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Exp Neurobiol 2017; 26(5): 295-306
Published online October 31, 2017
© The Korean Society for Brain and Neural Sciences
Jiwon Esther Han1, Pyung Won Lim1, Chul Min Na1, You Sik Choi1, Joo Young Lee1, Yona Kim1, Hyung Woo Park1, Hyo Eun Moon1, Man Seung Heo2, Hye Ran Park1, Dong Gyu Kim1 and Sun Ha Paek1,3,4*
1Department of Neurosurgery, Seoul National University College of Medicine, Seoul 03082, 2Smart Healthcare Medical Device Research Center, Samsung Medical Center, Seoul 06351, 3Cancer Research Institute, Seoul National University College of Medicine, Seoul 03082, 4Hypoxia Ischemia Disease Institute, Seoul National University College of Medicine, Seoul 03082, Korea
Correspondence to: *To whom correspondence should be addressed.
TEL: 82-2-744-8459, FAX: 82-2-2072-3993
Glioblastoma multiforme (GBM) is the most common and aggressive form of brain tumors. GBMs, like other tumors, rely relatively less on mitochondrial oxidative phosphorylation (OXPHOS) and utilize more aerobic glycolysis, and this metabolic shift becomes augmented under hypoxia. In the present study, we investigated the physiological significance of altered glucose metabolism and hypoxic adaptation in the GBM cell line U251 and two newly established primary GBMs (GBM28 and GBM37). We found that these three GBMs exhibited differential growth rates under hypoxia compared to those under normoxia. Under normoxia, the basal expressions of HIF1α and the glycolysis-associated genes, PDK1, PDK3, and GLUT1, were relatively low in U251 and GBM28, while their basal expressions were high in GBM37. Under hypoxia, the expressions of these genes were enhanced further in all three GBMs. Treatment with dichloroacetate (DCA), an inhibitor of pyruvate dehydrogenase kinase (PDK), induced cell death in GBM28 and GBM37 maintained under normoxia, whereas DCA effects disappeared under hypoxia, suggesting that hypoxic adaptation dominated DCA effects in these GBMs. In contrast, the inhibition of HIF1α with chrysin suppressed the expression of PDK1, PDK3, and GLUT1 and markedly promoted cell death of all GBMs under both normoxia and hypoxia. Interestingly, however, GBMs treated with chrysin under hypoxia still sustained higher viability than those under normoxia, and chrysin and DCA co-treatment was unable to eliminate this hypoxia-dependent resistance. Together, these results suggest that hypoxic adaptation is critical for maintaining viability of GBMs, and targeting hypoxic adaptation can be an important treatment option for GBMs.
Keywords: Glioblastoma, HIF1alpha, hypoxia
Glioblastoma multiforme (GBM) is the most frequent and aggressive subtype of primary brain tumors [1,2]. Current standard of care for GBMs includes the surgical resection of the primary tumor followed by radiotherapy and chemotherapy [3,4]. In spite of these multimodal therapeutic interventions, the overall survival period of GBM patients is, on average, only 12~15 months due to tumor recurrence . In fact, recurrent GBMs are repopulated with more malignant cells that are resistant to radio- and chemotherapy . Thus, more efficient therapeutic strategies targeting cellular features of GBMs should be developed.
GBMs, like other tumors, utilize a reprogrammed energy metabolism employing reduced mitochondrial oxidative phosphorylation (OXPHOS) and concomitantly increased aerobic glycolysis [7,8]. Although this metabolic shift, known as the Warburg Effect, is less efficient for generating ATP per unit glucose than mitochondrial OXPHOS, it is adopted by GBMs presumably because it rapidly supplies biosynthetic precursors, circumvents excess production of reactive oxygen species (ROS) in mitochondria, and decreases apoptotic risks of tumor cells . In addition, increased aerobic glycolysis provides acidic extracellular microenvironment, giving a survival advantage to tumor cells over normal cells .
Numerous investigations have been carried out to intervene increased aerobic glycolysis in tumor cells [7,8,11,12]. One such strategy is the use of dichloroacetate (DCA), a pyruvate mimetic that inhibits pyruvate dehydrogenase kinase (PDK), which is a negative regulator of pyruvate dehydrogenase (PDH) and blocks the conversions of pyruvate into acetyl-CoA . In fact, the expression levels of PDK isoforms are elevated in various types of tumors, and the administration of DCA offers improved prognosis of certain GBMs and other tumors by reducing aerobic glycolysis [12,14,15,16,17]. Although increased aerobic glycolysis is an important feature of GBMs, studying glycolytic adaptation only in normoxic condition (20% oxygen) might be insufficient to fully understand the actual physiological responses influencing tumorigenicity.
Hypoxia is a well-known feature of tumor microenvironment . GBMs, like other tumor cells, grow well in hypoxic condition by employing increased expression of hypoxia-inducible factor 1α (HIF1α) [19,20]. Indeed, HIF1α, together with its partner HIF1β, forms a complex which upregulates expression of almost all enzymes in glycolysis . HIF1α/β also upregulates the expression of genes involved in angiogenesis, survival, and migration, resulting in enhanced metastatic potential and resistance to radiotherapy and chemotherapy [21,22,23,24]. Since hypoxic adaptation is important for tumorigenicity , an effective strategy for treating GBM needs to be studied under hypoxia .
In the present study, we investigated the physiological significance of aerobic glycolytic adaptation and hypoxic adaptation with respect to cell viability of U251, a well-established GBM cell line, and the two primary GBMs, GBM28 (initial tumor) and GBM37 (recurrent tumor), which were derived from the same patient. Cellular responses were examined under hypoxia (approximately 1% oxygen) that recapitulates low oxygen levels of
The human glioblastoma cell line U251 and two primary glioblastoma cells (GBM28 and GBM37) were used in this study. The human glioblastoma U251 was purchased from the American Type Culture Collection (ATCC; Manassas, VA, USA). The primary glioblastoma cells, GBM28 and GBM37, were established from the same human patient who received a standard GBM treatment consisting of concurrent radiotherapy and temozolomide (TMZ) treatment followed by adjuvant TMZ as a primary treatment. GBM28 and GBM37 represented initial and recurrent tumors, respectively. The neural stem cell, NSC10, was used as normal control cell and described previously . These GBM cell line and primary cells were cultured with Dulbecco's modified Eagle's medium containing 10% fetal bovine serum (FBS, Gibco Corp., Grand Island, NY, USA) and 1% antibiotics/antimycotics in a 5% CO2 incubator at 37℃. Cells were routinely passaged by trypsinization using TrypLE™ (Gibco Corp., Grand Island, NY, USA) every 3~4 days. For the hypoxia treatments, O2 levels were maintained at 1%, with the residual gas mixture being 94% N2 and 5% CO2, as compared to the normoxia experiments, in which O2 levels were maintained at 20%. The experimental protocol for handling human subjects was reviewed and approved by the Institutional Review Board of Seoul National University Hospital (IRB No. H-1009-025-331).
Each of GBM cells was seeded on a 96-well plate at a density of 3×103 cells per well and maintained in a 5% CO2 incubator with 90% relative humidity at 37℃ for 24 h to ensure cell attachment. Subsequently, cells were put in a hypoxia incubator (1% O2) for 72 h. The control groups were prepared in a normoxia incubator (20% O2). In every 24 h for 3 consecutive days, the effects of hypoxic and normoxic conditions on cell proliferation were evaluated using the Cell Counting Kit-8 (CCK; Dojindo Molecular Technologies, Kumamoto, Japan). The 10 µl of the CCK solution was added per well and incubated for 2 h. Then, cell proliferation was analyzed according to the manufacturer's instructions by measuring the absorbance of the reaction solution at 450 nm with a Multiscan MS spectrophotometer (Labsystems, Stockholm, Sweden). All experiments were conducted in triplicate, and these were repeated 2~3 times.
GBM cells were lysed and total RNA was isolated using the RNeasy Mini Kit (Qiagen, Valencia, CA, USA) in accordance to the manufacturer's protocol. Then, cDNA was synthesized from 3 µg of the total RNA extracted using the Superscript III cDNA Synthesis Kit (Invitrogen, Carlsbad, CA, USA). A reverse transcriptase polymerase chain reaction (RT-PCR) was performed and the resulting PCR products were resolved by 1.5% agarose gel electrophoresis. Expression levels of transcripts were quantified using ImageJ (NIH, Bethesda, MD, USA). The primer sequences used in this study were as follows: 5′-GAAAGCGCAAGTCCTCAAAG-3′ and 5′-TGGGTAGGAGATGGAGATGC-3′ for HIF1α; 5′-CGGATCAGAAACCGACACA-3′ and 5′-ACTGAACATTCTGGCTGGTGA-3′ for PDK1; 5′-TTAATAAGTCCGCATGGCGC-3′ and 5′-TGAAGCATCCCTGGGTTCAC-3′ for PDK3; 5′-GATTGGCTCCTTCTCTGTGG-3′ and 5′-TCAAAGGACTTGCCCAGTTT-3′ for GLUT1; 5′-GTACCATGCGGAGACCATCA-3′ and 5′-GTAGGCGTTATCCAGCGTGA-3′ for PKM2; 5′-GCACTCTTCCAGCCTTCCTTCC-3′ and 5′-GAGCCGCCGATCCACACG-3′ for β-actin.
Cell viability of GBMs was evaluated with the
Mitochondria were isolated as described previously with minor modifications . Cells were harvested by centrifugation at 1,500 g for 5 min. After decanting the supernatant, cells were resuspended in 10 mM of ice-cold Tris buffer (pH 7.6) containing protease inhibitor cocktail (Roche, Mannheim, Germany). Cells were then disrupted mechanically by passing through a 1 ml syringe with a 21-gauge needle 20 times and chemically by the addition of 1.5 M ice-cold sucrose. Subsequently, the disrupted cells were centrifuged at 700 g for 10 min at 2℃. The supernatant containing mitochondria was collected and recentrifuged at 14,000 g for 10 min at 2℃. The resulting pellet was washed and resuspended in 100 µl of 10 mM ice-cold Tris buffer (pH 7.6) containing protease inhibitor cocktail. The 10 µl of isolated mitochondria was taken and lysed for the quantification of protein concentration, and the remaining volumes were stored on ice until functional analysis of mitochondrial complex activities.
Mitochondrial complex I, II, and IV activity assays were carried out as described previously with minor modifications [27,28,29]. For the complex I activity assay, 1 µg of the mitochondrial fraction was preincubated for 3 min at 37℃ in 240 µl of 25 mM potassium phosphate buffer (pH 7.8) containing 1.0 µM antimycin-A, 3.5 g/L BSA, 70 µM decylubiquinone, and 60 µM DCIP. The reaction was started by the addition of 5 µl of 160 mM NADH. The complex activity was measured spectrophotometrically at 600 nm at 30 sec intervals for 5 min at 37℃. After 5 min, 2.5 µl of rotenone in dimethyl sulfoxide was added to the reaction mixture and the absorbance was measured at 600 nm for additional 5 min at 30 sec intervals at 37℃ for the control groups.
For the complex II activity assay, 1 µg of the mitochondrial fraction was preincubated for 10 min at 37℃ in 240 µl of 80 mM potassium phosphate buffer (pH 7.8) containing 1 µM antimycin-A, 1 g/l BSA, 50 µM decylubiquinone, 60 µM DCIP, 3 µM rotenone, 2 mM EDTA, and 0.2 mM ATP. The reaction was started by the addition of 20 µl of 1.5 M succinate and 0.75 µl of 0.1 M KCN, and the activity was assessed spectrophotometrically at 600 nm at 30 sec intervals for 5 min at 37℃. For the control groups, 5 mM malonate was added to the reaction mixture prior to preincubation along with the mitochondrial sample.
For the complex IV activity assay, 1 µg of the mitochondrial fraction was preincubated for 10 min at 30℃ in 240 µl of 30 mM potassium phosphate buffer (pH 7.4) containing 2.5 mM dodecylmaltoside. The reaction was started by the addition of 20 µl of 34 µM ferrocytochrome c. The complex activity was measured spectrophotometrically at 550 nm at 30 sec intervals for 5 min at 30℃. For the control groups, 1 mM KCN was added to the reaction mixture prior to preincubation along with the mitochondrial sample.
The activity of citrate synthase, a tricarboxylic acid (TCA) cycle enzyme, was used to quantify mitochondrial mass and accordingly normalize the activity of mitochondrial complex enzymes as described previously . For the citrate synthase activity assay, 1 µg of the mitochondrial fraction was preincubated for 5 min at 30℃ in 240 µl of 50 mM Tris–HCl buffer (pH 7.5) containing 0.2 mM 5,5′-dithiobis-(2-nitrobenzoic acid) and 0.1 mM acetyl-CoA. The reaction was started by the addition of 2.5 µl of 50 mM oxaloacetate. The activity was measured spectrophotometrically at 412 nm at 30 sec intervals for 5 min at 30℃. For control groups, an equivalent volume of water was added instead of oxaloacetate.
Data are expressed as means±S.E.M. Two-sample comparisons were carried out using an unpaired two-tailed Student's
Since hypoxic adaptation is a prominent feature of GBMs [30,31,32], we first examined the proliferation rates of the GBM cell line U251 and two primary GBMs (GBM28, GBM37) under normoxia (20% O2) and hypoxia (1% O2). GBM28 and GBM37 are initial and recurrent tumors, respectively, generated from the same patient. Overall, GBM28 and GBM37 exhibited lower proliferation rates than U251 (Fig. 1A~C). U251 showed significantly higher proliferation rate under hypoxia than normoxia (Fig. 1A). GBM28 showed no difference in growth rates between normoxic and hypoxic conditions (Fig. 1B), which was in contrast to GBM37 exhibiting significantly reduced growth rate under hypoxia compared to that under normoxia (Fig. 1C).
Next, we examined the expression level of HIF1α in different GBMs under normoxic and hypoxic conditions. The basal expression level of HIF1α in U251 and GBM28 under normoxia was relatively low, while it was high in GBM37 (Fig. 1D and E). Under hypoxia, the transcript level of HIF1α markedly increased in both U251 and GBM28, whereas it was retained high in GBM37 (Fig. 1D and E). These results indicated that there is a correlation between the increase of proliferation rate and the induction of HIF1α under hypoxia (Fig.1A~E). Then, the expression levels of glycolysis-associated HIF1α target genes were examined. Under normoxia, the basal expression levels of PDK1, PDK3, and GLUT1 were relatively low in U251 and GBM28, while their basal expression levels were high in GBM37 (Fig. 1F~I). Under hypoxia, the expression levels of PDK1, PDK3, and GLUT1 were enhanced further in all GBMs examined, albeit at different levels. The expression levels of PKM2 varied in different GBMs under normoxia and were not altered under hypoxia (Fig. 1F and J). Considering that the overexpression of HIF1α and glycolysis-associated genes has been continuously reported to be correlated with a worse patient outcome [22,23], greater expression of HIF1α, PDK1, PDK3, and GLUT1 in GBM37 than U251 and GBM28 under normoxia might be a predictable feature of the recurrent tumor.
The intervention of glycolytic adaptation with DCA, an inhibitor of PDK, has been reported to offer improved therapeutic outcomes in various tumor types including breast, oral, lung, ovarian, and colon cancers [33,34,35,36,37]. Therefore, we examined the effects of DCA on viabilities of GBM cells cultured under normoxia or hypoxia. The results of MTT assay obtained after 48 h of DCA treatment indicated that no cell death was induced in U251 at all doses of DCA (5, 10, or 20 mM) under both normoxia and hypoxia (Fig. 2A). Interestingly, in GBM28 and GBM37, DCA treatment at 5, 10, or 20 mM induced significant levels of cell death under normoxia, but DCA at all doses failed to induce cell death under hypoxia (Fig. 2B and C), implying that hypoxic adaptation prevailed DCA effects in these cells. It is notable that DCA treatment in neural stem cells (NSC10) at the dose of 5 or 10 mM did not produce cell death under both normoxic and hypoxic conditions (Fig. 2D), although DCA treatment at 20 mM induced cell death under hypoxia. Overall, these results indicated that DCA effects on inducing cell death under normoxia varied, from no effect in U251 to moderate effects in GBM28 and GBM37, whereas all three GBMs were resistant to DCA treatment under hypoxia.
The results of DCA-induced cell death led us to examine the levels of mitochondrial OXPHOS activities in GBMs. The levels of mitochondrial complex I and II activities in U251 were, respectively, 77.1 and 83.5% of the control NSC10, and those in GBM28 and GBM37 were in the range of 67.4 and 85.1% of NSC10 (Fig. 2E and F). The level of mitochondrial complex IV activity in U251 was 60.8% of NSC10, whereas those in GBM28 and GBM37 were, respectively, 24.9 and 43.2% of NSC10 (Fig. 2G). These results indicated that the recurrent tumor GBM37 had relatively lower complex I and higher complex IV activities than the initial tumor GBM28 (Fig. 2E and G), and a similar remodeling of mitochondrial complex activities has been reported in a previous report studying TMZ-mediated chemoresistant gliomas . Taken together, these results suggest that compared to NSC10, GBMs had the reduced utilization of OXPHOS pathway and the extent of reduction was prominent for complex IV activity.
Next, we examined the physiological significance of hypoxic adaptation in cell viability of GBMs. Treatment of GBMs with chrysin, a naturally occurring flavonoid with an inhibitory activity of HIF1α , induced significant levels of cell death in all GBMs under both normoxia and hypoxia (Fig. 3A~C). Although chrysin effects were produced in a dose-dependent manner in all cell types, cells maintained under hypoxia had relatively higher viability than those cultured under normoxia, indicating the continued presence of hypoxia-dependent resistance (Fig. 3A~C). Chrysin treatment suppressed the expressions of HIF1α and the glycolysis-associated genes, PDK1, PDK3, and GLUT1 (Fig. 3E~I). However, the hypoxia-dependent increases of these genes partially remained in all cell types (Fig. 3E~I), which was consistent with chrysin-dependent partial reduction of cell viability of GBMs under hypoxia (Fig. 3A~C). In NSC10, chrysin treatment at 10 µM did not induce cell death under normoxic and hypoxic conditions, whereas chrysin at 50 and 100 µM induced cell death under both normoxic and hypoxic conditions (Fig. 3D). Overall, these results suggest that chrysin induced cell death of all GBMs examined via suppressing the expressions of hypoxia- and glycolysis-associated genes. However, chrysin effects were still weak under hypoxia than those under normoxia, and hypoxia-dependent resistance was sustained in all cell types.
Next, we examined whether blocking both hypoxic adaptation and glycolytic adaptation lessened hypoxia-dependent resistance of GBMs. The levels of cell death induced by chrysin at 10 µM increased in all GBMs when DCA at 10 or 20 mM was co-treated (Fig. 4A~C), including U251 which did not respond to DCA alone (Fig. 2A). However, to our surprise, co-treatment of chrysin (10 µM) and DCA (10 or 20 mM) reduced cell viability of NSC10 as well (Fig. 4D), whereas chrysin at 10 µM alone was not toxic to NSC10 (Fig. 3D). When a higher dose of chrysin (100 µM) was cotreated with DCA (10 or 20 mM), cell viability of all GBMs as well as NSC10 decreased compared to that induced by chrysin (100 µM) alone (Fig. 4E~H). Overall, these results suggest that chrysin and DCA co-treatment produced additive effects on inducing cell death of GBMs. Interestingly however, co-treatment even at high doses of chrysin and DCA was not potent enough to reduce cell viability of GBMs under hypoxia to the levels achieved under normoxia.
Mounting evidence has suggested that tumor cells utilize more aerobic glycolysis while using less mitochondrial OXPHOS activity [40,41]. This metabolic shift becomes enhanced under hypoxia in various tumor cells [21,42]. Indeed, all GBM cells had reduced oxidative glucose metabolism compared to NSC10, and, in particular, the newly established primary GBMs (GBM28 and GBM37) had significantly reduced mitochondrial complex IV activity compared to NSC10 (Fig. 2E~G). The mitochondrial complex I is the major step where mitochondrial ROS production takes place, and as long as the complex I activity remains low, tumor cells can evade ROS stress resulted from mitochondria [43,44]. On the other hand, the complex IV is the terminal step of the electron transport chain (ETC), and therefore, the complex IV activity represents the overall ETC activity [38,45]. The complex I activity in GBM37 tended to be lower than that in GBM28, while the complex IV activity in GBM37 was higher than that in GBM28 (Fig. 2E and G). The reduced demand for aerobic glycolysis in the recurrent tumor GBM37 compared to that in the initial tumor GBM28 might be indicative of reprogrammed tumorigenicity of GBM37 wherein its overall ETC activity in mitochondria was increased [38,45]. The reduced growth rate of GBM37 under hypoxia (Fig. 1C) might also be related in part to reprogrammed OXPHOS activity. We speculate that GBM37 can maintain its tumorigenicity presumably by regulating both glycolytic and reprogrammed oxidative metabolisms to fit its needs.
PDKs suppress PDH, a mitochondrial enzyme that converts pyruvate into acetyl-CoA to provide the initial substrate for the TCA cycle . U251 was resistant to DCA treatment under normoxia (Fig. 2A), probably because U251 utilized a relatively high level of mitochondrial OXPHOS pathway (Fig. 2E~G). However, as demonstrated in the present study, DCA treatment in GBM28 and GBM37 induced cell death under normoxia, and this DCA effect agrees well with previous reports showing cytotoxic effects of DCA in various tumors including GBMs [8,34,47]. DCA-induced cell death of GBM28 under normoxia was slightly higher than that of GBM37 (Fig. 2B and C), which might be correlated with significantly lower expressions of glycolysis-associated genes in GBM28 compared to GBM37 under normoxia (Fig. 1F~J). However, DCA effects on inducing cell death of GBM28 and GBM37 completely disappeared when these cells were cultured under hypoxia, which is considered more physiologically relevant tumor microenvironment than normoxia . These results are consistent in part with the increased expression levels of HIF1α and the glycolysis regulating genes, PDK1, PDK3, and GLUT1 in these GBMs under hypoxia (Fig. 1D~I). A similar induction pattern of these genes was detected in U251 maintained under hypoxia, which might contribute to the resistance of this cell line to DCA treatment. Overall, these results suggest that hypoxic adaptation dominates glycolytic adaptation in modulating survival of GBMs, and thus targeting hypoxic adaptation might be more important than intervening glycolytic adaption for treating GBM.
Hypoxia facilitates the metabolic adaptation of cancer cells via HIF1α-dependent regulation of glycolytic genes , and the up-regulation of HIF1α is associated with radio- and chemoresistance in GBMs [48,49]. Our cell viability assay indicated that chrysin treatment produced substantial cell death in all GBMs under normoxia (Fig. 3A~C). Upon chrysin treatment, U251, which was resistant to DCA treatment under normoxia (Fig. 2A), showed cell death to the levels of GBM28 (Fig. 3A and B). Furthermore, chrysin treatment produced cell death in all three GBMs under hypoxia, whereas DCA-induced cell death of these GBMs was absent under hypoxia. These results suggest that chrysin targeted the critical mechanism that controls the viability of GBMs. Similar to the case of DCA-induced cell death, however, GBM37 was still slightly more resistant to chrysin than GBM28 (Fig. 3B and C). For instance, 10 µM of chrysin induced cell death in GBM28 under normoxia but not in GBM37. Chrysin-dependent regulation of HIF1α expression is insufficient to explain the different responses to DCA and chrysin between the initial and recurrent tumors. In addition, considering that chrysin-treated GBM cells under hypoxia still displayed higher viability than those under normoxia, we speculate that chrysin was not potent enough to reduce hypoxia-dependent resistance of GBMs.
Cell death induced by chrysin was further increased when DCA was co-treated (Fig. 4). Co-treatment of chrysin and DCA at low doses (e.g. 10 µM for chrysin and 10 mM for DCA) induced cell death in GBMs under both normoxic and hypoxic conditions, but it is slightly toxic to NSC10 as well (Fig. 4A~D). Chrysin and DCA co-treatment at higher doses also induced cell death in GBMs and NSC10 (Fig. 4E~H). However, it is important to note that hypoxic condition stimulated GBMs to be resistant to both chrysin and DCA treatment, and the co-treatment even with high doses of chrysin and DCA was unable to lessen hypoxia-dependent resilience of GBMs (Fig. 5). Considering these complex results, further detailed studies regarding the mechanisms underlying the resilient response of GBMs to the two treatments under hypoxia should be addressed.
In summary, the inhibition of increased aerobic glycolysis, a metabolic feature of various tumors including GBMs, by DCA produced cell death in GBM28 and GBM37, but not in U251. The inhibition of HIF1α with chrysin, however, pulled down the viability of all three GBMs examined, supporting the possibility that hypoxic adaptation involving HIF1α induction is critical for maintaining cell viability of GBMs. Interestingly, the results of the present study indicated that cell viability of GBMs was still higher under hypoxia than normoxia. Moreover, modulating both glycolytic and hypoxic adaptations by DCA and chrysin, respectively, was ineffective in eliminating this hypoxia-dependent resistance of GBMs, although the co-treatment brought additive effects in inducing cell death. Further extensive studies are necessary to fully elaborate the detailed mechanisms underlying the differential responses of GBMs to DCA and/or chrysin under normoxia and hypoxia.