Acetate functions as an epigenetic metabolite to promote lipid synthesis under hypoxia

Gao, Xue; Lin, Shu-Hai; Ren, Feng; Li, Jin-Tao; Chen, Jia-Jia; Yao, Chuan-Bo; Yang, Hong-Bin; Jiang, Shu-Xia; Yan, Guo-Quan; Wang, Di; Wang, Yi; Liu, Ying; Cai, Zongwei; Xu, Ying-Ying; Chen, Jing; Yu, Wenqiang; Yang, Peng-Yuan; Lei, Qun-Ying

doi:10.1038/ncomms11960

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Published: 30 June 2016

Acetate functions as an epigenetic metabolite to promote lipid synthesis under hypoxia

Xue Gao^1,2^na1,
Shu-Hai Lin^3,4^na1,
Feng Ren¹,
Jin-Tao Li^1,2,
Jia-Jia Chen¹,
Chuan-Bo Yao¹,
Hong-Bin Yang¹,
Shu-Xia Jiang⁴,
Guo-Quan Yan^1,5,
Di Wang^1,2,
Yi Wang¹,
Ying Liu⁶,
Zongwei Cai⁴,
Ying-Ying Xu¹,
Jing Chen⁷,
Wenqiang Yu¹,
Peng-Yuan Yang¹ &
…
Qun-Ying Lei¹

Nature Communications volume 7, Article number: 11960 (2016) Cite this article

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Abstract

Besides the conventional carbon sources, acetyl-CoA has recently been shown to be generated from acetate in various types of cancers, where it promotes lipid synthesis and tumour growth. The underlying mechanism, however, remains largely unknown. We find that acetate induces a hyperacetylated state of histone H3 in hypoxic cells. Acetate predominately activates lipogenic genes ACACA and FASN expression by increasing H3K9, H3K27 and H3K56 acetylation levels at their promoter regions, thus enhancing de novo lipid synthesis, which combines with its function as the metabolic precursor for fatty acid synthesis. Acetyl-CoA synthetases (ACSS1, ACSS2) are involved in this acetate-mediated epigenetic regulation. More importantly, human hepatocellular carcinoma with high ACSS1/2 expression exhibit increased histone H3 acetylation and FASN expression. Taken together, this study demonstrates that acetate, in addition to its ability to induce fatty acid synthesis as an immediate metabolic precursor, also functions as an epigenetic metabolite to promote cancer cell survival under hypoxic stress.

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Introduction

Acetyl-CoA, as a central metabolic intermediate, is widely used in macromolecule biosynthesis and energy production to support cell growth and proliferation. As a donor of acetyl group, acetyl-CoA is also dynamically associated with acetylation modification to modulate protein functions. Therefore, maintenance of cellular acetyl-CoA pool is essential for the regulation of various cellular processes.

In human, acetyl-CoA is mainly produced from oxidation of glucose and other conventional carbon sources, such as glutamine and fatty acids^1,2. However, in human brain cancers, glucose contributes <50% carbons to cellular acetyl-CoA pool, suggesting the existence of a substitutive supply for acetyl-CoA³. Subsequent studies reveal that cancer cells avidly capture acetate as their alternative carbon source to support cell survival and proliferation under stressed conditions, in particular hypoxia^{3,4,5,6,7,8,9}. Moreover, various human cancers show enhanced acetate uptake in [¹¹C]-acetate PET studies^{10,11,12,13,14,15}. These findings suggest that cancer cells utilize acetate as an alternative carbon source to glucose to maintain cellular acetyl-CoA pool under stressed conditions.

Acetate has long been identified as a major carbon source in bacteria and yeasts. Yeast acetyl-CoA synthetases (Acs1p and Acs2p) fuel cell growth by converting acetate to acetyl-CoA¹⁶. Very recently, acetate is also found to be an alternative carbon source besides glucose, glutamine and fatty acids in human cancer, attracting intensive investigations^7,8. Generally, mammalian acetyl-CoA synthesis from acetate is carried out by ACSS2 to support lipid synthesis in the cytosol, and by ACSS1 to fuel ATP production in mitochondria^17,18,19. Acetate is mainly acquired from diet, but can also be generated in ethanol metabolism or deacetylation processes. The function of acetate has long been overlooked due to its relative low physiological concentration (0.2–0.3 mM) in blood²⁰. Recent studies reveal that cancer cells show increased acetate uptake under hypoxia even in the presence of low acetate concentration to support tumour growth^7,9,14. However, how cancer cells utilize acetate under hypoxia in such an efficient manner remains unclear.

Histone acetylation is intimately coordinated with cellular acetyl-CoA pool in response to metabolic state. As the downstream metabolite of carbon sources, acetyl-CoA represents a pivotal metabolic signal of nutrient availability^4,21,22. In yeast, histone is specifically acetylated at genes involved in lipogenesis, aminoacid biosynthesis and cell cycle progression upon entry into growth, in tune with intracellular acetyl-CoA level²³. ATP citrate lyase (ACLY), the enzyme converting glucose-derived citrate into acetyl-CoA, regulates histone acetylation by sensing glucose availability^1,22. Yeast acetyl-CoA Carboxylase (Acc1p) consumes acetyl-CoA to synthesize lipids and regulates global histone acetylation through competing for the same nucleocytosolic acetyl-CoA pool²⁴. Thus, the acetyl-CoA flux dynamically regulates gene expression profile by modulating histone acetylation state.

These observations led us to hypothesize that acetate induces a metabolic adaptation through modulating histone acetylation in hypoxic cancer cells. Consistent with this idea, we found that acetate predominately activates the expression of lipogenic genes through upregulating histone acetylation at their promoter regions, which in turn promotes lipid synthesis under hypoxia. Beyond a carbon source for macromolecular biosynthesis, our findings highlight an epigenetic role for acetate in metabolic adaptation of cancer cells to hypoxic stress.

Results

Acetate restores histone acetylation under hypoxia

Cancer cells demand distinctive extracellular nutrients and reprogram the metabolic pathways to survive and proliferate when facing harsh situation, such as hypoxia²⁵. Hypoxia stress is a critical player in tumorigenesis and tumour development²⁶. By performing exometabolome analysis based on ¹H-NMR spectra, we found that cancer cells absorbed around 20% acetate from the culture medium under normoxia while more than 80% acetate was consumed under hypoxia (Fig. 1a; Supplementary Fig. 1a,b), suggesting that cancer cells take up more acetate under hypoxia than nomoxia^7,9,14. Moreover, we carried out the quantification of acetate from five pairs of hepatocellular carcinoma (HCC) and adjacent samples by NMR. As shown in Supplementary Fig. 1c, acetate concentration range was from 0.56 to 2.67 μmol g⁻¹ in wet tissue (left) and acetate concentration in tissue would be roughly estimated around from 0.56 to 2.67 mM (right). In most cases acetate concentrations ranged around 0.5 mM and in two HCC samples acetate levels reached 2.5 mM (Supplementary Fig. 1c). To explore acetate effect on cancer cell, we treated HepG2 cells with acetate under hypoxia and found that acetate counteracted the declined histone acetylation under hypoxia compared with normoxia (Fig. 1b). Of particular interest, acetate induced significant increase of H3K9, H3K27 and H3K56 acetylation levels, but not H3K14, H3K18, H3K23 and H3K36 acetylation levels (Fig. 1b). This indicates that acetate rescues hypoxia-reduced histone acetylation with certain specificity. More importantly, by using metabolic-labelling technique based on liquid chromatography tandem mass spectrometry in multiple reaction monitoring mode (LC-MRM MS), we identified ¹³C₂-labelled acetylated H3K9 (Supplementary Fig. 1d), H3K27 (Supplementary Fig. 1e) and H3K56 (Supplementary Fig. 1f) peptides in HepG2 cells treated with [U-¹³C₂]-acetate, demonstrating that acetate-derived acetyl-CoA was indeed incorporated into histones. In addition, compared with normoxia, histone acetylation is more susceptible to acetate supplementation under hypoxia, even under low concentration, in HepG2, A549 and DU 145 cells (Fig. 1c; Supplementary Fig. 1g,h).

**Figure 1: Acetate rescues hypoxia-induced reduction of histone acetylation.**

To characterize the manner of epigenetic regulation by acetate, we treated cancer cells with acetate for different time points under hypoxia. We found that exogenous acetate supplement rapidly increased H3K9, H3K27 and H3K56 acetylation levels in a time-dependent manner under hypoxia in different cancer cell lines (Fig. 1d; Supplementary Fig. 1i,j). Acetate supplement dramatically increased H3K9, H3K27 and H3K56 acetylation levels within 2 h (Fig. 1d; Supplementary Fig. 1i,j). Furthermore, acetate was found to increase histone H3K9, H3K27 and H3K56 acetylation levels in a dose-dependent manner under hypoxia in HepG2 (Fig. 1e), A549 (Supplementary Fig. 1k), DU145 (Supplementary Fig. 1l), SkBr3 (Supplementary Fig. 1m) and HT29 cells (Supplementary Fig. 1n). Low concentration of acetate (1.25 mM) slightly increased histone acetylation levels, while higher concentrations of acetate (2.5–5 mM) significantly increased histone acetylation levels under hypoxia (Fig. 1e; Supplementary Fig. 1k, l, m, n). Collectively, these data imply that acetate is implicated in the regulation of histone acetylation under hypoxia.

Acetate epigenetically activates de novo lipogenesis

It is well known that histone acetylation is associated with transcription activity^27,28,29. To investigate the biological effect on cancer metabolism induced by the acetate-mediated histone acetylation, we detected the mRNA expression changes of metabolic genes. We designed quantitative PCR (qPCR) primers targeting 139 metabolic genes, covering a wide range of metabolic pathways (that is, glycolysis, TCA cycle, pentose phosphate pathway, glycogen metabolism, ketone body metabolism, aminoacid metabolism, fatty acid synthesis and β-oxidation, cholesterol synthesis, sphingolipid metabolism, and metabolism-related transporters) (Supplementary Table 1) and conducted qPCR to screen potential metabolic pathway(s) affected by acetate-induced epigenetic regulation. Scatter plot of mRNA expression data of 139 genes in HepG2 cells was shown in Fig. 2a, by comparing cell culture under hypoxia with normoxia. The expression of VEGF and LDHA, which were included as positive controls for verifying hypoxic effect, increased by more than two-fold under hypoxia (Fig. 2a). We then tested the effect of acetate on gene expression under either normoxia (Fig. 2b) or hypoxia (Fig. 2c), respectively. Compared with the normoxia group, the upregulated genes by two-fold in hypoxia group and hypoxia with acetate group showed a lot overlap (Supplementary Fig. 2a). Besides, FASN and ACSS2 were the unique genes in hypoxia with acetate group, indicating that acetate played an important role in lipid synthesis under hypoxia (Supplementary Fig. 2a). In line with this, FASN and ACACA mRNA levels were activated by more than two-fold with acetate under hypoxia, while no activation effect was observed under normoxia (Fig. 2b,c). Both FASN and ACACA are in the top-10 list of upregulated metabolic genes induced by acetate under hypoxia, indicating that acetate predominantly affects de novo lipid synthesis compared with other metabolic pathways (Fig. 2c,d). Interestingly, ACSS1 and ACSS2 mRNA levels were decreased under hypoxia compared with normoxia (Fig. 2a), which was discrepant with the other studies reported that ACSS2 was upregulated under hypoxia^9,14. This may be due to the experimental conditions, such as different cell lines used and treating time points. Moreover, mRNA level of ACSS1 and ACSS2, rather than ACLY, were also upregulated on acetate treatment under hypoxia (Fig. 2c,d). The rescue of hypoxia-reduced ACSS1 and ACSS2 expression by acetate may indicate that ACSS1 and ACSS2 play an important role in cancer cells adapting to hypoxia.

**Figure 2: Acetate predominately activates lipid synthesis pathway through epigenetic regulation under hypoxia.**

We further investigated whether acetate-induced histone acetylation was associated with acetate-promoted gene transcription on lipogenesis. We treated cancer cells with gradient concentrations of acetate and found that FASN and ACACA mRNA expression were upregulated in a dose-dependent manner under hypoxia (Fig. 2e,f). Intriguingly, change in FASN and ACACA mRNA expression are more prone to low concentration of acetate under hypoxia, compared with normoxic control (Fig. 2e,f). To characterize whether the upregulation of lipogenic genes is caused by histone acetylation induced by acetate supplementation, we treated cancer cells with trichostatin A (TSA), a histone deacetylase inhibitor, as a positive control. We found that TSA and acetate increased both the global acetylation level of histone H3 and the mRNA expression of FASN and ACACA, although TSA and acetate had no synergic effect (Supplementary Fig. 2b). In addition, acetate treatment led to time-dependent enhancement of H3K9, H3K27 and H3K56 acetylation levels (Supplementary Fig. 2c). In line with the increased histone H3 acetylation, FASN and ACACA mRNA expressions were also elevated in a time-dependent manner (Supplementary Fig. 2c). Furthermore, we carried out ChIP–qPCR assays to define the underlying mechanism of acetate-induced FASN and ACACA expressions. We found that acetate markedly enhanced H3K56 acetylation level at the promoters of FASN and ACACA (Supplementary Fig. 2d,e), but not ACLY (Supplementary Fig. 2f). Furthermore, we found that histone acetylation (H3K9ac, H3K27ac and H3K56ac) at the promoters of FASN and ACACA responded to acetate treatment in a dose-dependent manner and histone acetylation was more prone to be induced by low concentration (≤2.5 mM) of acetate under hypoxia, compared with normoxia (Fig. 2g,h). These results demonstrate that acetate is capable of epigenetically regulating lipogenic genes at pathophysiological concentration.

To rule out the possibility that promoter acetylation of FASN and ACACA reflects increased transcription, we alternatively performed ChIP–qPCR and tested promoter histone acetylation of LDHA and VEGF (Supplementary Fig. 2g, h, i, j). The acetylation levels (H3K9ac, H3K27ac and H3K56ac) at LDHA and VEGF promoters were increased under hypoxia, which was further upregulated by acetate supplementation (Supplementary Fig. 2g, h, i, j). Consistently, mRNA expression of LDHA and VEGF was also activated by acetate (Supplementary Fig. 2k,l). These observations suggest that acetate contributes to histone acetylation at the promoters of hypoxia-induced genes. Collectively, these data support our notion that acetate promotes lipid synthesis pathway through epigenetic regulation.

Acetate-induced lipogenesis does not reflect lipid demands

Cancer cells show increased demands for lipids, through scavenging extracellular lipids or de novo lipid synthesis^2,30,31. We tried to figure out whether acetate-induced expression of FASN or ACACA is affected by the extracellular lipids under hypoxia. Consistent with data in Fig. 2, FASN and ACACA mRNA expression was more prone to be induced by acetate under hypoxia, compared with normoxia (Fig. 3a,c) and acetate-activated promoter histone acetylation of FASN and ACACA more pronouncedly under hypoxia (Fig. 3b,d). Furthermore, lipid depletion in the serum did not alter acetate-induced expression of FASN and ACACA in HepG2 cells (Fig. 3e; Supplementary Fig. 3a). Similarly, FASN and ACACA promoter histone acetylation was not affected by lipid depletion as well (Fig. 3f; Supplementary Fig. 3b). Moreover, palmitate supplementation did not change the expression of FASN, VEGF and ACACA (Fig. 3g; Supplementary Fig. 3c,d). Consistently, histone acetylation at the promoter regions of FASN and ACACA did not respond to palmitate supplementation (Fig. 3h; Supplementary Fig. 3e). These results demonstrate that acetate increases histone acetylation at promoters of lipogenic genes and activates their mRNA expression, without reflecting lipid demand of hypoxic cells.

**Figure 3: Acetate epigenetically activates lipogenic genes without reflecting cellular lipid demands.**

ACSS1 and ACSS2 are both involved in acetate-induced lipogenesis

In mammals, two main enzymes involve in acetyl-CoA production from acetate: the cytosolic ACSS2 and its mitochondrial paralogue ACSS1. To dissect which enzyme is responsible for mediating the acetate-induced increase of histone acetylation, we knocked down ACSS1 or ACSS2 individually and assessed the corresponding histone acetylation levels. To our surprise, knockdown of either ACSS1 or ACSS2 was incapable of blocking acetate-induced increase in acetylation levels at H3K9, H3K27 and H3K56 sites (Supplementary Fig. 4a,b). Though discrepant with the other studies reporting that ACSS2, but not ACSS1, appeared to be the key enzyme involved in metabolizing acetate for lipid synthesis^5,9, our findings were consistent with those by Yun et al.³² and Bjornson et al.³³, which reported that ACSS1 was also vital for acetate-dependent lipid synthesis. These results indicate that ACSS1 and ACSS2 are functionally redundant. We then established ACSS1/2 double knockdown stable cell pools. Concurrent knockdown of ACSS1/2 by using two different sets of shRNAs dramatically inhibited acetate-induced increase in acetylation levels of H3K9, H3K27 and H3K56 under hypoxia (Fig. 4a; Supplementary Fig. 4c). Moreover, we conducted quantification based on LC-MRM MS to see whether ACSS1/2 double knockdown could reduce the abundance of histone acetylation peptides derived from acetate by using [U-¹³C₂]-acetate tracer. Acetate-derived acetyl-CoA was found actively incorporated into H3K9, H3K14 and H3K27 acetylation sites, compared with H3K18, H3K23 and H3K56 acetylation sites (Fig. 4b; Supplementary Fig. 4d). These results demonstrate that acetate could function as an acetyl donor to induce histone acetylation. ACSS1/2 double knockdown induced a more significant decrease in ¹³C₂-labelled acetylated H3K9, H3K27 peptides derived from [U-¹³C₂]-acetate (Fig. 4b; Supplementary Fig. 4d). As for the discrepancy of H3K56 acetylation between western blot data and MS quantification, it might be due to the fact that concentration of ¹³C₂-labelled acetylated H3K56 peptide was too low to meet the limit of quantification. In addition, we designed experiments to test whether ACSS1/2 double knockdown could block acetate-induced histone acetylation at FASN and ACACA promoter regions. ACSS1/2 knockdown efficiency was verified by qPCR (Supplementary Fig. 4e). We found that acetate-increased H3K9, H3K27 and H3K56 acetylation levels at FASN (Fig. 4c) and ACACA (Fig. 4d) promoter regions were blocked by ACSS1/2 double knockdown under hypoxia. ACSS1/2 knockdown had no effect on histone acetylation levels at ACLY promoter region (Fig. 4e). Consistently, ACSS1/2 knockdown blocked acetate-induced FASN expression (Supplementary Fig. 4f). Collectively, our data demonstrate that acetate contributes to histone acetylation in an ACSS1/2-dependent manner under hypoxia.

**Figure 4: Both ACSS1 and ACSS2 are involved in acetate-induced epigenetic regulation of *de novo* lipogenesis.**

Previous study linked ACLY-dependent glucose metabolism to histone acetylation^1,22. To dissect contributions of different carbon sources to histone acetylation, we performed qPCR and ChIP–qPCR assays in ACSS1/2-knockdown and/or ACLY-knockdown cells. The knockdown efficiency was verified (Supplementary Fig. 4g). Depletion of either ACSS1/2 or ACLY reduced gene expression of LDHA and VEGF, which were further decreased by knockdown of both ACSS1/2 and ACLY (Supplementary Fig. 4h,i). Consistently, either ACSS1/2 or ACLY knockdown reduced histone acetylation at the promoter regions of LDHA and VEGF (Supplementary Fig. 4j, k, l, m). These results demonstrate that both ACSS1/2 and ACLY contribute to histone acetylation under hypoxia.

Acetate-induced lipogenesis promotes cell survival

To test whether acetate would affect lipid pool in cancer cells, we cultured HepG2 stable cells in media with isotope-labelled [U-¹³C₂]-acetate for 24 h under hypoxia and measured the isotopologues of fatty acids pool by using gas chromatography coupled with mass spectrometry (GC-MS). ACSS1/2 double knockdown indeed decreased the fractional contribution of acetate to palmitate and stearate synthesis (Fig. 5a). These results are consistent with the model that acetyl-CoA derived from acetate stimulates de novo lipid synthesis^5,7,9. Furthermore, we knocked down FASN expression to the unstimulated state by using the low-efficiency siRNA as in Supplementary Fig. 5a, and treated cells with isotope-labelled [U-¹³C₂]-acetate under hypoxia. The de novo lipid synthesis from acetate-derived carbon was reduced in FASN knockdown cells, demonstrating that acetate functions as an acetyl donor to promote lipogenesis and acetate-induced FASN expression plays an important role in lipogenesis (Fig. 5b). Given that elevated lipid synthesis could promote tumour cell survival under hypoxia^2,25, we carried out CCK8 assay and found that acetate supplement enhanced cell survival under hypoxia, but not normoxia (Fig. 5c). To interrogate whether acetate promoted cancer cell survival through regulating lipogenic genes, we employed specific siRNAs targeting FASN or ACACA, of which the relative low knockdown efficiencies are capable of suppressing acetate-simulated FASN and ACACA mRNA expression to the level similar to unstimulated state (Supplementary Fig. 5a). We found that acetate-induced cell survival under hypoxia was inhibited by knockdown of either FASN or ACACA, indicating that acetate-promoted cell survival was dependent on lipogenesis and the induction of lipogenic gene expression was vital for cell survival (Fig. 5d). Strikingly, we found that inhibition of either FASN or ACACA to the untreated level had no effect on acetate-induced FASN promoter histone acetylation (Supplementary Fig. 5b). We attempted to further dissect whether acetate-induced de novo lipid synthesis is vital for cancer cell cultured under hypoxia. We found that HepG2 and SkBr3 cells were more sensitive to C75, a chemical inhibitor of FASN, when cultured under hypoxia (Supplementary Fig. 5c), consistent with previous report that cancer cells were dependent on de novo lipid synthesis under hypoxia⁹. Moreover, C75 treatment abrogated acetate-induced cancer cell survival without affecting FASN and ACACA mRNA levels (Supplementary Fig. 5d,e,f). Consistently, ACSS1/2 knockdown blocked the increased cell survival induced by acetate supplement, indicating that acetate-induced cell survival was ACSS1/2-dependent (Fig. 5e). In addition, we performed Matrigel assay and found that HepG2 cells supplied with acetate exhibited growth advantage over the control group, and this effect was blocked by ACSS1/2 double knockdown (Fig. 5f). Taken together, beyond functioning as a precursor for lipogenesis, acetate also serves as acetyl donor to induce histone acetylation and mRNA expression of lipogenic genes, contributing to the adaptation of cancer cells to hypoxia.

**Figure 5: Epigenetic regulation of *de novo* lipogenesis by acetate is vital for cell survival under hypoxia.**

ACSS1/2 positively correlates with FASN expression in HCC

ACSS2 is reported to be upregulated in various human cancers^3,5,9. ACSS1 is also reported to be increased in hepatocellular carcinoma³³. Through an analysis of 190 human HCC samples data from the Cancer Genome Atlas (TCGA)³⁴, we found a significant correlation between FASN and ACSS2 mRNA levels and no significant correlation between FASN and ACSS1 mRNA levels (Supplementary Fig. 6a). Our aforementioned results demonstrated that acetate could epigenetically regulate FASN expression, which prompts us to examine ACSS1/2, FASN, and H3 acetylation levels and their correlation in human HCC. We collected 53 pairs of primary human HCC samples with adjacent normal tissues, and determined protein changes (ACSS1, ACSS2 and FASN) and six histone acetylation marks (H3K9ac, H3K14ac, H3K18ac, H3K23ac, H3K27ac and H3K56ac) in all these samples (n=53) (Fig. 6a; Supplementary Fig. 6b). To perform a statistical analysis of the association between ACSS1/2 and histone acetylation, we divided tumour samples into two groups based on ACSS1/2 signature (high versus low). Tumours with 1.5-fold higher expression of ACSS1 or ACSS2 or both than that of adjacent normal control tissue are grouped into ACSS high-expression tumours (tumour/normal ≥1.5, n=26), while ACSS low-expression tumours (tumour/normal <1.5, n=27) express both ACSS proteins at 1.5-fold lower than its normal control (Fig. 6b). We found 16 cases with ACSS1 high expression, 8 cases with ACSS2 high expression and 2 cases with both ACSS1 and ACSS2 high expression (Fig. 6b). The higher percentage of ACSS1 high-expression cases than that of ACSS2 fully proves the importance of ACSS1 in acetate utilization by cancer. Our results demonstrate that four out of six histone acetylation marks, including H3K9ac (P=0.0220), H3K14ac (P=0.0289), H3K27ac (P=0.0034) and H3K56ac (P=0.0410), are significantly stronger in ACSS high-expression tumours than that from ACSS low-expression tumours with two-tailed unpaired Student’s t-test (Fig. 6c). In line with this observation, FASN protein level is also significantly upregulated in ACSS high-expression tumours (Fig. 6d). Consistently, we performed immunohistochemistry (IHC) analysis and found that ACSS1/2 expression was positively correlated with H3 acetylation and FASN expression in human HCC. A representative case was shown in Fig. 6e. By using Spearman correlation analysis, we have evaluated the relationship between histone acetylation and FASN expression. Surprisingly, each and every histone acetylation mark in our analysis exhibits significant positive correlation with FASN expression (P<0.0001 for H3K9ac, P=0.0003 for H3K14ac, P=0.0009 for H3K18ac, P=0.0385 for H3K23ac, P<0.0001 for H3K27ac and P<0.0001 for H3K56ac; Supplementary Fig. 6c). Collectively, these results strongly support a positive correlation between ACSS1/2, FASN and histone acetylation in human HCC.

**Figure 6: ACSS1/2 positively correlates with histone acetylation and FASN expression in human hepatocellular carcinoma.**

Discussion

Acetyl-CoA is a central metabolite derived mainly from glucose, glutamine and fatty acids in mammals. However, the capability of cancer cells to produce acetyl-CoA from these conventional carbon sources is dramatically decreased under hypoxia^3,7. Cancer cells, therefore, need alternative carbon sources and conduct metabolic reprogramming. Clinical studies report the increase in [¹¹C]-acetate uptake in multiple types of cancers including prostate, liver, lung and brain tumours. Recently, new studies further demonstrate that various tumours consume exogenous acetate to generate acetyl-CoA for lipid synthesis^3,7,9. These studies support that acetate serves as an important carbon source for cancer cells under unfavourable conditions. The underlying mechanism of how acetate functions as carbon source to fuel tumour growth, however, remains as an open question.

In this study, we elucidate that acetate functions as an epigenetic regulating metabolite to enhance lipid synthesis and to promote tumour survival under hypoxia. First, histone acetylation at H3K9, H3K27 and H3K56 sites can be stimulated by acetate in both time- and dose-dependent manners under hypoxia in mutiple cancer cell lines. Notably, acetate-derived acetyl-CoA is found to be incorporated not only into fatty acid synthesis but also into acetylated histone peptides. Second, acetate-induced histone acetylation is then associated with FASN and ACACA promoter regions, which activates FASN and ACACA expressions under hypoxia. Third, acetate-driven epigenetic regulation is mediated by ACSS1 and ACSS2. Last and most important, acetate-induced lipogenic genes expression is found to promote de novo lipid synthesis and cell survival under hypoxic stress. Collectively, besides its ability to induce fatty acid synthesis as an immediate metabolic precursor, our study reveals a new mechanism of epigenetic regulation by acetate to increase lipid synthesis and promote cell survival under unfavourable conditions (Fig. 7). However, we assess effects of acetate using limited set of cell lines in vitro, it needs further confirmation of the findings using more cell lines. In addition, the experiments in vivo would provide more validation.

**Figure 7: The model for acetate-induced epigenetic regulation of *de novo* lipogenesis.**

Alterations in histone modifications play important roles in transcriptional regulation in cancer cells³⁵. H3K9ac, H3K27ac and H3K56ac, three epigenetic marks of active gene transcription, are found to be altered in various cancers. H3K9 hyperacetylation is associated with specific genes in breast cancer³⁶. Similarly, H3K27 acetylation is upregulated in colorectal cancer³⁷. Intensive investigations also show that H3K56 acetylation is closely related with epigenetic activation and is increased in multiple types of cancer^{38,39,40,41,42,43}. In our work, we found that H3K9, H3K27 and H3K56 acetylation levels were increased in ACSS1/2-overexpressed human HCC, showing a significantly positive correlation between ACSS1/2 expression, histone acetylation and FASN expression. ACSS1/2 upregulation suggests that these cancer cells utilize acetate as both epigenetic metabolite and carbon source to meet their growth needs. These findings validate our model in vivo: ACSS1/2 links histone acetylation with FASN expression and acetate-induced epigenetic regulation may play a pivotal role in human HCC. As the major enzymes in acetate metabolism, ACSS1 and ACSS2 may merit further explorations as a therapeutic target for cancer.

Dysregulation of de novo lipid synthesis is a hallmark of cancer⁴⁴. Increased lipid synthesis fuels cancer engine and contributes to cancer cells survival when facing with stressed conditions, such as hypoxia^{25,30,44,45,46,47,48,49}. Upregulation of FASN is common in many cancers and FASN is targeted for cancer therapy^44,49,50,51. FASN inhibition with chemical inhibitors or RNAi can suppress tumour cell survival⁵¹. FASN upregulation in cancers is in part due to the transcriptional activation by SREBPs^44,45,52,53. In addition, USP2a was reported to enhance FASN protein stabilization⁴⁴. Our study proves that acetate increases histone acetylation at promoters of lipogenic genes and activates their mRNA expression, without reflecting lipid demand of hypoxic cells. Interestingly, our data show that both ACSS1/2-mediated acetate and ACLY-mediated glucose contribute to histone acetylation at the promoters of hypoxia-induced genes. VEGF or LDHA shows a similar response that occurs in hypoxia alone without acetate addition yet each remains responsive to acetate. This suggests the changes occurring at these promoters that lead to gene activation (presumably recruitment of co-activators) render them more sensitive to acetyl-CoA levels compared with other genes. How recruitment of co-activators modulates acetate-induced lipogenic genes expression is an interesting question.

In conclusion, our study adds a novel mechanism of epigenetic regulation to FASN upregulation. Besides glucose, cancer cells absorb more acetate under hypoxia. Besides functioning as an immediate metabolic precursor, acetate-derived acetyl-CoA increases H3K9, H3K27 and H3K56 acetylation levels at FASN and ACACA promoter regions, initiating epigenetic regulation of FASN and ACACA. Due to the limitation of our system in distinguishing the role of acetyl-CoA as a substrate rather than an epigenetic regulator, both pathways cannot be separated well and their relative contribution remains unclear. Histone acetylation modulated by acetate status might play a pivotal role in coordinating acetate and glucose availability with the intracellular level of FASN, hence fatty acid synthesis. Taken together, this study demonstrates that acetate, in addition to its ability to induce fatty acid synthesis as an immediate metabolic precursor, also functions as an epigenetic metabolite to promote cancer cell survival under hypoxic stress.

Methods

Cell culture

PLC-8024 cells were obtained from the Institute of Virology, Chinese Academy of Medical Sciences (Beijing, China). All the other cell lines were obtained from the Cell Bank of Type Culture Collection of Chinese Academy. HepG2, SkBr3, PLC-8024 and DU 145 cells were grown in DMEM medium (Gibco, high glucose) with 10% fetal bovine serum (FBS, biological industries (BI)), and penicillin/streptomycin. A549 cells were grown in F12K medium (Sigma) with 10% FBS (BI), and penicillin/streptomycin. HT29 cells were cultured in McCoy’s 5A medium (Sigma) with 10% FBS (BI), and penicillin/streptomycin. Mycoplasma contamination check was carried out on cultured cells. Hypoxia condition (1% O₂, 94% N₂, 5% CO₂) was achieved by gas chamber (C42, Biospherix).

Antibodies and reagents

Anti-histone H3 (9715S, CST), anti-H3K9 acetyl (1328-1, Epitomics), anti-H3K27 acetyl (EP865Y, Epitomics), anti-H3K56 acetyl (EPR996Y, Epitomics), anti-ACSS2 (ab133664, Abcam), anti-ACSS1 antibody (17138-1-AP, Proteintech), anti-FASN (C-20140, Santa Cruz), anti-ACLY antibody (1699-1, Epitomics) and anti-β-actin (a00702, Genescript) antibodies were commercially obtained. ChIP grade antibodies for H3K9 acetyl (ab4441, Abcam), H3K27 acetyl (ab4729, Abcam), H3K56 acetyl (17-10259, Milipore) were commercially obtained.

[U-¹³C₂]-Acetate (CLM-440-1, Cambridge Isotope Laboratories), lipoprotein depleted fetal bovine serum (LPDS) (880100, Kalen Biomedical), sodium palmitate (P9767, Sigma), BSA with low fatty acid (0219989925, MP) and TSA (9950, CST) were commercially obtained. Myristic acid (D27, 98%) as internal standard was obtained from Cambridge Isotope Laboratories (Tewksbury, MA, USA). Certified ACS grade sodium hydroxide was purchased from Standard Chemical and hydrochloric acid was from VWR International LLC (Pennsylvania, USA). Formic acid and BSTFA (with 1% TMCS) were purchased from Sigma-Aldrich. Methanol and chloroform were obtained from Tedia (Fairfield, USA) and hexane obtained from Duksan Pure Chemicals Co., Ltd (South Korea).

Western blotting

Standard western blot protocols were adopted. The quantification was carried out by subtracting background from the band intensity of western blots by using software ImageQuantTL (GE).

LC-MRM MS quantification of histone acetylation

Histone proteins were extracted and purified as previously described by Gao et al⁵⁴. Briefly, 3 × 10⁶ scramble (shscr) or shACSS1/2 HepG2 cells were cultured in DMEM medium (without glucose or glutamine, Sigma) by adding 10% dialysed FBS (BI), 2.5 mM [U-¹³C₂]-acetate, 5 mM label-free glucose, 2 mM label-free glutamine for 4 h under hypoxia (1% O₂), followed by lysis with hypotonic lysis buffer (10 mM Tris-HCl (pH 8.0), 1 mM KCl, 1.5 mM MgCl₂, 1 mM DTT and protease inhibitors) to isolate nuclei. Crude histone was extracted with 0.4 N HCl overnight. After removal of HCl by ultrafiltration, the crude histone was processed to derivatization in a final condition of 100 mM NHS-propionylate ester (home-made), 25 mM NH₄HCO₃, 50% ACN, 50 °C for 30 min. The proteins were SpeedVac to dryness and digested with trypsin in 25 mM NH₄HCO₃ overnight. Finally, the digested peptides were concentrated to dryness again and derivatized for a second round of histone NHS propionylation.

Stable isotope-labelled histone peptides containing various PTMs and chemical derivatization (purity >95%) were from New England Peptide LLC (Gardner, MA, USA). Prior to LC-MS analysis, internal standards were mixed with the diluted derivatized histone samples in a ratio of 1:2 in volume. In the final solution, the internal standard was doped as 25 nM stable isotope-labelled histone peptides, while the final concentration of stable isotope-labelled peptides with K27 acetylation was 2.5 nM. The peptides were first trapped to a nanoLC trap column at 2 ml min⁻¹ liquid flow that was diverted from the analytical column via a vent valve, while separation was performed by switching the valve to make the trap column in line with a C18 analytical column with a flow rate of 300 nl min⁻¹. The chromatographic gradient was 5–35% B from 0 to 45 min, 35–80% B from 45 to 60 min, and maintaining 80% B for 4 mins (mobile phase A: 0.1% formic acid in ddH₂O; mobile phase B: 0.1% formic acid in acetonitrile). Then, the peptides were directly analysed online using QTRAP 6500 mass spectrometer (Applied Biosystems, Singapore) set to multiple reaction monitoring (MRM) in positive electrospray ionization mode. The mass transitions were selected as previously described by Chen et al^54,55, with the addition of transitions corresponding to ¹³C₂-labelled acetylated peptides. All MRM data were processed through Skyline software (Washington University in St Louis, St Louis, MO, USA). The quantification was based on the peak area and the measured areas under the curve of indicated product ions were calculated. All data were manually inspected to ensure correct peak detection and accurate integration according to four criteria: (1) the correct m/z is selected for both the heavy and light traces of each peptide, and [U-¹³C₂]-acetylation for acetylated peptides; (2) the peak shapes are Gaussian-like and do not show excessively jagged appearance; (3) the retention time of certain peptide is similar between different runs; and (4) the relative contribution of each transition to the total signal is similar between each sample.

RNA extraction and qPCR

Total RNA was extracted with Trizol extraction kit (Invitrogen). qPCR was performed using SYBR Premix Ex Taq (TakaRa) in an ABI 7500 Sequence Detection System (Applied Biosystems). All reactions were performed in triplicate and relative mRNA expression was normalized to β-actin. All tested primers are listed in Supplementary Table 1.

ChIP–qPCR assays

ChIP–qPCR assays were performed as previously described⁵⁶. Briefly, 1 × 10⁶ HepG2 cells were cross-linked with 1% paraformaldehyde, lysed and sonicated using the Bioruptor at high-output power setting for eight cycles (30 s ON and 30 s OFF). Solubilized chromatin was immunoprecipitated with chip grade antibodies for H3K9 acetyl, H3K27 acetyl, H3K56 acetyl or rabbit IgG (negative control), following that the antibodies were preincubated with protein protein A sepharose overnight at 4 °C. Antibody–chromatin complexes were pulled down by protein A sepharose (Santa Cruz), washed and then eluted. After crosslink reversal in a water bath at 65 °C overnight and proteinase K treatment for 2 h at 55 °C, the immunoprecipitated DNA was extracted with phenol–chloroform, and ethanol precipitated. The DNA fragments were detected by qPCR. Histone acetylation marks were mapped at promoters spanning −3 to 3 kb of target genes (ACACA, FASN, ACLY, VEGF and LDHA). Primers spanning the regions with peaks were adopted for ChIP–qPCR analysis. All tested primers targeting FASN, ACACA, ACLY, LDHA and VEGF are listed in Supplementary Table 1.

RNA interference

For RNA interference experiments, si ACSS1 #1 (EHU040311) and si ACSS2 #1 (EHU135341) were purchased from Sigma. The others were obtained from commercial synthetic siRNA oligonucleotide (Genepharma, Shanghai).

siACSS1 #2: 5′-UCACCGUAUUUCAGCAACAGCCGG-3′;

siACSS2 #2: 5′-UAUGCUUGGUGACAGGCUCAUCUCC-3′;

siACLY #1: 5′-GAUCAAACGUCGUGGAAAAUU-3′;

siACLY #2: 5′-GAGGAAGCUGAUGAAUAU-3′;

siFASN #1: 5′-UGGAGCGUAUCUGUGAGAAUU-3′;

siFASN #2: 5′-AACCCUGAGAUCCCAGCGCUG-3′;

siACACA #1: 5′- CAAUGGCAUUGCAGCAGUGUU-3′;

siACACA #2: 5′-UAUGAGGUGGAUCGGAGAUUUUU-3′.

All siRNA transfection experiments were performed with Lipofectamine RNAiMAX Transfection Reagent (Invitrogen) according to the reverse transfection instructions. A volume of 5 μl siRNA (10 μM) plus with 5 μl Lipofectamine RNAiMAX per well (six wells) was used. The cell density was 8 × 10⁵ HepG2 cells per well (six wells). The fresh culture media was exchanged after 24 h. The samples with indicated treatment were collected at 72 h after transfection. The knockdown efficiency were detected by western blot or qPCR.

Generation of stable knockdown cell lines

The retroviral knockdown constructs of shACSS1 were constructed in pMKO.1-hyg vector as previously described⁵⁷. The lentiviral knockdown constructs of shACSS2 were constructed in pLKO.1-puro vector. The targeting sequences were as follows:

shACSS1#1: 5′-TCACCGTATTTCAGCAACAGCCGG-3′;

shACSS1#2: 5′-AGGTGGTTATCACCTTCAA-3′;

shACSS2#1: 5′-TACAATGTACTGGATCGAA-3′;

shACSS2#2: 5′-AACGCTTTGAGACAACCTA-3′;

The virus was packaged in HEK293T cells by co-transfecting pLKO.1-shACSS2 or pLKO.1-shACSS2 or corresponding vectors (pLKO.1-shscr and pLKO.1-shscr) with package vectors expressing gag and vsvg genes (from vesicular stomatitis virus G). The virus in the culture media was harvested every 24 h for 48 h after initial plasmid transfection. For infection, HepG2 cells were infected for 12 h with pMKO.1-shACSS1 and pMKO-shscr retrovirus with 8 μg ml⁻¹ polybrene to increase the infection efficiency and selected in 200 μg ml⁻¹ hygromycin B for 1 week first. Then the stable shACSS1 or shscr HepG2 cells were infected with pLKO.1-shACSS2 and pLKO.1-shscr retrovirus and selected in 1 μg ml⁻¹ puromycin for 1 week to obtain ACSS1 and 2 double knockdown (shACSS1/2) stable cell lines. The knockdown efficiency was verified by western blot.

NMR spectroscopy

To detect acetate uptake in cancer cells, the cancer cells were grown in DMEM (Gibco, high glucose) supplemented with 10% FBS (BI) and penicillin/streptomycin. A volume of 200 μl of each medium was transferred into a 1.5-ml Eppendorf tube, then 600 μl of cold CD₃OD with 0.09 mg ml⁻¹ trimethylsilyl propionate (TSP) was added to the sample and vortexed for 30 s. The mixture was centrifuged at 20,817g for 10 min at 4 °C. Then 600 μl each supernatant was transferred to a standard 5-mm NMR tube for analysis.

To quantitatively analyse acetate in HCC tissues and adjacent normal tissue, 150 mg was weighted for each wet tissue. By adding 700 μl methanol and 280 μl H₂O into each sample, homogenization was carried out by using TissueLyser (Qiagen, Germany) for 90 s at 20 Hz. Subsequently, more 700 μl chloroform and 350 μl H₂O were added for further homogenization. The homogenate was centrifuged for 10 min at 10,621g at 4 °C. The supernatant was dried under gentle nitrogen stream. And then the dried residue was added with 600 μl D₂O containing 0.05% TSP, vortexed and centrifuged for 8 min at 20,817g. The supernatant was removed into a standard 5-mm NMR tube for analysis.

One-dimensional NMR (¹H-NMR) spectra were obtained on a 600-MHz Bruker Avance III HD spectrometer equipped with a 5-mm PABBO BB-probe. The spectra were acquired with presaturation of the water signal using the Bruker pulse program ‘noesygpprld’ with a 16-p.p.m., 9,590.793 Hz width and 4.27 s acquisition time. All spectra were recorded with 81,920 data points at 25 °C. The time domain data were Fourier-transformed, phase-corrected and baseline-corrected manually. The signal intensities were normalized against the intensity of the TSP signal at 0.00 p.p.m.

GC–MS analysis of lipid pool

HepG2 stable cells (5 × 10⁶) were cultured in DMEM medium (without glucose or glutamine, Sigma) by adding 10% dialysed FBS (BI), 25 mM label-free glucose, 4 mM label-free glutamine and 5 mM [U-¹³C₂]-acetate for 24 h under 1% O₂. Cells were rapidly washed twice with 37 °C PBS, then 1 ml PBS was added to the dish and quickly detached from the dish using a cell lifter. The liquid-containing cells were transferred into a 2-ml tube and centrifuged at 15,294g for 10 min at 4 °C. The supernatant was discarded and the residue was added to 400 μl of cold (80:20 methanol: water) extraction solution for quenching. The sample was vortexed and then stored at −20 °C for 1 h. The samples will be vortexed and centrifuged at 15,294g for 10 min at 4 °C. The supernatant will be moved to new tubes for dryness and used for further detection.

The sample preparation of the saponified fatty acids has been described previously⁷. Briefly, 1 ml MeOH/H₂O (1:1, v/v) solution with 0.1 M HCl and 1.6 p.p.m. internal standard (myristic acid-D27) in solution was added into cell debris at 0 °C, vortexed for 30 s, followed by addition of 0.5 ml chloroform, and vortexed for 1 min, then centrifuged at 12,000g for 5 min at 4 °C. The resulting chloroform layer was transferred to a new glass vial, and the extract was dried under N₂. The desiccation was reconstituted into 1 ml MeOH/H₂O (9:1, v/v) containing 0.3 M NaOH, and incubated at 80 °C for 1 h to saponify the fatty acids. Then 0.1 ml of formic acid was added for acidification, followed by extraction twice with 1 ml hexane. The hexane layer was transferred to an Eppendorf tube for dryness under N₂. The dried residue was added to 20 μl of pyridine and 80 μl of BSTFA (with 1% TMCS) and vortexed for 30 s, then derivatized at 70 °C for 1 h. The mixture was vortexed for 30 s and centrifuged at 12,000g for 5 min at room temperature before GC–MS analysis.

Analysis was performed using an Agilent 7890B gas chromatography system coupled to an Agilent 5977A mass spectrometric detector (MSD, Agilent Technologies). Derivatives were separated using an Agilent 122-5532 UI, DB-5MS capillary column (30 m × 250 μm × 0.25 μm). The injection volume was 1 μl in split mode in the ratio 5:1, and the solvent delay time was set to 6 min. The initial oven temperature was set at 100 °C for 1 min, then ramped to 200 °C at a rate of 20 °C min⁻¹, to 260 °C at a rate of 10 °C min⁻¹, then to 300 °C at a rate of 20 °C min⁻¹ and finally set at 300 °C for 5 min. Helium was used as a carrier gas at a constant flow rate of 1 ml min⁻¹ through the column. The temperatures of the front inlet, Aux-2 temperature and electron MS source were set at 280 °C, 250 °C and 230 °C, respectively. The electron energy was 70 eV, and the mass spectral data were collected in a full-scan mode (m/z 50–600). The mass isotopomer distribution of ¹³C-labelled palmitate and stearate was normalized to internal standard and cell number. Natural isotope abundance was corrected using IsoCor⁵⁸.

Cell survival assay

shACSS1/2 or shscr HepG2 stable cells were plated in 96-well plates at 1.5 × 10⁴ per well in 200 μl of medium treated with indicated conditions. The cell viability was measured after 48 h treatment by Cell Counting Kit-8 (Beyotime Biotechnology) according to the manufacturer’s instructions. CCK-8 solution was incubated with cells for 1 h at 37 °C. Absorbances at 450 nm were collected by ELx800 Universal Microplate Reader (BioTek).

Matrigel assay

Matrigel assay was performed as previously reported with minor modification⁵⁹. Briefly, shscr or shACSS1/2 HepG2 cells were plated in 96-well plates (at 5,000 per well) precoated with 100% Matrigel bed and cultured in DMEM supplemented with 10% FBS, and 2% Matrigel for 12 days. The fresh culture media was exchanged every 2 days. The spheroids were monitored by microscopy. Spheroid size was determined at the times indicated by automated imaging on an inverted microscope (Olympus Fluorescence Microscope IX18).

Hepatocarcinoma cancer samples

Hepatocarcinoma cancer samples were obtained from the 10th People’s Hospital, Shanghai (affiliated to Tongji University), with consents from the patients. The procedure related to human subjects was approved by the Ethics Committee of the Institutes of Biomedical Sciences (IBS), Fudan University. Direct immunoblotting was performed as mentioned above.

IHC staining

Human HCC sections were de-paraffinized in xylene and hydrated in graded ethanol, followed by deionized water. Endogenous peroxidase activity was inactivated with 3% hydrogen peroxidase in methanol for 30 min. Antigen retrieval was performed by boiling the sections in 0.01 M citrate buffer (pH 6.0) for 25 min in microwave oven. The sections were incubated with normal goat serum for 20 min to block non-specific staining and then incubated overnight at 4 °C with primary antibodies. After incubation for 45 min with a HRP-conjugated anti-rabbit secondary antibody, the sections were detected using diaminobenzidine (DAB) kit according to the manufacturer’s instructions, followed by counterstaining with hematoxylin. The primary antibodies were used as follows: anti-H3K9 acetyl (ab32129, Abcam, 1:150 dilution), anti-H3K27 acetyl (ab45173, Abcam, 1:80 dilution), anti-H3K56 acetyl (ab76307, Abcam, 1:60 dilution), anti-ACSS2 (ab133664, Abcam, 1:50 dilution), anti-ACSS1 antibody (HPA041014, Sigma, 1:50 dilution), and anti-FASN (C-20140, Santa Cruz, 1:40 dilution). The negative controls were performed by omitting the primary antibodies.

Statistical analysis

All data were statistically analysed with two-tailed unpaired Student’s t-test. The correlation analysis between FASN and ACSS1/2 mRNA levels in 190 human HCC samples from TCGA data base were carried out by using Pearson’s coefficient test and the correlation analysis between histone acetylation and FASN expression in human HCC samples by Spearman’s correlation test. The values of P<0.05 were considered as statistically significant.

Data availability

The analysis result in Supplementary Fig. 6a is based upon data generated by the TCGA Research Network: http://cancergenome.nih.gov/ Data supporting the findings of this study are available within the article and its Supplementary Information files, and from the corresponding author upon request.

Additional information

How to cite this article: Gao, X. et al. Acetate functions as an epigenetic metabolite to promote lipid synthesis under hypoxia. Nat. Commun. 7:11960 doi: 10.1038/ncomms11960 (2016).

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Acknowledgements

We thank the members of the Fudan Cancer Metabolism Laboratory for participation throughout this study. We also thank Biomedical Core Facility, Fudan University for technical support. This work was supported by MOST(2015CB910401 and 2013CB911201), NSFC (grant no. 81430057, 81225016, 31271454 and 21377106), Shanghai Key basic research program (12JC1401100), Scholar of ‘Dawn’ Program of Shanghai Education Commission, Shanghai Outstanding Academic Leader (grant no. 13XD1400600), Cheung Kong Scholar’s Program, and the Youth Science and Technology Leading Talent by MOST to Q.Y.L.

Author information

Xue Gao and Shu-Hai Lin: These authors contributed equally to this work.

Authors and Affiliations

and Department of Biochemistry and Molecular Biology, Key Laboratory of Metabolism and Molecular Medicine, Ministry of Education, School of Basic Medical Sciences, and Institutes of Biomedical Sciences, Fudan University, Shanghai, 200032, China
Xue Gao, Feng Ren, Jin-Tao Li, Jia-Jia Chen, Chuan-Bo Yao, Hong-Bin Yang, Guo-Quan Yan, Di Wang, Yi Wang, Ying-Ying Xu, Wenqiang Yu, Peng-Yuan Yang & Qun-Ying Lei
School of Life Science, Fudan University, Shanghai, 200433, China
Xue Gao, Jin-Tao Li & Di Wang
Department of Biochemistry and Molecular Cell Biology, Shanghai Key Laboratory for Tumor Microenvironment and Inflammation, Shanghai Jiao Tong University School of Medicine (SJTU-SM), Shanghai, 200025, China
Shu-Hai Lin
Department of Chemistry, State Key Laboratory of Environmental and Biological Analysis, Hong Kong Baptist University, Hong Kong, 999077, China
Shu-Hai Lin, Shu-Xia Jiang & Zongwei Cai
Department of Chemistry, Fudan University, Shanghai, 200032, China
Guo-Quan Yan
Department of Pathology, School of Basic Medical Sciences, Fudan University, Shanghai, 200032, China
Ying Liu
Department of Hematology and Medical Oncology, Winship Cancer Institute of Emory, Emory University School of Medicine, Atlanta, 30322, Georgia, USA
Jing Chen

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Xue Gao
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X.G. performed part of the experiments, analysed the data and co-wrote the manuscript. F.R., J.T.L. and C.B.Y. assisted in doing the experiments, data analysis and proofreading the manuscript. S.H.L., J.J.C., P.Y.Y. and S.X.J. contributed to GC–MS analysis of lipid pool and ¹H-NMR spectra of acetate. J.J.C., Y.W., G.Q.Y. and P.Y.Y. contributed to MS analysis. H.B.Y. contributed to analysis of human hepatocarcinoma cancer samples and proofreading the manuscript. D.W. and Y.Y.X. contributed to performing experiments and proofreading the manuscript. Y.L. contributed to analysis of IHC samples. J.C. and W.Q.Y. contributed to proofreading the manuscript. S.H.L., Z.W.C. and P.Y.Y. contributed to design MS experiment and data analysis. Q.Y.L. conceived the idea, designed and supervised the study, analysed the data and co-wrote the manuscript.

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Correspondence to Peng-Yuan Yang or Qun-Ying Lei.

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Supplementary Figures 1-7 and Supplementary Table 1 (PDF 7087 kb)

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Gao, X., Lin, SH., Ren, F. et al. Acetate functions as an epigenetic metabolite to promote lipid synthesis under hypoxia. Nat Commun 7, 11960 (2016). https://doi.org/10.1038/ncomms11960

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Received: 11 January 2016
Accepted: 17 May 2016
Published: 30 June 2016
DOI: https://doi.org/10.1038/ncomms11960

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