 5% CO2 at 37 °C for 6 h. Cells were washed twice with transfection medium, fresh culture media were replaced, and hypoxia experiments were performed 2 days later.
Time course experiments in HUVE cells showed that 18F–FDG uptake began to rapidly rise after 4 h of hypoxia and reached a relative plateau between 6 and 16 h (Fig. 1A). This was accompanied by a similar temporal increase of lactate release by hypoxia that reached 198.0 ± 17.2% of controls by 24 h (Fig. 1B). These findings are consistent with a hypoxiainduced shift of metabolism toward glycolytic flux. When the two major determents of cellular glucose uptake were evaluated, hexokinase activity also displayed a temporal increase paralleling that of 18F–FDG uptake and lactate release. This reached 2.8 ± 0.1 fold of norrmoxic cells by 6 h of hypoxia (Fig. 1B). Plasma membrane Glut-1 expression was also increased by hypoxia, which was completely blocked by cycloheximide (Fig. 1B).
3.2. HIF-1α accumulates by hypoxia and is sufficient to enhance uptake
18
F–FDG
2.9. Data analysis Results are expressed as mean ± SE of data obtained from two separate experiments or mean ± SD of triplicate samples from a single representative experiment. Student's tests were used to compare 2 groups and one-way ANOVA with Tukey post-hoc tests were used to compare 3 or more groups. P b 0.05 was considered significant. 3. Results 3.1. Hypoxia stimulates endothelial cell hexokinase and glut-1
18
F–FDG uptake and glycolysis via
HUVE cells, HMVE cells and CPAE cells all showed significant augmentation of 18F–FDG uptake by 16 h hypoxia exposure to 178.8 ± 6.2%, 215.4 ± 7.6% and 179.7 ± 15.4% of normoxic cells, respectively (Fig. 1A).
When HUVE cells were exposed to 16 h of hypoxia, immunoblots displayed a substantial accumulation of HIF-1α protein to 325.6 ± 64.2% of controls (Fig. 2 A). In contrast, there was no increase of HIF2α protein (data not shown). Upon oxygen restoration after hypoxia, there was a prompt 32.3% decline of HIF-1α protein level by 4 h (Fig. 2A). However, in the presence of the HIF-1α stabilizing agent dimethyloxallylglycine (DMOG), a high HIF-1α level was maintained at 316.2 ± 100.4% of controls after 4 h of reoxygenation (Fig. 2A). Similar to HIF-1α level, 18F–FDG uptake that reached 236.2 ± 100.4% of controls by hypoxia returned to 133.9 ± 3.5% after 2 h of oxygen restoration (Fig. 2A). However, high 18F–FDG uptake was maintained in the presence of DMOG (Fig. 2A). DMOG also increased 18F– FDG uptake of normoxic cells to 124.0 ± 4.5% of untreated cells by 16 h (Fig. 2B). Taken together, these results indicate that HIF-1α plays a central role in mediating hypoxia-stimulated HUVE cell glycolysis.
Fig. 1. Effect of hypoxia on endothelial cell glucose metabolism. (A) Effect of 1% O2 on 18F–FDG uptake of endothelial cell lines (left), and time-course of hypoxia-stimulated 18F–FDG uptake in HUVE cells (right). (B) Time-course of hypoxia effects on HUVE cell lactate release (left) and hexokinase activity (middle), and effect of 24 h hypoxia with or without cycloheximde on membrane Glut-1 expression (right). Curves are fitted by non-linear regression. Results are mean ± SD of triplicate samples from a single experiment (A, left), or mean ± SE of values of six samples obtained from 2 independent experiments (time course data). †: p b 0.001, ‡ p b 0.0001 compared to normoxic controls.
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Fig. 2. Association between HIF1α accumulation and hypoxia-stimulated 18F–FDG uptake. (A) Effect of 16 h hypoxia and subsequent 4 h reoxygenation on HUVE cell HIF1α level (left), and time course of reversal of hypoxia-stimulated 18F–FDG uptake during reoxygenation with or without HIF1α stabilization by 50 μM DMOG (right). Data are mean ± SE of 6 samples from 2 separate experiments (HIF1α) or 9 samples from 3 separate experiments (18F–FDG uptake). (B) Stimulatory effect of DMOG on 18F–FDG uptake of normoxic HUVE cells. Data are mean ± SD of triplicate samples from a single experiment. *, p b 0.05; †, p b 0.001; ‡, p b 0.0001, compared to normoxic controls (A, left and B) or cells reoxygenated without DMOG (A, right).
3.3. HIF-1α inhibition or silencing completely abrogates hypoxia-stimulated 18 F–FDG uptake Accumulation of HIF-1α protein by hypoxia was completely blocked when HUVE cells were treated with a specific HIF-1α inhibitor or underwent gene silencing by transfection with specific siRNA (Fig. 3A). In both of these treatments, this was accompanied by complete
abrogation of the ability of hypoxia to augment 18F–FDG uptake, which decreased from 204.5 ± 13.4% to 115.7 ± 9.0% and 104.3 ± 2.6% of normoxic controls, respectively (Fig. 3B). Normoxic cell 18F–FDG uptake was not affected by the treatments. Also, transfection with control siRNA did not influence hypoxia-stimulated 18F–FDG uptake. Together, these results demonstrate that increased HIF-1α accumulation is necessary for the stimulatory effect of hypoxia on HUVE cell glycolysis.
Fig. 3. HIF1α is required for hypoxia to augment 18F–FDG uptake. (A) HUVE cell HIF1α accumulation stimulated by 24 h hypoxia is completely blocked by a specific inhibitor or by gene silencing with HIF1α specific siRNA. (B) Effects of treatment with an HIF1α inhibitor (left) or transfection with HIF1α specific or nonspecific siRNA (right) on normoxic and hypoxic HUVE cell 18F–FDG uptake. All data are mean ± SD of triplicate samples from a single experiment. **, p b 0.005; †, p b 0.001, compared to hypoxic cells without treatment.
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3.4. ROS plays a central role in hypoxia-stimulated HIF-1α accumulation and 18F–FDG uptake ROS and nitric oxide play important roles in angiogenesis as well as endothelial cell survival and metabolism. Measurements of these gases revealed that HUVE cell ROS production is markedly elevated by 1 h hypoxic exposure to 447.6 ± 117.2% of controls (Fig. 4A). On the other hand, whereas NO concentration markedly increased by adding sodium nitroprusside as donor (546.3 ± 15.5% of controls), there was no increase of nitric oxide production by hypoxia alone (Fig. 4A). HUVE cells were then evaluated for the effects of the ROS scavenger NAC and NADPH oxidase inhibitor apocynin. These treatments near completely blocked the increase of ROS production by hypoxia, whereas ROS production in normoxic cells were largely unaffected (Fig. 4B). Both of these treatments also completely abolished the capacity of hypoxia to stimulate 18F–FDG uptake (Fig. 4C) and HIF-1α accumulation (Fig. 4D). 3.5. Involvement of protein synthesis, and PKC and PI3K pathways Involvement of candidate intracellular signaling pathways in the metabolic effect of hypoxia was tested using specific inhibitors. Firstly, cycloheximide completely blocked hypoxia-stimulated 18F–FDG uptake and HIF-1α accumulation (Fig. 5A), indicating the requirement of both responses for new protein synthesis. Similarly, protein kinase C (PKC) inhibition with staurosporine completely blocked hypoxia-induced 18 F–FDG uptake and HIF-1α accumulation (Fig. 5B). In comparison, phosphatidylinositol 3-kinase (PI3K) inhibition with wortmannin effectively blocked the ability to augment 18F–FDG uptake, but not HIF-1α accumulation (222.1 ± 13.3% of controls; Fig. 5B). The requirement of PI3K activity for hypoxia-stimulated 18F–FDG uptake was further confirmed with PI3K specific siRNA that completely suppressed this response whereas nonspecific siRNA did not (Fig. 5C). Finally, HIF-1α binding to HRE in nuclear extracts from HUVE cells was enhanced to
2.51 ± 0.02 fold of controls by hypoxia, but this was significantly suppressed by wortmannin as well as by cycloheximide (Fig. 5C). This suggests that although PI3K activity is not required for hypoxia-induced HIF1α accumulation, it is necessary for HIF1α binding to HRE. 4. Discussion Tumor 18F–FDG uptake is directly and indirectly influenced by the metabolic state of vascular endothelial cells. Oxygen deprivation is a strong stimulus for new vessel formation, and endothelial cells respond by adjusting their metabolism to meet the demands required for survival and proliferation. Our study confirms that hypoxia shifts HUVE cell metabolism toward glycolytic flux. This metabolic effect was accompanied by upregulated Glut-1 expression and hexokinase activity. Furthermore, HIF1α accumulation driven by increased ROS production played a key role in the metabolic response. Stimulation of glucose uptake by low oxygen tension has previously been observed in HUVE and bovine endothelial cells [15], and in retinal capillary endothelial cells [16]. While HUVE cells are the most commonly studied endothelial cells, endothelial function and phenotype can be influenced by the vascular bed of origin. For instance, microvascular endothelial cells may respond differently with HUVE cells to cytokines [22]. However, our findings show that HUVE cells, HMVE cells and CPAE cells respond to hypoxia with similar elevations in FDG uptake. In our study, enhancement of 18F–FDG uptake was accompanied by a closely paralleled temporal rise in lactate release, indicating that the consumed glucose largely underwent glycolytic breakdown rather than oxidative phosphorylation. Previous reports attributed the augmented glucose uptake by hypoxia to glucose transporter upregulation [15,16]. Our results corroborated these findings by showing hypoxiainduced increase of plasma membrane-localized Glut-1, the predominant glucose transporter isoform in endothelial cells. Our results further showed that enhanced hexokinase activity also contributes to high 18F–
Fig. 4. Role of reactive oxygen species (ROS) on hypoxia-stimulated 18F–FDG uptake. (A) Hypoxia substantially increased HUVE cell production of ROS (left) but not nitric oxide (NO; right). Sodium nitroprusside (SNP; 100 μM) treated cells were used as a positive control for NO production. (B,C) Complete abrogation of hypoxia-stimulated ROS production (B) and 18F–FDG uptake (C) by N-acetyl cysteine (NAC; left) and apocynin (NAPDH oxidase inhibitor, right). (D) Effects of NAC and apocynin on hypoxia-induced HIF1α accumulation. Results are mean ± SE of values obtained from 2 separate experiments (n = 6; A-C), or mean ± SD of triplicate samples from a single experiment (D). †, p b 0.001; ‡, p b 0.0001, compared to normoxic controls (A) or hypoxic cells without treatment (D).
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Fig. 5. Inhibition of candidate signaling pathways on hypoxia effect. (A,B) Effects of 200 nM cycloheximide (CHX; A), 4 μM staurosporine (ST; B) and 200 nM wortmannin (WM; B) on hypoxia-stimulated HUVE cell 18F–FDG uptake (left) and HIF1α accumulation (right). (C) 18F–FDG uptake in normoxic and hypoxic HUVE cells gene-silenced with PI3K specific or control siRNA (left). Increased hypoxia-responsive element (HRE) binding of HIF1α extracted from HUVE cells after 24 h hypoxia is suppressed by WM or CHX (right). Results are mean ± SE of values of 9 samples from 3 separate experiments (FDG uptake in A and B), or mean ± SD of values of triplicate samples from a single experiment (C and HIF1α levels). †: p b 0.001; ‡ p b 0.0001 compared to hypoxic cells without treatment.
FDG uptake. Whereas glycolytic enzyme activation is considered a characteristic response of cancer cells to hypoxia, we are not aware of its description in endothelial cells. HIF1α is a transcriptional factor subunit that serves as a master regulator of oxygen homeostatsis [13]. Our results confirm that HUVE cells exposed to hypoxia buildup HIF1α protein level as they augment 18F– FDG uptake. The two responses were further linked in that both responses were revered when normal oxygen tension was restored after hypoxic treatment. Furthermore, stabilizing HIF1α protein by retarding its degradation during reoxygenation with the prolyl hydroxylase inhibitor, DMOG, was sufficient to maintain a high level of 18F–FDG uptake. In addition, normoxic cells also showed modest but significant elevation 18 F–FDG uptake by DMOG treatment alone. Importantly, blocking of HIF1α accumulation by treatment with a specific HIF1α inhibitor or gene silencing with specific siRNA completely abolished the ability of hypoxia to augment 18F–FDG uptake. Collectively, these findings demonstrate a key role played by HIF1α accumulation that is both necessary and sufficient for hypoxia to promote HUVE cell glycolysis and 18 F–FDG uptake. Tightly regulated generation of ROS and nitrogen species that can fine tune downstream signaling in cell-specific manners are critical constituents of cellular responses to various stimuli [23]. ROS production regulates several aspects of vascular and endothelial function including angiogenesis [23]. A previous study showed that endothelial cells increase ROS production under hypoxic conditions via the mitochondrial respiratory chain or NADPH oxidase activation [24]. In our study, the ROS scavenger, NAC, totally suppressed ROS production and HIF1α accumulation as well as 18F–FDG uptake by hypoxia. Furthermore, NADPH oxidase-mediated ROS production was a major effector for the metabolic effect of hypoxia on HUVE cells, since both HIF1α accumulation and 18F–FDG uptake responses were completely blocked by the NADPH oxidase inhibitor, apocynin. These findings put ROS production
at an early event upstream of HIF1α accumulation in the metabolic response to hypoxia. This notion is consistent with previous proposals that ROS generation may be an initial step for hypoxia-sensing in endothelial cells [25]. We also tested the possibility of NO involvement, since hypoxia has been shown to stimulate NO release in certain endothelial cells [26] and our group previously demonstrated NO to be a powerful simulator of HUVE cell 18F–FDG uptake [4]. However, the results of the present study showed that hypoxia does not increase NO release, excluding the involvement of NO signaling in the metabolic effect of hypoxia on HUVE cells. HIF1α accumulation can occur either through slowing of its proteosomal degradation or by increased synthesis through growth factor signaling [27]. The role of protein synthesis on a cell response can be tested by treatment with cycloheximide. In our study, this caused a total loss of hypoxia-stimulated HIF1α accumulation and 18F–FDG uptake. Similar losses of hypoxia-induced HIF1α buildup [28] and deoxyglucose uptake by cycloheximide have been previously observed in other cell types [29]. These results suggest that new protein synthesis is required to mediate the metabolic effect of hypoxia. HIF1α is rapidly destroyed in normoxic tumor cells with a half-life of approximately 5 min, whereas stabilization in hypoxic states will prolong its half-life to 2–4 h [10]. In HUVE cells, HIF1α half-life may be shorter, even under hypoxic conditions (about 30 min) [30]. Our data imply that hypoxic HUVE cells require continuous protein synthesis for HIF1α accumulation and increasing 18F–FDG uptake. On exploring candidate signaling pathways that may be involved, the potent PKC inhibitor staurosporine completely blocked the ability of hypoxia to stimulate both HIF1α accumulation and 18F–FDG uptake. This finding could be related to the fact that the PKC pathway can be activated by hypoxia [31] and is implicated in the angiogenic response [32]. The specific PI3K inhibitor wortmannin also inhibited stimulation of 18F–FDG uptake by hypoxia, whereas it had no influence on HIF1α
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Fig. 6. Scheme of proposed molecular mechanism for hypoxia-stimulated endothelial cell glycolysis.
accumulation. The PI3K pathway is activated by growth factors and is closely involved in glucose metabolism as well as angiogenesis. A previous study reported that certain HRE promoters bind HIF1α in a PI3Kdependent manner [33]. Hence, it is possible that PI3K is required for HIF1α binding to HRE and subsequent stimulation of glycolysis, even though it is not necessary for HIF1α accumulation. A schematic of the proposed molecular mechanism involved in hypoxia-stimulated endothelial cell glycolysis and 18F–FDG uptake is illustrated in Fig. 6. In conclusion, hypoxia stimulates HUVE cell glycolysis, in a manner accompanied by upregulated Glut-1 expression and hexokinase activity. HIF1α accumulation driven by increased ROS generation plays a central role in this metabolic effect, which also involves PKC and PI3K signaling. The resultant increase of endothelial cell 18F–FDG uptake may contribute to greater transport of the substrate to underlying cancer cells, which could together contribute to higher tumor 18F–FDG uptake in oxygen-deprived regions. Sources of funding This work was supported by the Basic Science Research Program through the National Research Foundation of Korea (NRF) funded by the Ministry of Science, ICT and Future Planning (NRF-2015R1A2A2A01006419). Disclosures None. References [1] Warburg O. On the origin of cancer cells. Science 1956;123:309–14. [2] Farwell MD, Pryma DA, Mankoff DA. PET/CT imaging in cancer: current applications and future directions. Cancer 2014;120:3433–45. [3] Maschauer S, Prante O, Hoffmann M, Deichen JT, Kuwert T. Characterization of FDG uptake in human endothelial cells in vitro. J Nucl Med 2004;45:455–60. [4] Paik JY, Lee KH, Ko BH, Choe YS, Choi Y, Kim BT. Nitric oxide stimulates FDG uptake in human endothelial cells through increased hexokinase activity and GLUT1 expression. J Nucl Med 2005;46:365–70. [5] Paik JY, Koh BH, Jung KH, Lee KH. Fibronectin stimulates endothelial cell FDG uptake via focal adhesion kinase mediated phosphatidylinositol 3-kinase/Akt signaling. J Nucl Med 2009;50:618–24.
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Hypoxia upregulates glucose transport activity through an adenosine-mediated increase of GLUT1 expression in retinal capillary endothelial cells. Diabetes 1998;47:1480–8. [17] Cantelmo AR, Brajic A, Carmeliet P. Endothelial metabolism driving angiogenesis: emerging concepts and principles. Cancer J 2015;21:244–9. [18] Carmeliet P, Dor Y, Herbert JM, Fukumura D, Brusselmans K, Dewerchin M, et al. Role of HIF-1alpha in hypoxia-mediated apoptosis, cell proliferation and tumour angiogenesis. Nature 1998;394:485–90. [19] Hickey MM, Simon MC. Regulation of angiogenesis by hypoxia and hypoxiainducible factors. Curr Top Dev Biol 2006;76:217–57. [20] Coutts AS, Pires IM, Weston L, Buffa FM, Milani M, Li JL, et al. Hypoxia-driven cell motility reflects the interplay between JMY and HIF-1α. Oncogene 2011;30:4835–42. [21] zur Nedden S, Tomaselli B, Baier-Bitterlich G. HIF-1 alpha is an essential effector for purine nucleoside-mediated neuroprotection against hypoxia in PC12 cells and primary cerebellar granule neurons. J Neurochem 2008;105:1901–14. [22] Lidington EA, Moyes DL, McCormack AM, Rose ML. A comparison of primary endothelial cells and endothelial cell lines for studies of immune interactions. Transpl Immunol 1999;7:239–46. [23] Panieri E, Santoro MM. ROS signaling and redox biology in endothelial cells. Cell Mol Life Sci 2015;72:2281–303. [24] Schäfer M, Schäfer C, Ewald N, Piper HM, Noll T. Role of redox signaling in the autonomous proliferative response of endothelial cells to hypoxia. Circ Res 2003;92:1010–5. [25] Pearlstein DP, Ali MH, Mungai PT, Hynes KL, Gewertz BL, Schumacker PT. Role of mitochondrial oxidant generation in endothelial cell responses to hypoxia. Arterioscler Thromb Vasc Biol 2002;22:566–73. [26] Justice JM, Tanner MA, Myers PR. 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Nuclear Medicine and Biology journal homepage: www.elsevier.com/locate/nucmedbio
Reactive oxygen species-driven HIF1α triggers accelerated glycolysis in endothelial cells exposed to low oxygen tension☆ Jin-Young Paik a,1, Kyung-Ho Jung a,b,1, Jin-Hee Lee a,b, Jin-Won Park a,b, Kyung-Han Lee a,b,⁎ a b
Department of Nuclear Medicine, Samsung Medical Center, Seoul, Republic of Korea Samsung Advanced Institute for Health Sciences & Technology, Sungkyunkwan University School of Medicine, Seoul, Republic of Korea
a r t i c l e
i n f o
Article history: Received 3 September 2016 Received in revised form 12 October 2016 Accepted 24 October 2016 Available online xxxx Keywords: Endothelial cell Glycolysis 18 F–FDG Hypoxia HIF1α ROS
a b s t r a c t Endothelial cells and their metabolic state regulate glucose transport into underlying tissues. Here, we show that low oxygen tension stimulates human umbilical vein endothelial cell 18F–fluorodeoxyglucose (18F–FDG) uptake and lactate production. This was accompanied by augmented hexokinase activity and membrane Glut-1, and increased accumulation of hypoxia-inducible factor-1α (HIF1α). Restoration of oxygen reversed the metabolic effect, but this was blocked by HIF1α stabilization. Hypoxia-stimulated 18F–FDG uptake was completely abrogated by silencing of HIF1α expression or by a specific inhibitor. There was a rapid and marked increase of reactive oxygen species (ROS) by hypoxia, and ROS scavenging or NADPH oxidase inhibition completely abolished hypoxiastimulated HIF1α and 18F–FDG accumulation, placing ROS production upstream of HIF1α signaling. Hypoxiastimulated HIF1α and 18F–FDG accumulation was blocked by the protein kinase C (PKC) inhibitor, staurosporine. The phosphatidylinositol 3-kinase (PI3K) inhibitor, wortmannin, blocked hypoxia-stimulated 18F–FDG uptake and attenuated hypoxia-responsive element binding of HIF1α without influencing its accumulation. Thus, ROS-driven HIF1α accumulation, along with PKC and PI3K signaling, play a key role in triggering accelerated glycolysis in endothelial cells under hypoxia, thereby contributing to 18F–FDG transport. © 2016 Elsevier Inc. All rights reserved.
1. Introduction Warburg's original observation of a heightened rate of glycolysis in cancer cells [1] forms the basis for 18F–fluorodeoxyglucose ( 18F–FDG) positron emission tomography (PET) imaging of patients with malignancies [2]. In addition to the widespread use of 18F–FDG PET for cancer diagnosis, the level of tumor 18F–FDG uptake serves as an indicator of treatment response and patient prognosis. This makes it imperative to have a full understanding of key factors that regulate tumor 18F–FDG uptake. The vascular endothelium serves as a conduit for supplying nutrients and oxygen to all types of tissues including tumors. Recognition of endothelia as a significant component of total tumor mass has recently prompted interest in understanding how these cells handle 18F–FDG [3–5]. Endothelial cells share with cancer cells a strong preference for glycolysis as their energy source [6,7]. Interestingly, endothelial cells can influence the metabolic state of neighboring cancer cells [8]. Moreover, a recent study revealed that endothelial cell glucose metabolism is ☆ This research was supported by the Basic Science Research Program through the National Research Foundation of Korea (NRF) funded by the Ministry of Science, ICT and Future Planning (NRF-2015R1A2A2A01006419). ⁎ Corresponding author at: Department of Nuclear Medicine, Samsung Medical Center, 50 Ilwondong, Kangnamgu, Seoul, Republic of Korea. Tel.: +82 2 3410 2630; fax: +82 2 3410 2639. E-mail address: [email protected] (K.-H. Lee). 1 Both authors contributed equally to this work. http://dx.doi.org/10.1016/j.nucmedbio.2016.10.006 0969-8051/© 2016 Elsevier Inc. All rights reserved.
a vital mediator of glucose transport from the circulation into underlying tissues [9]. These findings underscore the importance of endothelial cell metabolism on overall tumor 18F–FDG uptake. A major characteristic of tumors is their regional heterogeneity, which includes fluctuation of oxygen concentration that leads to areas with hypoxia, a key regulatory factor in tumor growth [10]. A large amount of evidence indicates that low oxygen state is associated with increased tumor 18F–FDG uptake [11,12]. Because vascular endothelia in hypoxic tissues need to survive and establish new blood vessels [13], they have evolved to thrive under low oxygen environments [14]. An important adaptive response to hypoxia programmed in endothelial cells to help ensure energy and biomass required for angiogenesis is to shift their metabolism toward glycolytic flux [15,16]. In fact, endothelial cell glycolysis could be a phenotypic switch that promotes angiogenesis when tissues are exposed to hypoxia [17]. The best understood mediator of response to oxygen deficiency is hypoxia-inducible factor-1α (HIF1α). Whereas HIF1α protein normally undergoes rapid degradation via prolyl residue hydroxylation, degradation is suppressed when oxygen tension is low. This causes a buildup of HIF1α that orchestrates the expression of genes necessary for hypoxic adaptation [10,18,19]. A recent study showed that endothelial cellspecific deletion of HIF1α significantly hinders tissue glucose uptake [9]. This underscores the importance of understanding the roles of HIF1α and subsequent mediators on 18F–FDG uptake of endothelial cells. However, in contrast to extensive studies on hypoxia-driven
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cancer cell metabolism [11,12], insight on the regulation of endothelial cell metabolism under low oxygen is relatively scarce. In this study, we thus evaluated endothelial cell glycolysis and 18F–FDG uptake under low oxygen tension, and investigated the molecular mechanisms that drive the metabolic response. 2. Materials and methods 2.1. Endothelial cell culture and exposure to hypoxia Human umbilical vein endothelial (HUVE) cells, human microvascular endothelial (HMVE) cells and calf pulmonary artery endothelial (CPAE) cells (American Type Cell Culture) were maintained in a humidified atmosphere of 5% CO2 at 37 °C in endothelial cell basal medium (Lonza, Basel, Switzerland) supplemented with epidermal growth factor, 12% fetal bovine serum, 2 g/L glucose and 100 U/ml penicillin–streptomycin. Culture media was changed every 3 days, and cells from passage 4–5 were used for experiments performed when confluence reached 80%. Exposure to hypoxia was performed by placing cells in an anaerobic chamber (ANAEROBIC SYSTEM-1029, Forma Scientific) containing 1% O2, 5% CO2 and 94% N2. A cell-permeable amidophenolic compound, 3-(2-(4-Adamantan-1yl-phenoxy)-acetylamino)-4-hydroxybenzoic acid methyl ester, that specifically inhibits hypoxia-induced HIF1α [20] was from Calbiochem (Germany). All other reagents were from Sigma-Aldrich (MO) unless otherwise specified. The effects of HIF1α inhibitor, sodium nitroprusside (SNP), N-acetyl cysteine (NAC), apocynin, cycloheximide, staurosporine and wortmannin were tested by addition to culture media 1 h prior to hypoxic exposure. 2.2. Measurement of 18F–FDG uptake 18
F–FDG was produced on a FASTlab synthesizer (GE Healthcare, Loncin, Belgium) and passed all required quality control tests. Its radiochemical purity was higher than 99%, and specific activity was approximately 20–22 Ci/mmol based on the amount of precursor used. Cells seeded at 1 × 10 5 cells per wells of a 12-well culture plate were incubated for 40 min with 370 kBq 18F–FDG added to 1.5 ml of media. Cells were washed with phosphate buffered saline (PBS), lysed in distilled water, and measured for radioactivity on a high-energy gamma counter (Wallac). 18F–FDG uptake of each sample was normalized by protein content as determined by Bradford assays. 2.3. Measurement of total hexokinase activity Total cellular hexokinase activity was measured as previously described [4]. Briefly, cells were homogenized in homogenizing buffer (50 mM triethanolamine and 5 mM MgCl2; pH 7.6) and centrifuged at 1000 g at 4 °C for 5 min. Supernatants were mixed with 20 °C preincubated homogenizing buffer supplemented with 0.5 mM glucose, 5 mM adenosine triphosphate, 0.25 mM reduced nicotineamide adenine dinucleotide phosphate and 6 units of glucose–6-phosphate dehydrogenase. The reaction mixture was repeatedly measured for 340 nm absorbance, and hexokinase activity in mU/mg protein was determined from a standard curve with 1 unit defined as enzyme activity that phosphorylates 1 μmol of glucose per min at 20 °C. 2.4. Measurement of nitric oxide release Nitric oxide release was measured from 100 μl of culture media that was sampled and incubated with and an equal volume of Griess reagent containing 1% sulfanilamide, 0.1% N-(1-naphthyl)ehtylendiamine dihydrochloride, and 2.5% phosphoric acid for 10 min. Spectrophotometric absorbance at 540 nm was measured and results were expressed as concentration relative to controls.
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2.5. Immunoblot of membrane Gkut-1 expression Two 150 mm plates of cells were washed with PBS and solubilized in 500 μl lysis buffer containing 0.0856 g/ml sucrose, 10 mM/ml HEPES, 25 μM/ml EDTA, 10 μg/ml aprotinin, 10 μg/ml leupeptin, and 1 mM phenylmethylsulfonylfluoride. Following centrifugation at 1000 g for 20 min, the supernatant was transferred to an ultracentrifugation tube containing 1.5 ml lysis buffer (0.0856 g/ml sucrose, 10 mM/ml HEPES and 10 mM/ml MgCl2) and incubated at 4 °C for 1 h. Membrane fraction protein was obtained by high-speed centrifugation at 45,000 rpm for 60 min, dissolved in PBS, and separated (20 μg) on a 10% polyacrylamide gel. After transfer to a hydrobond ECL nitrocellulose membrane (Amersham Biosciences, NJ, USA) and overnight reaction at 4 °C with a polyclonal anti-human Glut-1 antibody (Calbiochem, Frankfurter, Germany; 1:1000), immunoreactive protein was visualized by 1 h incubation at room temperature with a horse-radish peroxidase-conjugated anti-rabbit IgG secondary antibody (Caltag, Buckingham, UK; 1:1000) and exposure on a high performance chemiluminescence film. 2.6. HIF-1α immunoblotting HIF-1α immunoblotting was performed with immune-precipitated protein. Briefly, cells were dissolved in ice-cold RIPA buffer containing 0.079 g/ml Tris base, 0.09 g/ml NaCl, 10% NP-40, 10 μM/L EDTA, 10 μg/ ml aprotinin, 10 μg/ml leupeptin, 1 mM phenylmethyl sulfonylfluoride, and centrifugation was performed at 14,000 g for 15 min. The supernatant was reduced of nonspecific binding by 10 min incubation with 1 ml pre-cleared protein-G sepharose at 4 °C, centrifuged at 14,000 g for 10 min, and transferred to a new tube. The sample was diluted in PBS to 1 mg/ml and incubated with anti-HIF-1α antibody (Novus Biological, CO; 1:500) at 4 °C for 2 h on a shaker. The immunocomplex was captured by gentle rocking overnight at 4 °C after adding 50 μl of proteinG sepharose bead slurry, followed by centrifugation. The immunoprecipitate pellet was washed 2–4 times with 1 ml RIPA buffer and resuspended in 50 μl of 2× electrophoresis sample buffer. Protein samples were boiled for 4 min and separated on a 10% polyacrylamide gel and membrane transferred as above. The membrane was reacted at 4 °C for 24 h with anti-HIF-1α antibody (BD, NJ; 1:1000) and anti-HIF-1α antibody (Novus Biologicals, CO; 1:1000). Immune-reactive protein was visualized as above using an anti-mouse IgG secondary antibody (Caltag, Buckingham, UK; 1:1000). 2.7. HIF-1α binding to hypoxia response element (HRE) Cells were washed twice with ice-cold PBS and centrifuged at 250 g for 5 min. The cell pellet was washed in hypotonic buffer containing 10 mM HEPES (pH 7.9), 1.5 mM MgCl2, 10 mM KCl, 2 mM DTT, 2 mM sodium vandate, 4 μg/ml pepstatin, 4 μg/ml leupeptin, and 4 μg/ml aprotitnin, and incubated on ice for 15 min. Cells were homogenized by repeated passage through a 26G needle, and the nuclear fraction was obtained by centrifugation at 8000 g at 4 °C for 20 min. The pellet was suspended in an extraction buffer containing 20 mM HEPES (pH 7.9), 1.5 mM MgCl2, 0.42 M NaCl, 0.2 mM EDTA, 2 mM DTT, 1.0% Igepal CA-630, 25% glycerol, sodium vandate, 4 μg/ml pepstatin, 4 μg/ml leupeptin, and 4 μg/ml aprotitnin. Nuclear debris was removed by centrifugation at 16,000 g, 4 °C for 30 min, and the final nuclear extract was adjusted to 2 mg/ml with cold PBS. HIF-1α binding to HRE was measured with Trans AM™ HIF-1α kit [21] from Active Motif (Carlsbad, CA). Briefly, 5 μg/ml of nuclear extracts were added to a 96-well plate to which oligonucleotides containing an HRE (5´-TACGTGCT-3´) from the erythropoietin gene were immobilized. This oligonucleotide binds HIF-1α as a transcription factor with specific DNA binding activity. The plate was reacted with a primary antibody against HIF-1α followed by a secondary HRP-conjugated antibody, and spectrophotometry was performed at 450 nm using a reference wavelength of 655 nm.
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2.8. Inhibition of HIF-1α by RNA interference HIF-1α silencing was performed with 20–25 nucleotide targetspecific small interfering RNA (siRNA) designed to knock-down human HIF-1α expression (Santa Cruz Biotechnology, CA). Briefly, to a sterile tube was added 100 μl of transfection medium (Opti-MEM, Lonza, Basel, Switzerland) and 6 μl of lipofectamine™ 2000 (invitrogen, CA). HIF-1α inactivating siRNA was added to a final concentration of 100 nM and the mixture was incubated at room temperature for 45 min. The mixture was added to media of 60% confluent cells on 6-well plates that was seeded 24 h earlier. After vigorous agitation (for even siRNA dispersion), cells were incubated in a humidified atmosphere of 5% CO2 at 37 °C for 6 h. Cells were washed twice with transfection medium, fresh culture media were replaced, and hypoxia experiments were performed 2 days later.
Time course experiments in HUVE cells showed that 18F–FDG uptake began to rapidly rise after 4 h of hypoxia and reached a relative plateau between 6 and 16 h (Fig. 1A). This was accompanied by a similar temporal increase of lactate release by hypoxia that reached 198.0 ± 17.2% of controls by 24 h (Fig. 1B). These findings are consistent with a hypoxiainduced shift of metabolism toward glycolytic flux. When the two major determents of cellular glucose uptake were evaluated, hexokinase activity also displayed a temporal increase paralleling that of 18F–FDG uptake and lactate release. This reached 2.8 ± 0.1 fold of norrmoxic cells by 6 h of hypoxia (Fig. 1B). Plasma membrane Glut-1 expression was also increased by hypoxia, which was completely blocked by cycloheximide (Fig. 1B).
3.2. HIF-1α accumulates by hypoxia and is sufficient to enhance uptake
18
F–FDG
2.9. Data analysis Results are expressed as mean ± SE of data obtained from two separate experiments or mean ± SD of triplicate samples from a single representative experiment. Student's tests were used to compare 2 groups and one-way ANOVA with Tukey post-hoc tests were used to compare 3 or more groups. P b 0.05 was considered significant. 3. Results 3.1. Hypoxia stimulates endothelial cell hexokinase and glut-1
18
F–FDG uptake and glycolysis via
HUVE cells, HMVE cells and CPAE cells all showed significant augmentation of 18F–FDG uptake by 16 h hypoxia exposure to 178.8 ± 6.2%, 215.4 ± 7.6% and 179.7 ± 15.4% of normoxic cells, respectively (Fig. 1A).
When HUVE cells were exposed to 16 h of hypoxia, immunoblots displayed a substantial accumulation of HIF-1α protein to 325.6 ± 64.2% of controls (Fig. 2 A). In contrast, there was no increase of HIF2α protein (data not shown). Upon oxygen restoration after hypoxia, there was a prompt 32.3% decline of HIF-1α protein level by 4 h (Fig. 2A). However, in the presence of the HIF-1α stabilizing agent dimethyloxallylglycine (DMOG), a high HIF-1α level was maintained at 316.2 ± 100.4% of controls after 4 h of reoxygenation (Fig. 2A). Similar to HIF-1α level, 18F–FDG uptake that reached 236.2 ± 100.4% of controls by hypoxia returned to 133.9 ± 3.5% after 2 h of oxygen restoration (Fig. 2A). However, high 18F–FDG uptake was maintained in the presence of DMOG (Fig. 2A). DMOG also increased 18F– FDG uptake of normoxic cells to 124.0 ± 4.5% of untreated cells by 16 h (Fig. 2B). Taken together, these results indicate that HIF-1α plays a central role in mediating hypoxia-stimulated HUVE cell glycolysis.
Fig. 1. Effect of hypoxia on endothelial cell glucose metabolism. (A) Effect of 1% O2 on 18F–FDG uptake of endothelial cell lines (left), and time-course of hypoxia-stimulated 18F–FDG uptake in HUVE cells (right). (B) Time-course of hypoxia effects on HUVE cell lactate release (left) and hexokinase activity (middle), and effect of 24 h hypoxia with or without cycloheximde on membrane Glut-1 expression (right). Curves are fitted by non-linear regression. Results are mean ± SD of triplicate samples from a single experiment (A, left), or mean ± SE of values of six samples obtained from 2 independent experiments (time course data). †: p b 0.001, ‡ p b 0.0001 compared to normoxic controls.
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Fig. 2. Association between HIF1α accumulation and hypoxia-stimulated 18F–FDG uptake. (A) Effect of 16 h hypoxia and subsequent 4 h reoxygenation on HUVE cell HIF1α level (left), and time course of reversal of hypoxia-stimulated 18F–FDG uptake during reoxygenation with or without HIF1α stabilization by 50 μM DMOG (right). Data are mean ± SE of 6 samples from 2 separate experiments (HIF1α) or 9 samples from 3 separate experiments (18F–FDG uptake). (B) Stimulatory effect of DMOG on 18F–FDG uptake of normoxic HUVE cells. Data are mean ± SD of triplicate samples from a single experiment. *, p b 0.05; †, p b 0.001; ‡, p b 0.0001, compared to normoxic controls (A, left and B) or cells reoxygenated without DMOG (A, right).
3.3. HIF-1α inhibition or silencing completely abrogates hypoxia-stimulated 18 F–FDG uptake Accumulation of HIF-1α protein by hypoxia was completely blocked when HUVE cells were treated with a specific HIF-1α inhibitor or underwent gene silencing by transfection with specific siRNA (Fig. 3A). In both of these treatments, this was accompanied by complete
abrogation of the ability of hypoxia to augment 18F–FDG uptake, which decreased from 204.5 ± 13.4% to 115.7 ± 9.0% and 104.3 ± 2.6% of normoxic controls, respectively (Fig. 3B). Normoxic cell 18F–FDG uptake was not affected by the treatments. Also, transfection with control siRNA did not influence hypoxia-stimulated 18F–FDG uptake. Together, these results demonstrate that increased HIF-1α accumulation is necessary for the stimulatory effect of hypoxia on HUVE cell glycolysis.
Fig. 3. HIF1α is required for hypoxia to augment 18F–FDG uptake. (A) HUVE cell HIF1α accumulation stimulated by 24 h hypoxia is completely blocked by a specific inhibitor or by gene silencing with HIF1α specific siRNA. (B) Effects of treatment with an HIF1α inhibitor (left) or transfection with HIF1α specific or nonspecific siRNA (right) on normoxic and hypoxic HUVE cell 18F–FDG uptake. All data are mean ± SD of triplicate samples from a single experiment. **, p b 0.005; †, p b 0.001, compared to hypoxic cells without treatment.
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3.4. ROS plays a central role in hypoxia-stimulated HIF-1α accumulation and 18F–FDG uptake ROS and nitric oxide play important roles in angiogenesis as well as endothelial cell survival and metabolism. Measurements of these gases revealed that HUVE cell ROS production is markedly elevated by 1 h hypoxic exposure to 447.6 ± 117.2% of controls (Fig. 4A). On the other hand, whereas NO concentration markedly increased by adding sodium nitroprusside as donor (546.3 ± 15.5% of controls), there was no increase of nitric oxide production by hypoxia alone (Fig. 4A). HUVE cells were then evaluated for the effects of the ROS scavenger NAC and NADPH oxidase inhibitor apocynin. These treatments near completely blocked the increase of ROS production by hypoxia, whereas ROS production in normoxic cells were largely unaffected (Fig. 4B). Both of these treatments also completely abolished the capacity of hypoxia to stimulate 18F–FDG uptake (Fig. 4C) and HIF-1α accumulation (Fig. 4D). 3.5. Involvement of protein synthesis, and PKC and PI3K pathways Involvement of candidate intracellular signaling pathways in the metabolic effect of hypoxia was tested using specific inhibitors. Firstly, cycloheximide completely blocked hypoxia-stimulated 18F–FDG uptake and HIF-1α accumulation (Fig. 5A), indicating the requirement of both responses for new protein synthesis. Similarly, protein kinase C (PKC) inhibition with staurosporine completely blocked hypoxia-induced 18 F–FDG uptake and HIF-1α accumulation (Fig. 5B). In comparison, phosphatidylinositol 3-kinase (PI3K) inhibition with wortmannin effectively blocked the ability to augment 18F–FDG uptake, but not HIF-1α accumulation (222.1 ± 13.3% of controls; Fig. 5B). The requirement of PI3K activity for hypoxia-stimulated 18F–FDG uptake was further confirmed with PI3K specific siRNA that completely suppressed this response whereas nonspecific siRNA did not (Fig. 5C). Finally, HIF-1α binding to HRE in nuclear extracts from HUVE cells was enhanced to
2.51 ± 0.02 fold of controls by hypoxia, but this was significantly suppressed by wortmannin as well as by cycloheximide (Fig. 5C). This suggests that although PI3K activity is not required for hypoxia-induced HIF1α accumulation, it is necessary for HIF1α binding to HRE. 4. Discussion Tumor 18F–FDG uptake is directly and indirectly influenced by the metabolic state of vascular endothelial cells. Oxygen deprivation is a strong stimulus for new vessel formation, and endothelial cells respond by adjusting their metabolism to meet the demands required for survival and proliferation. Our study confirms that hypoxia shifts HUVE cell metabolism toward glycolytic flux. This metabolic effect was accompanied by upregulated Glut-1 expression and hexokinase activity. Furthermore, HIF1α accumulation driven by increased ROS production played a key role in the metabolic response. Stimulation of glucose uptake by low oxygen tension has previously been observed in HUVE and bovine endothelial cells [15], and in retinal capillary endothelial cells [16]. While HUVE cells are the most commonly studied endothelial cells, endothelial function and phenotype can be influenced by the vascular bed of origin. For instance, microvascular endothelial cells may respond differently with HUVE cells to cytokines [22]. However, our findings show that HUVE cells, HMVE cells and CPAE cells respond to hypoxia with similar elevations in FDG uptake. In our study, enhancement of 18F–FDG uptake was accompanied by a closely paralleled temporal rise in lactate release, indicating that the consumed glucose largely underwent glycolytic breakdown rather than oxidative phosphorylation. Previous reports attributed the augmented glucose uptake by hypoxia to glucose transporter upregulation [15,16]. Our results corroborated these findings by showing hypoxiainduced increase of plasma membrane-localized Glut-1, the predominant glucose transporter isoform in endothelial cells. Our results further showed that enhanced hexokinase activity also contributes to high 18F–
Fig. 4. Role of reactive oxygen species (ROS) on hypoxia-stimulated 18F–FDG uptake. (A) Hypoxia substantially increased HUVE cell production of ROS (left) but not nitric oxide (NO; right). Sodium nitroprusside (SNP; 100 μM) treated cells were used as a positive control for NO production. (B,C) Complete abrogation of hypoxia-stimulated ROS production (B) and 18F–FDG uptake (C) by N-acetyl cysteine (NAC; left) and apocynin (NAPDH oxidase inhibitor, right). (D) Effects of NAC and apocynin on hypoxia-induced HIF1α accumulation. Results are mean ± SE of values obtained from 2 separate experiments (n = 6; A-C), or mean ± SD of triplicate samples from a single experiment (D). †, p b 0.001; ‡, p b 0.0001, compared to normoxic controls (A) or hypoxic cells without treatment (D).
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Fig. 5. Inhibition of candidate signaling pathways on hypoxia effect. (A,B) Effects of 200 nM cycloheximide (CHX; A), 4 μM staurosporine (ST; B) and 200 nM wortmannin (WM; B) on hypoxia-stimulated HUVE cell 18F–FDG uptake (left) and HIF1α accumulation (right). (C) 18F–FDG uptake in normoxic and hypoxic HUVE cells gene-silenced with PI3K specific or control siRNA (left). Increased hypoxia-responsive element (HRE) binding of HIF1α extracted from HUVE cells after 24 h hypoxia is suppressed by WM or CHX (right). Results are mean ± SE of values of 9 samples from 3 separate experiments (FDG uptake in A and B), or mean ± SD of values of triplicate samples from a single experiment (C and HIF1α levels). †: p b 0.001; ‡ p b 0.0001 compared to hypoxic cells without treatment.
FDG uptake. Whereas glycolytic enzyme activation is considered a characteristic response of cancer cells to hypoxia, we are not aware of its description in endothelial cells. HIF1α is a transcriptional factor subunit that serves as a master regulator of oxygen homeostatsis [13]. Our results confirm that HUVE cells exposed to hypoxia buildup HIF1α protein level as they augment 18F– FDG uptake. The two responses were further linked in that both responses were revered when normal oxygen tension was restored after hypoxic treatment. Furthermore, stabilizing HIF1α protein by retarding its degradation during reoxygenation with the prolyl hydroxylase inhibitor, DMOG, was sufficient to maintain a high level of 18F–FDG uptake. In addition, normoxic cells also showed modest but significant elevation 18 F–FDG uptake by DMOG treatment alone. Importantly, blocking of HIF1α accumulation by treatment with a specific HIF1α inhibitor or gene silencing with specific siRNA completely abolished the ability of hypoxia to augment 18F–FDG uptake. Collectively, these findings demonstrate a key role played by HIF1α accumulation that is both necessary and sufficient for hypoxia to promote HUVE cell glycolysis and 18 F–FDG uptake. Tightly regulated generation of ROS and nitrogen species that can fine tune downstream signaling in cell-specific manners are critical constituents of cellular responses to various stimuli [23]. ROS production regulates several aspects of vascular and endothelial function including angiogenesis [23]. A previous study showed that endothelial cells increase ROS production under hypoxic conditions via the mitochondrial respiratory chain or NADPH oxidase activation [24]. In our study, the ROS scavenger, NAC, totally suppressed ROS production and HIF1α accumulation as well as 18F–FDG uptake by hypoxia. Furthermore, NADPH oxidase-mediated ROS production was a major effector for the metabolic effect of hypoxia on HUVE cells, since both HIF1α accumulation and 18F–FDG uptake responses were completely blocked by the NADPH oxidase inhibitor, apocynin. These findings put ROS production
at an early event upstream of HIF1α accumulation in the metabolic response to hypoxia. This notion is consistent with previous proposals that ROS generation may be an initial step for hypoxia-sensing in endothelial cells [25]. We also tested the possibility of NO involvement, since hypoxia has been shown to stimulate NO release in certain endothelial cells [26] and our group previously demonstrated NO to be a powerful simulator of HUVE cell 18F–FDG uptake [4]. However, the results of the present study showed that hypoxia does not increase NO release, excluding the involvement of NO signaling in the metabolic effect of hypoxia on HUVE cells. HIF1α accumulation can occur either through slowing of its proteosomal degradation or by increased synthesis through growth factor signaling [27]. The role of protein synthesis on a cell response can be tested by treatment with cycloheximide. In our study, this caused a total loss of hypoxia-stimulated HIF1α accumulation and 18F–FDG uptake. Similar losses of hypoxia-induced HIF1α buildup [28] and deoxyglucose uptake by cycloheximide have been previously observed in other cell types [29]. These results suggest that new protein synthesis is required to mediate the metabolic effect of hypoxia. HIF1α is rapidly destroyed in normoxic tumor cells with a half-life of approximately 5 min, whereas stabilization in hypoxic states will prolong its half-life to 2–4 h [10]. In HUVE cells, HIF1α half-life may be shorter, even under hypoxic conditions (about 30 min) [30]. Our data imply that hypoxic HUVE cells require continuous protein synthesis for HIF1α accumulation and increasing 18F–FDG uptake. On exploring candidate signaling pathways that may be involved, the potent PKC inhibitor staurosporine completely blocked the ability of hypoxia to stimulate both HIF1α accumulation and 18F–FDG uptake. This finding could be related to the fact that the PKC pathway can be activated by hypoxia [31] and is implicated in the angiogenic response [32]. The specific PI3K inhibitor wortmannin also inhibited stimulation of 18F–FDG uptake by hypoxia, whereas it had no influence on HIF1α
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Fig. 6. Scheme of proposed molecular mechanism for hypoxia-stimulated endothelial cell glycolysis.
accumulation. The PI3K pathway is activated by growth factors and is closely involved in glucose metabolism as well as angiogenesis. A previous study reported that certain HRE promoters bind HIF1α in a PI3Kdependent manner [33]. Hence, it is possible that PI3K is required for HIF1α binding to HRE and subsequent stimulation of glycolysis, even though it is not necessary for HIF1α accumulation. A schematic of the proposed molecular mechanism involved in hypoxia-stimulated endothelial cell glycolysis and 18F–FDG uptake is illustrated in Fig. 6. In conclusion, hypoxia stimulates HUVE cell glycolysis, in a manner accompanied by upregulated Glut-1 expression and hexokinase activity. HIF1α accumulation driven by increased ROS generation plays a central role in this metabolic effect, which also involves PKC and PI3K signaling. The resultant increase of endothelial cell 18F–FDG uptake may contribute to greater transport of the substrate to underlying cancer cells, which could together contribute to higher tumor 18F–FDG uptake in oxygen-deprived regions. Sources of funding This work was supported by the Basic Science Research Program through the National Research Foundation of Korea (NRF) funded by the Ministry of Science, ICT and Future Planning (NRF-2015R1A2A2A01006419). Disclosures None. References [1] Warburg O. On the origin of cancer cells. Science 1956;123:309–14. [2] Farwell MD, Pryma DA, Mankoff DA. PET/CT imaging in cancer: current applications and future directions. Cancer 2014;120:3433–45. [3] Maschauer S, Prante O, Hoffmann M, Deichen JT, Kuwert T. Characterization of FDG uptake in human endothelial cells in vitro. J Nucl Med 2004;45:455–60. [4] Paik JY, Lee KH, Ko BH, Choe YS, Choi Y, Kim BT. Nitric oxide stimulates FDG uptake in human endothelial cells through increased hexokinase activity and GLUT1 expression. J Nucl Med 2005;46:365–70. [5] Paik JY, Koh BH, Jung KH, Lee KH. Fibronectin stimulates endothelial cell FDG uptake via focal adhesion kinase mediated phosphatidylinositol 3-kinase/Akt signaling. J Nucl Med 2009;50:618–24.
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