Apoptosis-induced mitochondrial dysfunction causes cytoplasmic lipid droplet formation

A characteristic of apoptosis is the rapid accumulation of cytoplasmic lipid droplets, which are composed largely of neutral lipids. The proton signals from these lipids have been used for the non-invasive detection of cell death using magnetic resonance spectroscopy. We show here that despite an apoptosis-induced decrease in the levels and activities of enzymes involved in lipogenesis, which occurs downstream of p53 activation and inhibition of the mTOR signaling pathway, the increase in lipid accumulation is due to increased de novo lipid synthesis. This results from inhibition of mitochondrial fatty acid β-oxidation, which coupled with an increase in acyl-CoA synthetase activity, diverts fatty acids away from oxidation and into lipid synthesis. The inhibition of fatty acid oxidation can be explained by a rapid rise in mitochondrial membrane potential and an attendant increase in the levels of reactive oxygen species.

The responses of cells to stress are intimately connected to cellular metabolism. 1 Central among the signaling pathways connecting the two are the phosphoinositide 3-kinase-AKT-mTOR pathway, which promotes cell growth and survival by increasing energy production and protein synthesis, 2 and the transcription factor p53, which under conditions of mild or transient stress, induces cell-cycle arrest and DNA repair before the cells re-enter a normal proliferative state when the stress has been resolved. In this situation p53 modulates the expression of several enzymes involved in metabolism, including a fructose-2,6-bisphosphatase, which diverts flux from glycolysis into the pentose phosphate pathway, 3 increasing the production of NADPH, which acts, via reduced glutathione, to lower the raised levels of reactive oxygen species (ROS) and a mitochondrial glutaminase, which through increasing glutamate concentration increases the levels of glutathione. 4 When the stress results in severe or irreparable damage, p53 produces a permanent inhibition of cell proliferation, through induction of apoptosis or senescence. 5 Under these circumstances, p53 inhibits the AKT and mTOR signaling pathways by activating transcription of negative regulators such as PTEN and tuberous sclerosis complex 2 (TSC2). 6 The re-direction of cellular metabolism during apoptosis is also controlled by AMP-activated protein kinase (AMPK). Increases in the levels of AMP in stressed cells stimulate AMPK activity, which increases the activities of those pathways that generate ATP while at the same time inhibiting those pathways that consume ATP, such as those involved in biosynthesis. 7 Following induction of cell death, p53 induces the expression of Sestrins1 and 2, which possess antioxidant activity, and, through Sestrin 2, also activates AMPK, which forms a complex that activates TSC2 and inhibits the mTORC1 pathway 8 (Figure 1). DNA damage induced by drugs, such as etoposide, also results in increased AMPK activity, via activation of ATM, as a mechanism to fulfill the energetic requirements of the DNA repair process. 9 The metabolic changes resulting from the induction of cell death should result in an inhibition of lipid biosynthesis. Inhibition of the AKT and mTORC1 pathways results in a decrease in nuclear translocation of the sterol-regulatory element-binding protein 1 (SREBP-1), which regulates expression of lipogenic enzymes, 10 (Figure 1) and activated AMPK has a direct effect on lipid synthesis by phosphorylating and inhibiting hydroxymethylglutaryl-CoA reductase and acetyl-CoA carboxylase (ACC1), 11 therefore decreasing the levels of malonyl-CoA ( Figure 1). Malonyl-CoA regulates the balance between lipogenesis and b-oxidation as it is both a substrate for fatty acid synthase (FASN) and an inhibitor of carnitine palmitoyltransferase I (CPT-1), which mediates uptake of fatty acids into the mitochondria. 12 However, despite the expected inhibition of cellular lipid synthesis, an accumulation of cytoplasmic lipid droplets after induction of apoptosis has long been reported. [13][14][15] As these give rise to a high-resolution 1 H nuclear magnetic resonance signal, 14 which can be detected in vivo, they have been used as a non-invasive marker of tumor cell death post-treatment. 13,16 However, why these droplets accumulate following the induction of apoptosis is not clear, although various suggestions have been made as to their origin, including redistribution of different lipid species between cellular compartments, 17 increased neutral lipid synthesis during an active phase of apoptosis, 18 increased lipid content caused by organelle degradation or autophagy, 19 or enhanced phospholipase A2 activity. 16 Lipid droplets are dynamic organelles consisting of a central core of neutral lipids, mainly triacylglycerides (TAG) and cholesterol esters, surrounded by a monolayer of phospholipids that contains several proteins associated with lipid metabolism and membrane trafficking. 20 Lipid droplets have been associated with several processes, including energy regulation, fatty acid storage, control of lipid signaling molecules and mediation of lipid toxicity. 21 They also interact with other structures inside the cell, such as mitochondria and the endoplasmic reticulum (ER), 20 which are intimately involved in the progression of the apoptotic program.
We show here that the accumulation of cytoplasmic lipid droplets during apoptosis can be explained by an early inhibition of mitochondrial fatty acid oxidation (FAO) and that, despite the decrease in the cell's capacity for lipid synthesis, which occurs downstream of activation of p53 and inhibition of the mTOR signaling pathway, this results in increased TAG synthesis and lipid droplet formation.

Results
Induction of apoptosis leads to an early increase in cytoplasmic lipid droplet formation and mitochondrial membrane potential. Etoposide-induced apoptosis of murine lymphoma cells (EL4) resulted in an increase in the Figure 1 Apoptosis affects cell signaling pathways that control lipid synthesis and energy metabolism. The enzymes involved in de novo lipid synthesis are depicted as well as the signaling pathways that regulate them. Lipids are synthesized from citrate, which is transported into the cytosol following synthesis in the mitochondria by condensation of acetyl-CoA with oxaloacetate. Glucose is the principal origin of citrate carbon, although other metabolites can also be used as carbon donors. Lipid synthesis also requires NADPH, which originates from the pentose phosphate pathway and from the reaction catalyzed by malic enzyme. Pictures of the mitochondrion and cell membrane were obtained from Servier Medical Art (www.servier.com). ACC1, acetyl-CoA carboxylase; ACLY, ATP-citrate lyase; ACS, acyl-CoA synthetase; AMPK, AMPactivated kinase; ATM, ataxia-telangiectasia mutated; CPT-1, carnitine palmitoyltransferase 1; DAG, diacylglycerides; DAGAT, diacylglycerol acyltransferase; DHAP, dihydroxyacetone phosphate; FASN, fatty acid synthase; FATP, fatty acid transport protein; FFA, free fatty acids; G3P, glyceraldehyde 3-phosphate; G6P, glucose 6phosphate; Glut, glucose transporter; HK, hexokinase; LDH, lactate dehydrogenase; PDH, pyruvate dehydrogenase; PK, pyruvate kinase; SREBP-1, sterol-receptor element binding protein; TAG, triacylglycerides; TIGAR, Tp53-induced glycolysis and apoptosis regulator; TSC2, tuberous sclerosis complex 2 number of cytoplasmic lipid droplets (Figure 2a). This increase was observed as early as 2 h after drug treatment when the levels of early apoptosis and late apoptosis/ necrosis, determined by annexin V (AV) and 7aminoactinomycin D (7AAD) staining respectively, were similar to those in untreated cells (Figures 2b and c). The increase in lipid staining was evident in etoposide-treated cells that appeared to be viable (AV and 7AAD negative), indicating that lipid droplet formation is an early event following induction of apoptosis in these cells. This increase in lipid droplet number following the induction of apoptosis, which has been reported in previous studies, 18,22 was also observed in other cell lines that had been treated with different apoptosis-inducing agents, including the prostate cancer cell lines C4-2b and DU145 and the human colon adenocarcinoma, HCT116 (Supplementary Figure S1). This phenomenon is not specific to tumor cells as we also observed it in mouse embryo fibroblasts (MEFs) treated with etoposide (Supplementary Figure S2).
The effect of apoptosis induction on mitochondrial function was analyzed with Mitotracker orange (MTO), which stains functionally active mitochondria, accumulating in a membrane potential-dependent manner. 23 Increased staining intensity in drug-treated cells indicated an increase in mitochondrial membrane potential (Figures 2a and c), which was already apparent by 2 h after drug treatment in the viable and early apoptotic cell populations. To confirm the increase in mitochondrial membrane potential, cells were also stained with tetramethylrhodamine methyl ester (TMRM), which is a specific indicator of membrane potential. 24 There was a fivefold increase in dye intensity by 2 h after etoposide treatment (507 ± 19% in drug-treated cells versus 100 ± 1.5% in control cells; n ¼ 3, Po0.00001; mean ± S.E.M.).
Increased lipid droplet formation correlates with increased lipogenesis. To investigate the origin of lipid droplet accumulation following induction of apoptosis in EL4 cells, we analyzed de novo lipid synthesis by measuring the incorporation of [1,2-14 C 2 ]acetate into cellular lipids. The labeled acetate was added to the cell suspension 1 h before cell harvest and the incorporation of 14 C label into neutral and polar lipids (Figures 3a and b) was determined. As early as 2 h after drug treatment, there was a threefold increase in acetate incorporation into triacylglycerol (TAG) relative to untreated control cells (Po0.03), and this continued to rise until 8 h after treatment, whereupon there was a subsequent decrease ( Figure 3c). The decrease coincided with a significant loss of cell viability, which correlated with increased levels of cleaved caspase 3 (Figures 3g and h). By 14 and 16 h after drug treatment, acetate incorporation into TAG was less than in untreated control cells ( Figure 3c). Next, we investigated incorporation of fatty acids into TAG by incubating the cells with 14 C-palmitate (Figures 3d and e). There was no significant increase in incorporation compared with untreated control cells until 4 h after drug treatment, when this increased incorporation continued to rise until peaking at 12 h and was still higher than in untreated control cells at 14 and 16 h (Figure 3f). Both [1,2-14 C 2 ]acetate and 14 C-palmitate showed a general decrease in incorporation into polar lipids following drug treatment (Figures 3b and e), as observed previously. 25 MEFs also showed increased lipid synthesis at 24 h after treatment with etoposide (Supplementary Figure S2B).
Cell signaling pathways affected by the induction of apoptosis can inhibit lipogenesis. Etoposide-induced apoptosis in EL4 cells produced the expected increase in p53 protein ( Figure 4a) and in the expression of the p53responsive genes, p21 and Sestrin 2 (Figures 1 and 4b).
There was an increase in AMPK phosphorylation, which coincided with increased expression of Sestrin 2, and a decrease in phosphorylated p70-S6K (Figure 4a), consistent with inhibition of the activity of the mTORC1 complex. There appeared to be no significant change in Akt serine 473 phosphorylation, although there was a decrease in the level of the total protein at later time points (Figure 4a). The increase in AMPK phosphorylation and activity and the phosphorylation and consequent inhibition of mTORC1 are expected to decrease the expression of the lipogenic enzymes, 10 and also to inhibit lipogenesis more acutely through phosphorylation and inhibition of ACC1 7 ( Figure 1).  FASN (Figure 4d). The expression of ACS is activated by increased levels of FFA and by lipid droplets. 26 Accordingly, the observed increase in ACS protein (Figure 4d) paralleled the increase in lipid droplet number (Figure 2c) and could explain the sustained incorporation of 14 C-palmitate into TAG at the later time points after drug treatment (Figure 3f). Furthermore, the activity of ACS is enhanced by deacetylation by sirtuin 1 (Sirt1) in response to a decrease in NAD þ concentration. 27 The observed NAD(H) depletion in EL4 cells treated with etoposide, which has been shown to be due to poly(ADP-ribose)polymerase activation, 28 correlated with increased Sirt1 protein levels (Supplementary Figure S3A), indicating that the activity of the ACS enzyme could also be enhanced at later time points after drug treatment.

Induction of apoptosis affects the expression and
To confirm the importance of ACS and de novo TAG synthesis for lipid droplet formation, cells were treated with Triacsin C (TrC), which is a specific inhibitor of the enzyme. Despite enhancing the induction of apoptosis by etoposide (Supplementary Figure S3B), the drug inhibited lipid droplet formation (Supplementary Figure S3C), decreased the incorporation of 14 C-acetate into TAG and increased 14 C incorporation into FFA (Supplementary Figures S3D  and S3E). However, the increase in ACS content was not observed until 6 h after etoposide treatment and therefore cannot explain the earlier increase in lipid droplet content ( Figure 2c) and TAG synthesis from 14 C-acetate (Figure 3c), which were already apparent at 2 h after drug treatment, and the increased TAG synthesis from 14 Cpalmitate, which was already observed at 4 h after drug treatment (Figure 3f).
Induction of apoptosis increases formation of ROS and decreases lipid b-oxidation. The delayed changes in ACS activity suggest that there must be other factors contributing to the early increases in TAG synthesis and subsequent lipid droplet formation following the induction of apoptosis. The  Figure  S2B). Glucose oxidation in EL4 cells, however, only decreased after 6 h of drug treatment, reaching a 20% decrease by 10 h (Figure 5a). The decrease in glucose oxidation paralleled caspase 3 activation (Figure 3h), suggesting that the decrease in glucose oxidation might be the result of cytochrome c release from the mitochondria, which through formation of the apoptosome could be responsible for the observed caspase 3 cleavage. These results suggest that increased TAG synthesis and lipid droplet accumulation following etoposide treatment could be due to reduced mitochondrial FAO. This apparent inverse correlation between TAG synthesis and lipid droplet formation on the one hand and mitochondrial lipid oxidation on the other was further tested by treating cells with etomoxir, a CPT-1 inhibitor that blocks mitochondrial fatty acid uptake. 29 Treatment of cells with etomoxir for 2 h had no effect on cell viability but resulted in a 20% decrease in oleate oxidation, whereas oleate incorporation into lipids was increased by 50%, indicating a redirection of fatty acids from oxidation into biosynthesis (Figures 5b and c). There was no evidence for involvement of CIDE (cell death-inducing DNA fragmentation factor 45-like effector) proteins 30 in these changes in lipid metabolism (Supplementary Figure S4). The recently demonstrated ROS-dependent inhibition of mitochondrial FAO 31 could provide an explanation for the early decrease in FAO following the induction of apoptosis with etoposide. Consistent with this proposal, we observed that when mitochondrial ROS production was induced by treating EL4 cells for 2 h with antimycin, an inhibitor of complex III that increases mitochondrial H 2 O 2 formation, 32 there was a 40% decrease in oleate oxidation (Figure 5b) and an accompanying increase in 14 C-oleate incorporation into TAG (Figure 5c).
The effect of inducing apoptosis on cellular ROS production was examined by staining cells with carboxy-dichlorodihydrofluorescein diacetate (DCFDA) (Figure 6a). Etoposide treatment resulted in a rapid and marked increase in ROS levels, which were four times the levels observed in control cells by 2 h after drug treatment. This increase in ROS levels paralleled the marked increase in mitochondrial membrane potential determined using MTO (Figures 2a and c) and TMRM. However, while tocopherol appeared to reduce the levels of ROS in etoposide-treated cells to those observed in control cells (Figure 6b), it neither restored oleate oxidation to control levels (Figure 6c) nor decreased oleate incorporation into TAG (Figure 6d). Similar results were obtained with other cytosolic antioxidants (data not shown), suggesting, as indicated by the experiments with antimycin, that it is ROS levels in the mitochondria, and not in the cytosol, that account for the observed inhibition of lipid b-oxidation. To prove that mitochondrial ROS were responsible for inhibition of lipid b-oxidation, we treated cells with mitoubiquinone (MitoQ), which is a mitochondrially targeted antioxidant that blocks mitochondrial oxidative damage. 33 The antioxidant ubiquinone moiety is targeted to the mitochondria by covalent attachment of a lipophilic triphenylphosphonium cation via an aliphatic carbon chain. As a control for the effects of the triphenylphosphonium ion on mitochondrial function, we used MitoQ lacking the ubiquinone moiety (DecylTPP). Cells  (Figure 6c). However, in cells treated with etoposide and MitoQ lipid oxidation increased to above control levels and there was a corresponding decrease in lipid synthesis (Figure 6d).

Discussion
Increased lipid droplet formation following the induction of cell death has been widely observed [16][17][18][19] and has formed the basis of a non-invasive magnetic resonance imaging technique to detect tumor cell death post-treatment. 13 However, the mechanisms responsible for the accumulation of these droplets in dying cells are unclear.
We have shown here, in different cell types, that the induction of cell death resulted in the rapid accumulation of cytoplasmic lipid droplets. In EL4 murine lymphoma cells, this increased droplet accumulation, which was already evident by 2 h after drug treatment, was accompanied by an increase in mitochondrial membrane potential, an increase in ROS production, an increase in de novo neutral lipid synthesis and a decrease in lipid b-oxidation. The increase in lipid synthesis occurred despite the expected overall downregulation or inhibition of enzymes involved in lipogenesis, downstream of p53 and AMPK activation. There was a decrease in FASN mRNA by 4 h and a decrease in protein by 10 h and there was AMPKdependent phosphorylation and inhibition of ACC1 by 2 h and a decrease in its expression by 4 h. There was, however, an increase in the levels of ACS protein at later time points (after 6 h), when the activity of the enzyme is expected to be increased by deacetylation by Sirt1 in response to the decrease in NAD þ concentration. The incorporation of fatty acids into diacylglycerides (DAG) and TAG will be enhanced by the increase in ACS activity and indeed we observed sustained incorporation of 14 C label from palmitate into TAG at a time when incorporation of label from acetate, which relies on the activities of ACC1 and FASN, was declining.
Thus, we propose that lipid droplet accumulation following the induction of apoptosis is due to de novo neutral lipid synthesis as a result of inhibition of mitochondrial fatty acid b-oxidation and re-direction of fatty acids away from oxidation and into lipid synthesis. That inhibition of lipid oxidation could result in such a reciprocal upregulation of lipid synthesis was demonstrated here by treating cells with the CPT-1 inhibitor etomoxir, which inhibited oleate oxidation and increased its incorporation into TAG, and by inhibiting complex III in the mitochondrial respiratory chain with antimycin, which had a similar effect. Treatment of cells with compounds that result in mitochondrial toxicity has been shown previously to result in lipid droplet formation, although this was attributed to lysosomal processing of lipids from damaged mitochondria rather than inhibition of b-oxidation. 19 The demonstration that increases in ROS levels in mammary epithelial cells, caused by matrix detachment, resulted in inhibition of FAO 31 suggested a potential mechanism whereby apoptosis induction could lead to inhibition of lipid oxidation and led us to investigate whether the increased ROS levels observed here could be responsible for the observed decrease in b-oxidation. Consistent with this proposal was the observation that inhibition of the respiratory chain with antimycin, which results in the generation of intramitochondrial H 2 O 2 , 32 both inhibited oleate oxidation and increased its incorporation into lipid. However, while a-tocopherol treatment appeared to prevent completely the etoposide-induced increase in cellular ROS, it neither restored oleate oxidation nor decreased oleate incorporation into TAG; similar results were obtained with other antioxidants. However, when etoposide was combined with a mitochondrially targeted antioxidant, MitoQ, there was recovery of lipid b-oxidation, which was accompanied by a decrease in fatty acid incorporation into TAG. A similarly targeted version of a-tocopherol has been shown to be much more effective at protecting mitochondrial function from oxidative damage than a-tocopherol alone. 34 These results corroborate the relationship between mitochondrial ROS production and inhibition of lipid b-oxidation and confirm the reciprocal relationship between lipid oxidation and its incorporation into neutral lipid droplets. These results are in agreement with previous studies showing that H 2 O 2 inhibits fatty acid b-oxidation 35 and that many of the proteins in heart mitochondria, whose thiol redox state was affected by generation of intramitochondrial ROS, either at complex I by reverse electron transfer or at complex III by inhibition with antimycin, were those involved in FAO. 36 Thus, the model that emerges from our results is that induction of apoptosis leads to a rapid rise in mitochondrial membrane potential and the generation of intramitochondrial ROS, which includes H 2 O 2 . The mitochondrial H 2 O 2 reacts reversibly with thiol groups in the enzymes involved in fatty acid b-oxidation and inhibits them. The inhibition of b-oxidation, when combined with the increase in ACS activity, diverts fatty acids into TAG synthesis and the formation of lipid droplets.
Lipid droplets could have several functions in the apoptotic and pre-apoptotic cell. In damaged cells, where there is a ROS-dependent increase in mitochondrial membrane potential and inhibition of fatty acid b-oxidation, 31 increased TAG synthesis and lipid droplet formation may prevent the accumulation of toxic FFAs. Consistent with this proposal, we showed that inhibition of fatty acid synthesis with TrC increased FFA accumulation and enhanced cell death and that inhibition of FAO, with the CPT-I inhibitor etomoxir, enhanced lipid synthesis. In skeletal muscle cells, oleate has been shown recently to protect against palmitate-mediated lipotoxicity by channeling palmitate into TAG synthesis and by promoting palmitate oxidation 37 and in pancreatic b-cells increased lipogenesis or b-oxidation were shown to protect against the toxicity associated with increased FFAs. 38 There is also evidence that macrophages take advantage of lipid droplet accumulation as a mechanism to recognize and clear apoptotic cells, using the lipids from the engulfed apoptotic cell as peroxisome proliferator-activated receptord transcriptional activators that accelerate apoptotic cell removal and inhibit the inflammatory response. 39 The lipid droplets have been shown to provide a signal for the non-invasive detection of tumor cell death post-treatment using 1 H magnetic resonance spectroscopy (MRS) techniques. 13 The studies presented here have shown that droplet formation accompanies the very earliest events in the apoptotic program and therefore that these 1 H MRS measurements can potentially detect the very earliest signs of cell death.

Materials and Methods
Cell culture. HCT116, EL4, DU145 (ATCC, Teddington, UK) and C4-2b (ViroMed Laboratories, Minnetonka, MN, USA) cell lines were maintained using standard cell culture methods. HCT116 cells were grown in McCoy's 5A supplemented with 10% FBS, MEFs were grown in DMEM supplemented with 10% FBS and all the other cell lines were grown in RPMI medium supplemented with 10% FBS. All cell culture reagents were obtained from Invitrogen (Paisley, UK). Cell number and viability were measured by the Trypan blue dye exclusion assay using a Vi-Cell counter (Beckman Coulter, High Wycombe, UK). Etoposide (Eposin) was obtained from PCH Pharmachemie (Haarlem, The Netherlands). DecylTPP and MitoQ were obtained from Dr. MP Murphy (Cambridge, UK). All other reagents were obtained from Sigma Aldrich (Gillingham, UK), unless specified otherwise.
Assessment of apoptosis and lipid droplet content. Cells (5 Â 10 6 ) were centrifuged and resuspended in 1 ml pre-warmed medium containing 100 nM MTO CMTMRos (Invitrogen, Paisley, UK), incubated for 30 min at 37 1C, centrifuged and then resuspended in ice-cold 3% BSA in HBS (HEPES-buffered saline: 20 mM HEPES, 150 mM NaCl, 2 mM CaCl 2 , pH 7.4) for 30 min at 4 1C. After washing with HBS, cells were incubated in 100 ml HBS containing 5 ml AV-Alexa Fluor 647 (Invitrogen) for 30 min at room temperature. The resulting mixture was then washed three times with HBS and incubated for 20 min at room temperature with 1 ml HBS containing 10 ng/ml Bodipy 493/503 (Invitrogen). Finally, cells were washed twice with HBS and resuspended in 100 ml ice-cold HBS containing 20 mg/ml of 7AAD (Invitrogen). Samples were kept on ice briefly before being loaded into an ImageStream flow cytometer system (Amnis Corporation, Seattle, WA, USA). For each sample, 10 000 events were collected in standard mode. Data files were analyzed using the IDEAS software (Amnis Corporation) to gate the different populations according to their fluorescence staining, to acquire fluorescence images of individual cells and to measure the median intensity of the different dyes. Lipid droplet number (number of spots in the Bodipy (500-560 nm) channel) was obtained by applying the 'Spot Count' feature in the software, which compares the distribution of pixel intensities in an image, providing the number of continuously connected regions.  40 and dissolved in an amount of chloroform proportional to the total cell number. Neutral lipids were resolved by thinlayer chromatography on a silica gel 60 plate (Whatman, Maidstone, UK) using a solvent system containing hexane : diethyl ether : acetic acid (70 : 30 : 1), whereas for polar lipids the solvent system contained chloroform : methanol : acetic acid : water (50 : 37.5 : 3.5 : 2). 18 Phosphor images were obtained using a Typhoon Trio System (GE Healthcare) and densitometric analysis of the different lipid bands was performed with the ImageQuant TL software (GE Healthcare).
Measurement of fatty acid b-oxidation. Cells were pretreated with 100 mM oleate or palmitate dissolved in 10% BSA in PBS for 24 h before experiments; 0.5 mCi/ml [1-14 C]oleate (Perkin Elmer) or 1 mCi/ml [1-14 C]palmitate (GE Healthcare) was added to the culture medium 1 h before cell harvest. Trapping of 14 CO 2 was performed as described in Fu et al. 9 and counts determined using a liquid scintillation counter (Perkin Elmer). The measured radioactivity was normalized to cell number. To measure glucose oxidation, 2 mCi/ml D-[U-14 C]glucose (GE Healthcare) was added to the culture medium 1 h before cell harvest and the same procedure was used to measure 14 CO 2 production.
Microscopy. For the microscopy experiments, 10 000 cells were seeded into each well of a m-slide 8-well plate (Ibidi, Martinsried, Germany) in 300 ml medium. Cells were fixed with 3% paraformaldehyde in PBS and then washed thoroughly with ice-cold PBS. Cells were stained with 0.3 mM DAPI (Invitrogen) for 5 min at room temperature, washed with ice-cold PBS and incubated for 30 min at room temperature with 1 Â HCS LipidTOX Deep Red neutral lipid stain (Invitrogen). Fluorescence images were acquired using a Nikon C1Si confocal microscope (Nikon, Kingston, UK).
The actin gene was used as an internal control to normalize the results of the real-time reverse transcription-PCR. Power SYBR Green PCR Master Mix (Applied Biosystems, Carlsbad, CA, USA) was used to setup the reactions. Amplification and detection were performed on a 7900 HT Fast Real-Time PCR System (Applied Biosystems) using the following cycling conditions: 50 1C for 2 min, 95 1C for 10 min, followed by 40 cycles of 95 1C for 15 s and 60 1C for 1 min. A dissociation stage of 95 1C for 15 s and 60 1C for 15 s was added to check reaction specificity. The Ct values obtained were converted to relative expression values by the comparative Ct method, correcting with the values obtained for actin and normalizing to the untreated control levels.
Western blots. Cell pellets were washed with ice-cold PBS and stored at À80 1C until use. Pellets were solubilized in M-PER Mammalian Protein Extraction Reagent (Thermo Scientific, Cramlington, UK) supplemented with PhosSTOP Phosphatase Inhibitor Cocktail (Roche Applied Science, Burgess Hill, UK) and COMPLETE-mini Protease Inhibitor Cocktail (Roche Applied Science) and sonicated for 5 min. Cell debris was removed by centrifugation. Protein concentrations were determined using QuickStart Bradford Assay (Bio-Rad, Hemel Hempstead, UK). In all, 20mg of protein were loaded into each channel on a 4-20% gel and transferred on to polyvinylidene fluoride membranes (Invitrogen). Membranes were probed with the following antibodies: p53, Akt, phospho-Akt