Changes in metabolic processes play a critical role in the survival or death of cells subjected to various stresses. In the present study, we have investigated the effects of ER (endoplasmic reticulum) stress on cellular metabolism. A major difficulty in studying metabolic responses to ER stress is that ER stress normally leads to apoptosis and metabolic changes observed in dying cells may be misleading. Therefore we have used IL-3 (interleukin 3)-dependent Bak−/−Bax−/− haemopoietic cells which do not die in the presence of the ER-stress-inducing drug tunicamycin. Tunicamycin-treated Bak−/−Bax−/− cells remain viable, but cease growth, arresting in G1-phase and undergoing autophagy in the absence of apoptosis. In these cells, we used NMR-based SIRM (stable isotope-resolved metabolomics) to determine the metabolic effects of tunicamycin. Glucose was found to be the major carbon source for energy production and anabolic metabolism. Following tunicamycin exposure, glucose uptake and lactate production are greatly reduced. Decreased 13C labelling in several cellular metabolites suggests that mitochondrial function in cells undergoing ER stress is compromised. Consistent with this, mitochondrial membrane potential, oxygen consumption and cellular ATP levels are much lower compared with untreated cells. Importantly, the effects of tunicamycin on cellular metabolic processes may be related to a reduction in cell-surface GLUT1 (glucose transporter 1) levels which, in turn, may reflect decreased Akt signalling. These results suggest that ER stress exerts profound effects on several central metabolic processes which may help to explain cell death arising from ER stress in normal cells.
The ER (endoplasmic reticulum) is a vital organelle that plays an important role in the regulation of cellular homoeostasis and communication [1,2]. The ER is the cellular organelle where proteins and lipids are synthesized and modified. Many protein chaperones in the ER facilitate the proper folding of individual proteins and the formation of macromolecular complexes. The disruption of ER functions by depletion of ER Ca2+ stores, inhibition of asparagine-linked (N-) protein glycosylation, disturbance of disulfide bond formation or viral infection leads to protein misfolding and subsequent protein aggregation. The accumulation of unfolded proteins subsequently induces an ER stress response via the UPR (unfolded protein response) pathway. In mammalian cells, UPR signalling involves three pathways: IRE1 (inositol-requiring kinase-1), ATF6 (activating transcription factor 6) and PERK (RNA-dependent protein kinase-like ER kinase) . These sense the presence of unfolded proteins in the ER lumen and trigger various downstream signalling events. In response to different levels of UPR, there may be various types of cellular responses, including apoptosis and autophagy.
Whereas apoptosis is a tightly regulated cellular suicide programme, autophagy is a cellular process for degradation of cellular components in lysosomes and is activated by a number of stressors including the UPR [3–6]. During autophagy, double-membraned vesicles, autophagosomes, are formed de novo to sequester cytoplasmic contents. Once the outer membranes of autophagosomes have fused with lysosomal membranes, the cytoplasmic contents are delivered to the lysosome lumen, where they are degraded. The resulting degradation products are released into the cytosol and may be reutilized. Autophagy is a highly regulated cellular catabolism system and deficiency in autophagy has been invoked in the pathogenesis of many human diseases, including neurodegeneration, infections and cancer. ER stress has been reported to induce autophagy in many cellular systems and may represent a defence mechanism which promotes cell survival . However, more extreme ER stress can lead to autophagic cell death. Although it is not clear how pro-survival and pro-death outcomes of autophagy are regulated, it appears that the extent of autophagy may determine cell fate .
Cells undergoing autophagy typically exit the cell cycle and maintain a minimal metabolic rate commensurate with maintenance of cellular homoeostasis and repair. A large fraction of ATP consumed is used for maintaining ion gradients across the plasma membrane and intracellular membranes, and for protein synthesis [9,10]. An important issue for cell survival is the production of sufficient metabolic energy for repair and membrane potential maintenance. How metabolic changes in ER stress-induced cellular metabolism are involved in cell-fate decisions is largely unknown. In the present study, we examined the metabolic effects of ER stress on IL-3 (interleukin 3)-dependent Bak−/−Bax−/− cells using a NMR-based SIRM (stable isotope-resolved metabolomics) approach. We found that ER stress induces progressive autophagy and a relative inability to utilize extracellular glucose, resulting in reduced glycolysis and tricarboxylic acid cycle activity. This appears to be accompanied by a reduction in GLUT1 (glucose transporter 1) levels on the cell surface. Together, these data suggest that ER stress has marked effects on central metabolic processes, particularly glucose metabolism.
MATERIALS AND METHODS
Cell lines and reagents
Bak−/−Bax−/− IL-3-dependent cells were cultured at 37 °C (95% air/5% CO2) in glucose-free RPMI 1640 medium (Sigma) supplemented with 10% (v/v) dialysed FBS (fetal bovine serum) (Clontech), 5 mM glucose (Sigma), 2 mM glutamine (Mediatech), 100 units/ml penicillin (Mediatech), 100 μg/ml streptomycin (Mediatech) and 3.4 ng/ml IL-3 (Invitrogen). Wild-type murine Bax or Bak cDNA was re-expressed in IL-3-dependent Bak−/−Bax−/− cells by retroviral infection, and stable clones expressing Bax or Bak were selected as described previously . cDNAs of Myc-tagged mouse GLUT1 or mouse Akt1 with the myristoylation sequence GSSKSKPKSR at its N-terminus were retrovirally expressed in Bak−/−Bax−/− IL-3-dependent cells with GFP (green fluorescent protein) as a marker expressed from an IRES (internal ribosome entry site) as described previously . Cells stably expressing Myc-tagged GLUT1 or myristoylated Akt1 were obtained using FACS (Moflow, Dako). [U-13C]Glucose was purchased from Sigma Isotec. Tunicamycin was purchased from Sigma. MitoTracker Green and MitoTracker Red were from Invitrogen. Antibodies used for Western blot analysis were anti-BiP (immunoglobulin heavy-chain-binding protein)/GRP78 (glucose-regulated protein of 78 kDa) pAb (polyclonal antibody) (Assay Designs), anti-CHOP [C/EBP (CCAAT/enhancer-binding protein)-homologous protein] mAb (monoclonal antibody) (Santa Cruz Biotechnology), anti-β-actin mAb (Sigma), anti-Bak pAb (Upstate Biotechnology), anti-Bax pAb (Santa Cruz Biotechnology), anti-LC3 (microtubule-associated protein 1 light chain 3) pAb (Cell Signaling Technology), anti-GLUT1 pAb (Abcam), anti-Myc mAb (Millipore), anti-Akt pAb (Cell Signaling Technology), anti-hexokinase-2 pAb (Santa Cruz Biotechnology), anti-(p70 S6 kinase) mAb (BD Transduction Laboratories), anti-(phospho-p70 S6 kinase) (Thr421/Ser424) mAb (Cell Signaling Technology) and anti-phospho-Akt (Ser473) mAb (Cell Signaling Technology).
Cell viability, cell size and cell-cycle analysis
Cell viability was measured using propidium iodide (Invitrogen) exclusion assays carried out using flow cytometry (FACSCalibur™, Becton Dickinson) as described previously . Cell sizes were measured by forward scatter as described previously . For cell-cycle analysis, 5×105 cells were sedimented at 300 g for 5 min and washed twice with 500 μl of PBS. Cells were then fixed with 1 ml of 70% ethanol in PBS at 4 °C overnight. After centrifugation at 300 g for 5 min, cells were washed twice with 500 μl of PBS and resuspended in 500 μl of PBS. RNase A (50 units) (Qiagen) was added to samples and incubated at 37 °C for 1 h. Propidium iodide (5 μg) was added to samples which were incubated for 30 min at 37 °C before flow cytometric analysis.
Immunofluorescence staining of Myc–GLUT1
Cells (3×105) were sedimented at 300 g for 5 min and washed twice with 200 μl of blocking buffer (PBS with 2% FBS). Cells were then resuspended with 25 μl of blocking buffer containing 1.25 μl of rat serum (Sigma) and 0.25 μl of Fc Block™ (Becton Dickinson), and incubated on ice for 10 min. Then cells were sedimented at 300 g for 5 min and resuspended with 25 μl of blocking buffer containing 3 μl of anti-Myc antibodies. Following 20 min of incubation on ice, 150 μl of blocking buffer was added and the cells were sedimented. After a wash with 200 μl of blocking buffer, cells were resuspended with 50 μl of blocking buffer containing 1 μl of phycoerythrin-conjugated rat anti-(mouse IgG1) antibodies (Becton Dickinson) and incubated on ice for 20 min in the dark. Blocking buffer (150 μl) was added to the samples followed by centrifugation at 300 g for 5 min. After washing with 200 μl of blocking buffer, immunofluorescence was assessed by flow cytometry (FACSCalibur™) or by fluorescence microscopy (EVOS®, Advanced Microscopy Group).
Cells (4×106) were collected in a 1.5 ml Eppendorf tube by centrifugation at 300 g for 5 min, and cell pellets were fixed with 3% (w/v) glutaraldehyde in 0.1 M sodium cacodylate at 4 °C overnight. Before embedding, cells were treated with 2% (w/v) osmium tetroxide followed by an increasing gradient dehydration step using ethanol and propylene oxide. Cells were then embedded in LX-112 epoxy plastic (Ladd Research). Ultrathin sections of 80 nm were cut, placed on uncoated copper grids and stained with lead citrate and saturated aqueous uranyl acetate. Images were obtained with a Philips CM12 transmission electron microscope at 80 kV as described previously .
Measurement of lactate, ATP, mitochondrial potential and oxygen consumption
Lactate in the growth medium was measured using Lactate Reagent (Trinity Biotech). In this assay, lactate is converted into pyruvate and H2O2 by lactate oxidase. The resulting H2O2 is detected by peroxidase-catalysed oxidative condensation of a chromogen, yielding a product with maximal absorption at 540 nm. For this assay, 1 μl of sample was mixed with 100 μl of Lactate Reagent in a 96-well plate. Following incubation for at least 5 min at 25 °C, absorbance of samples at 540 nm was measured using a plate reader (Bio-tek).
For measurement of intracellular ATP levels, 0.5×106 cells were collected and resuspended in 0.5 ml of boiling water. The ATP concentration in the supernatant was determined by a bioluminescence reaction using firefly luciferase and D-luciferin (Invitrogen). This assay is based on ATP-mediated light production by luciferase (emission maximum ~560 nm). Briefly, 7 μl of samples or standard solutions was added into 100 μl of reaction solution in a 96-well plate. The readings at emission wavelength 560 nm were obtained using a luminometer (Molecular Devices).
To measure mitochondrial mass and membrane potential, cells were incubated with 200 nM MitoTracker Green or MitoTracker Red at 37 °C for 1 h. The fluorescence intensity of the dye was determined by flow cytometry according to the manufacturer's instructions (Invitrogen).
For oxygen consumption, 5×106 cells were collected and resuspended in 0.55 ml of fresh RPMI 1640 medium. Samples were loaded into the chamber of a Strathkelvin Oxygen System and the loss of oxygen in the chamber was monitored.
NMR sample preparation
IL-3-dependent Bak−/−Bax−/− cells were cultured in RPMI 1640 medium containing 5 mM unlabelled glucose. Cells (5×106) were collected and washed three times with 50 ml of glucose-free RPMI 1640 medium, followed by resuspension in RPMI 1640 medium containing 5 mM [U-13C]glucose at a final cell concentration of 0.20×106/ml. DMSO (control) or tunicamycin (final concentration of 3 μg/ml) were added to cells. The growth medium was collected, frozen in liquid nitrogen and stored at −80 °C at 0, 3, 6, 9, 24, 30 and 48 h time points. Cells (5×106) were collected by centrifugation at 300 g for 5 min and washed three times with 20 ml of ice-cold PBS at 24 and 48 h time points. Cell pellets were frozen in liquid nitrogen and stored at −80 °C. To extract metabolites, 200 μl of growth medium sample was mixed with 200 μl of 40% ice-cold TCA (trichloroacetic acid) or 300 μl of 10% TCA was added to cell pellets. Samples were centrifuged at 16000 g for 20 min and freeze-dried as described previously [15,16].
NMR spectra were recorded at 14.1 T on a Varian Inova spectrometer equipped with a 5 mm inverse triple resonance cold probe (cells), at 20 °C. One-dimensional NMR spectra were recorded with an acquisition time of 2 s and a recycle time of 5 s. Concentrations of metabolites and 13C incorporation were determined by peak integration of the 1H-NMR spectra referenced to the DSS (2,2-dimethyl-2-silapentane-5-sulfonic acid) methyl groups, with correction for differential relaxation, as described previously [15–18]. 1H Spectra were typically processed with zero filling to 131 k-points, and apodized with an unshifted Gaussian and a 0.5 Hz line-broadening exponential. 13C profiling was achieved using one-dimensional 13C-edited HSQC (heteronuclear single-quantum coherence) 1H spectra recorded with a recycle time of 1.5 s, with 13C GARP (globally optimized alternating-phase) decoupling centred at 80 p.p.m. (1JCH set to 150 Hz) and during the proton acquisition time of 0.15 s.
TOCSY spectra were recorded with a mixing time of 50 ms and a B1 field strength of 8 kHz with acquisition times of 0.341 s in t2 and 0.05 s in t1. The fits were zero-filled once in t2, and linear-predicted and zero-filled to 4096 points in t2. The data were apodized using an unshifted Gaussian and a 1 Hz line-broadening exponential in both dimensions. Specific 13C isotopomers and fractional incorporation of 13C were determined by comparing the areas or volume of satellite peaks with the total integrated area or volume, with appropriate corrections for differential relaxation as described previously [15–18].
Glucose consumption was quantified by NMR using the 13C-1αH satellite signals centred at 5.22 p.p.m. This accounts for 36% of the total glucose. The 13C-labelled and unlabelled lactate and alanine concentrations were determined by integration of the methyl peak and its satellites and normalized to the concentration of the standard DSS. From this, the amount of glucose consumed, ΔGlc, and the fraction converted into lactate plus alanine, F, could be estimated, according to eqns (1) and (2) [15–18]:
The expression 1−F then represents the glucose carbon that enters other metabolites and macromolecules in the cell mass or otherwise not accounted for. Abundant media components (e.g. threonine and valine) were similarly integrated and their concentrations were determined as a function of sampling time to assess the utilization of essential amino acids. The concentrations of choline metabolites (choline, phosphocholine and glycerophosphocholine) were determined from the areas of the trimethyl ammonium resonance near 3.22 p.p.m. as described previously [15–18]. NMR assignments were made according to the chemical shifts and scalar coupling patterns in TOCSY and by reference to our databases [15–18].
Tunicamycin inhibits cell proliferation and induces autophagy in the absence of apoptosis
ER stress has profound effects on cellular homoeostasis . However, the details of how ER stress might influence cellular metabolism are largely unknown. Prolonged ER stress often induces multiple forms of cell death, including apoptosis and autophagy . It is difficult to evaluate the effects of prolonged ER stress on cellular metabolism as membrane integrity of organelles in cells undergoing apoptosis is often disrupted, leading to the leak of cellular metabolites from the dying cells. To avoid the perplexing variable of apoptosis, we used IL-3-dependent Bak−/−Bax−/− haemopoietic cells as a system to evaluate the effects of prolonged ER stress on cellular metabolism. Bak and Bax double deficiency in these cells make them resistant to apoptosis induced by IL-3 withdrawal . Among several ER stress inducers examined in IL-3-dependent Bak−/−Bax−/− cells, tunicamycin, an inhibitor of protein N-glycosylation , induced dramatic ER stress and minimal cell death (Figures 1A and 1C). In the presence of 3 μg/ml tunicamycin, Bak−/−Bax−/− cells did not proliferate significantly, but remained essentially 100% viable (Figures 1B and 1C). In addition, tunicamycin induced cell arrest at G1-phase (Figures 1D and 1E). Re-expression of Bak or Bax in Bak−/−Bax−/− cells caused cells to undergo apoptosis in the presence of tunicamycin (see Supplementary Figure S1 at http:www.BiochemJ.org/bj/435/bj4350285add.htm).
Tunicamycin affects proliferation and cell cycle, but not viability of IL-3-dependent cells lacking Bak and Bax
To characterize cells under ER stress further, high-power transmission electron microscopy experiments were carried out to examine subcellular structures. At 12 h after tunicamycin treatment, early autophagosomes were observed in the treated cells (Figure 2A). When quantified, the number of autophagosomes in tunicamycin-treated cells was greatly increased compared with that of the untreated cells (Figure 2B). Autophagy persisted over 48 h of tunicamycin treatment as the cytoplasm of the treated cells was increasingly occupied with vesicular structures that contained degraded intracellular organelle remnants and exhibited characteristics of lysosomes (Figure 2A). The sizes of the treated cells were also noticeably smaller in comparison with those of the untreated cells (Figure 2C), indicating that cells underwent atrophy. Furthermore, expression of LC3 upon tunicamycin treatment was examined (Figure 2D). The conversion of the LC3-I form into the LC3-II form, a hallmark of autophagy, was observed in tunicamycin-treated cells, providing more evidence that tunicamycin induced autophagy in the absence of apoptosis (Figure 2E). The observation of autophagy in tunicamycin-treated Bak−/−Bax−/− cells is consistent with previously reported autophagy induced by ER stress in many other cell types [3,6,7].
Tunicamycin induces autophagy in Bak−/−Bax−/− cells
Tunicamycin inhibits glucose uptake and glycolysis
To understand how tunicamycin might affect cellular metabolism, we first used high-resolution NMR to measure changes in the concentrations of metabolites in the growth medium. The concentrations of essential amino acids such as threonine remained almost the same, and the concentrations of unlabelled lactate in the medium were largely unchanged over 48 h of tunicamycin treatment (Figure 3A). In contrast, the consumption of [13C]glucose from the medium was considerably higher in the control cells than that in the treated cells (Figure 3B). Furthermore, the production of [13C]lactate from glucose via glycolysis accounted for more than 50% of the glucose carbon in the control cells at 24 h, but was less than 35% in the treated cells (Figure 3C). Interestingly, consumption of glutamine from the medium was increased in tunicamycin-treated cells (Figure 3D). Using an enzymatic assay, it was confirmed that the total amount of lactate in the medium was reduced at least 2-fold in the presence of tunicamycin (Figure 3E). Hence, glycolytic flux to lactate was considerably attenuated in the presence of tunicamycin.
Tunicamycin reduces glucose consumption and glycolysis
Tunicamycin reduces oxidative metabolism
In addition to glucose utilization and lactate production, we also examined the effects of tunicamycin on oxidative metabolism. The relative concentrations of critical intracellular metabolites that reflect glycolysis, the pentose phosphate pathway, the tricarboxylic acid cycle and pyrimidine biosynthesis were determined. As we monitored the incorporation of 13C atoms derived from [13C]glucose into metabolites, these events represent de novo biosynthesis of metabolites (see Supplementary Figure S2 at http:www.BiochemJ.org/bj/435/bj4350285add.htm). In one-dimensional HSQC experiments, only protons attached to 13C were observed (Figure 4A). As the natural abundance of 13C is only 1.1%, the peaks detected in this experiment largely reflect de novo biosynthesis of compounds that utilize glucose carbon. The concentrations of newly synthesized metabolites were generally higher in the control cells than those in the treated cells, consistent with the decreased glucose consumption in the presence of tunicamycin (Figure 3B).
Major 13C-labelled soluble metabolites in cells in the absence or presence of tunicamycin were detected by SIRM
More details of de novo biosynthesis of cellular metabolites were revealed by the TOCSY experiments (Figures 4B and 4C). The 13C-labelling patterns in the metabolite markers of glycolysis and the tricarboxylic acid cycle, including lactate, alanine, glutamate and aspartate, were determined (Figure 4B). Both alanine and lactate exhibited the characteristic square pattern of 13C satellites surrounding a central resonance that represents unlabelled compound, indicating that alanine and lactate were synthesized directly from glucose (via glycolysis) with no significant metabolic scrambling [15–18]. In contrast, both glutamate and aspartate resonances showed a more complex pattern of satellites, corresponding to unlabelled species, molecules that contain 13C at both positions (e.g. C-2 and C-3 in aspartate), as well as two additional species. For aspartate, the additional species correspond to 13C-2 12C-3 and 12C-2 13C-3. The latter pattern arises from tricarboxylic acid cycle activity in which doubly labelled acetyl-CoA from uniformly labelled pyruvate condense with unlabelled oxaloacetate, producing first 2-OG (2-oxoglutarate) labelled at C-4 and C-5 followed by further transformation to oxaloacetate via the tricarboxylic acid cycle (see Supplementary Figure S2). Because of the scrambling at the succinate step, the oxalate comprises a 1:1 mixture of 13C-1 13C-2 and 13C-3 13C-4, which is converted into aspartate by transamination. In a single turn of the tricarboxylic acid cycle, this would give rise to a different satellite pattern from what was observed. Only with a second addition of labelled acetyl-CoA, does the multiply labelled aspartate appear, in which both C-2 and C-3 are labelled in the same molecule. Similar remarks apply to the glutamate-labelling pattern, as glutamate is synthesized by transamination of 2-OG (see Supplementary Figure S2).
Hence, the tricarboxylic acid cycle is fully operative in IL-3-dependent Bak−/−Bax−/− cells. The relative abundance of the various isotopomers was quantified as described previously [15–18]. As shown in Table 1, there was a substantial decrease in the amount of multiply labelled glutamate and aspartate in the tunicamycin-treated cells, consistent with the decreased glucose uptake and flow through glycolysis. As aspartate is a direct precursor of pyrimidine ring biosynthesis, the uracil in UTP exhibits the same labelling pattern as the aspartate (see Supplementary Figure S2). Shown in the TOCSY spectra of the free nucleotide pools of the cells (Figure 4C), the degree of labelling of uracil rings in the control cells was substantially higher than in the treated cells (Table 1). As the orotate dehydrogenase step occurs in mitochondria, this provides further evidence that functional respiration and tricarboxylic acid cycle activity in the mitochondria of Bak−/−Bax−/− cells was compromised by tunicamycin treatment. Interestingly, significant ribose biosynthesis (pentose phosphate pathway) was observed in tunicamycin-treated cells, albeit at a slightly reduced rate compared with that in the control cells (Table 1). Nevertheless, even at the 48 h time point, tunicamycin-treated cells were still turning over RNA, necessitating replacement by de novo nucleotide biosynthesis.
|Uracil in UTP||12C-6 12C-5||63||83||1.32|
|Uracil in UTP||12C-6 12C-5||63||83||1.32|
The results presented so far demonstrate that tunicamycin induces autophagy, with a concomitant decrease in demand for glycolysis and tricarboxylic acid cycle activity. This should lead to a decreased demand for ATP synthesis. To corroborate this, we measured the mitochondrial membrane potential in response to tunicamycin using mitochondrion-selective dyes. As shown in Figures 5(A) and 5(B), tunicamycin treatment reduced the fluorescent staining of both mitochondrial potential-independent MitoTracker Green and mitochondrial potential-sensitive MitoTracker Red, indicating that both mitochondrial mass and mitochondrial membrane potential were decreased in tunicamycin-treated cells, consistent with the catabolism of intracellular organelles in cells undergoing autophagy. We also measured cellular respiration rate as well as the steady-state levels of ATP in response to tunicamycin. The respiration rate decreased more than 3-fold in the presence of tunicamycin (Figures 5C and 5D), with a concomitant 2-fold decrease in the steady-state ATP level (Figure 5E), consistent with decreased oxidative phosphorylation.
Tunicamycin reduces metabolic activities of mitochondria
GLUT1 levels on the cell surface are reduced in tunicamycin-treated cells
Glucose uptake is partially controlled by the cell-surface expres-sion of glucose transporters [21,22], and the ubiquitously expressed glucose transporter GLUT1 is primarily responsible for glucose uptake in haemopoietic cell lines [23,24]. To explore the molecular mechanism of the reduced glucose uptake in tunicamycin-treated IL-3-dependent cells, the effect of tunicamycin on GLUT1 expression was examined. As shown in Figures 6(A) and 6(B), tunicamycin did not significantly affect either total GLUT1 or hexokinase-2 protein expression levels. Although tunicamycin inhibits glycosylation and GLUT1 is a highly glycosylated protein, tunicamycin did not seem to alter glycosylation patterns of GLUT1. As GLUT1 transportation to the cell surface is critical for its function, we examined cell-surface expression levels of GLUT1 using an immunofluorescence staining approach. Owing to the limitations in commercially available anti-GLUT1 antibodies, we failed to detect endogenous GLUT1 expression on the cell surface. As previous studies show that trafficking of exogenously expressed GLUT1 reflects that of endogenous GLUT1 in haemopoietic cells [25,26], Myc-tagged GLUT1 was stably expressed in the IL-3-dependent Bak−/−Bax−/− cells to achieve highly specific tracking of surface localization of GLUT1. In the absence of tunicamycin, Myc-tagged GLUT1 seemed to be on the cell surface, but it was largely localized intracellularly after 48 h of tunicamycin treatment (Figure 6C). Flow cytometric analyses demonstrated further that tunicamycin treatment substantially reduced the surface levels of Myc-tagged GLUT1, possibly accounting for the reduction in glucose uptake in tunicamycin-treated cells (Figures 6D and 6E). Although unlikely, it is still possible that that trafficking of exogenously expressed GLUT1 is different from that of endogenous GLUT1.
Tunicamycin reduces GLUT1 levels on the cell surface and Akt signalling
Akt signalling is involved in GLUT1 trafficking and autophagy induction caused by tunicamycin
As the PI3K (phosphoinositide 3-kinase)/Akt signalling pathway has been demonstrated to be important for maximal cell-surface expression of GLUT1 [25,27], we examined whether the PI3K/Akt signalling pathway might be influenced by tunicamycin. At 24 h, whereas the expression levels of Akt and the Akt downstream target p70 S6 kinase were not significantly affected, phosphorylation of p70 S6 kinase was completely abolished (Figure 6F), indicating that the Akt signalling pathway was down-regulated in the treated cells. At 48 h, both Akt and p70 S6 kinase expression levels were reduced. These results indicate that tunicamycin down-regulates Akt signalling.
To elucidate further the role of the Akt signalling pathway, a constitutively active form of Akt (myristoylated Akt1) was exogenously expressed in IL-3-dependent Bak−/−Bax−/− cells (Figure 7A). Activated Akt signalling enabled cells to maintain exogenously expressed GLUT1 levels on the cell surface, suggesting that the tunicamycin-induced decrease in GLUT1 surface levels is caused by Akt inactivation (Figure 7B). Furthermore, myristoylated Akt1 overexpression largely blocked the conversion of the LC3-I form into the LC3-II form and cell atrophy caused by tunicamycin, providing evidence that tunicamycin-induced autophagy is blocked by activation of Akt signalling (Figures 7C–7E). Thus reduced Akt signalling is involved in decreased cell-surface GLUT1 expression, which might lead to a decrease in glucose uptake and subsequent changes in cellular metabolism and cell-fate determination (Figure 6F).
Akt activation abrogates the effects of tunicamycin on GLUT1 trafficking and autophagy induction
We have utilized a NMR-based metabolomics approach to systematically analyse the effects of prolonged ER stress on nutrient utilization, lactate fermentation and de novo biosynthesis of metabolites that reflect important cellular metabolic processes, including glycolysis, the tricarboxylic acid cycle, pyrimidine biosynthesis and the pentose phosphate pathway. In response to sustained ER stress, IL-3-dependent Bak−/−Bax−/− haemopoietic cells demonstrate co-ordinated bioenergetic responses, leading to an increase in catabolic metabolism and a decrease in anabolic metabolism. As the major carbon source for energy production and anabolic metabolism in IL-3-dependent cells, extracellular glucose consumption decreased upon ER stress. Bioenergetic function of mitochondria in cells undergoing ER stress was compromised, probably as a result of both autophagy and reduced pyruvate supply from glucose oxidation. The observed changes in cellular metabolism could be attributed to the loss of cell-surface GLUT1 expression, which is probably caused by down-regulation of the Akt signalling pathway.
IL-3-dependent cells deficient in both Bak and Bax were used to investigate ER stress-induced changes in cellular metabolism because, unlike normal cells, these cells will survive exposure to such stress. The inhibition of anaerobic metabolism has been reported to lead to compromised mitochondrial function . One example is the reduction in glucose utilization caused by growth factor withdrawal in IL-3-dependent cells, which is associated with disruption of mitochondrial function and subsequent apoptosis . In the presence of the pro-apoptotic Bcl-2 proteins Bak or Bax, the alterations in glucose metabolism induced by ER stress may lead to decreased mitochondrial membrane integrity and subsequent release of apoptogenic factors, such as cytochrome c, and ultimately apoptosis. Therefore changes in key cellular metabolic processes revealed in the present study could represent a common bioenergetic response to ER stress in haemopoietic cells.
Our systematic analysis of nutrient utilization, lactate fermentation reactions and de novo biosynthesis of intra-cellular metabolites has revealed comprehensive information about metabolic changes in IL-3-dependent cells undergoing ER stress-induced autophagy. Whereas the de novo synthesis of metabolites reporting tricarboxylic acid cycle activities, such as glutamate and aspartate, was significantly reduced in the treated cells, the synthesis of ribose moieties in free nucleotides was almost unchanged, indicating that the pentose phosphate pathway was still active. In addition to generating ribose 5-phosphate used in the synthesis of nucleotides, the pentose phosphate pathway is also critical in producing NADPH from NADP+ in cells. NADPH is required for maintaining the antioxidant activities of reduced glutathione . Several recent reports support the importance of the pentose phosphate pathway in protecting cells from death through scavenging toxic free radicals and re-reducing the oxidized biomolecules. For instance, a p53-inducible protein, TIGAR [TP53 (tumour protein 53)-induced glycolysis and apoptosis regulator], has been shown to act by lowering ROS (reactive oxygen species) levels through the pentose phosphate pathway and protecting against cell death caused by oxidative stress . Similarly, NADPH generated by the pentose phosphate pathway is critical for the survival of oocytes in vitro through regulating caspase 2 activity . It is conceivable that cells undergoing ER stress must maintain the activity of the pentose phosphate pathway and NADPH levels to survive under autophagic conditions.
The regulation of cellular metabolism by different death stimuli in IL-3-dependent cells has been reported previously and the underlying mechanisms may be distinct and complex [14,25,28,33,34]. Genotoxic drugs inhibit both anaerobic and aerobic metabolism in IL-3-dependent cells, probably due to the reduction in expression levels of GLUT1 and several key enzymes involved in glycolytic metabolism, including hexokinase, phosphofructokinase and pyruvate kinase . In this earlier study, within 4 h of drug treatment, a reduction in mRNA levels for these proteins could be observed. However, in the present study, we found that the expression levels of GLUT1 and hexokinase appear unchanged during 48 h of tunicamycin treatment. Likewise, the total GLUT1 expression level in Bak−/−Bax−/− cells is maintained for at least 1 week following IL-3 withdrawal . Similar to the bioenergetic responses observed in our studies, IL-3 withdrawal also leads to co-ordinated suppression of glucose utilization and promotion of autophagy . Growth factor withdrawal also causes reduction in cell-surface GLUT1 expression [25,28]. As autophagy induced by growth factor withdrawal is essential for maintaining cell survival , it is conceivable that the autophagy observed in cells undergoing ER stress could actually be important for cell survival over an extended period of ER stress.
Emerging evidence indicates that ER stress negatively regulates the oncogenic PI3K/Akt signalling pathway [35–37]. BiP/GRP78, an ER luminal protein, appears to be a major chaperone able to suppress PI3K/Akt signalling. Our studies confirm the negative impact of ER stress on PI3K/Akt signalling with subsequent inhibition of glucose uptake and bioenergetic responses leading to cell-fate determination. Although not transformed, Bak−/−Bax−/− cells share important characteristics with cancer cells as they cannot undergo apoptosis, an important survival mechanism in cancer cells, but proliferate rapidly in the presence of nutrients and the growth factor IL-3. Furthermore, in the presence of tunicamycin, they exhibit the metabolic characteristics of non-dividing cells that can be maintained in culture under defined conditions. Thus these apoptosis-deficient cells provide a good model system for investigating the metabolic consequences of cytostasis and transformation in the absence of apoptosis.
A recent elegant study by Wang's group has identified ENTPD5 (ectonucleoside triphosphate diphosphohydrolase 5), an ER UDPase to hydrolyse UDP to UMP, as an integral part of the PI3K/Akt/PTEN (phosphatase and tensin homologue deleted on chromosome 10) signalling loop . Upon Akt activation, ENTPD5 is transcriptionally up-regulated to promote protein N-glycosylation and folding in the ER, resulting in the increase in lactate production and cell proliferation. On the other hand, our studies demonstrate that inhibition of protein N-glycosylation by tunicamycin leads to the decreased Akt activities and subsequent reduction in glucose uptake and glycolysis, providing more evidence that targeting protein N-glycosylation in cancer cells could be a potential therapeutic strategy.
immunoglobulin heavy-chain-binding protein
C/EBP (CCAAT/enhancer-binding protein)-homologous protein
ectonucleoside triphosphate diphosphohydrolase 5
fetal bovine serum
glucose transporter 1
glucose-regulated protein of 78 kDa
heteronuclear single-quantum coherence
microtubule-associated protein 1 light chain 3
stable isotope-resolved metabolomics
unfolded protein response
Xiaoli Wang, Colins Eno, Brian Altman, Yanglong Zhu, Guoping Zhao and Kristen Olberding designed experiments, performed experiments, and contributed to data analysis and preparation of the paper. Jeffrey Rathmell and Chi Li designed the study and analysed results. Chi Li wrote the paper.
This work was supported by the National Institutes of Health [grant numbers P20RR018733, K01CA106599 and K01CA106599S1 (to C.L.) and R01CA123350 (to J.C.R.)], and the James Graham Brown Cancer Center (to C.L.).