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Hypoxia causes triglyceride accumulation byHIF-1-mediated stimulation of lipin 1 expression

Ilias Mylonis1, Hiroshi Sembongi2, Christina Befani1, Panagiotis Liakos1, Symeon Siniossoglou2 andGeorge Simos1,3,*1Laboratory of Biochemistry, Medical School, University of Thessaly, BIOPOLIS, Larissa 41110, Greece2Cambridge Institute for Medical Research, University of Cambridge, Wellcome Trust/Medical Research Council Building, Hills Road, Cambridge,CB2 0XY, UK3Institute of Biomedical Research and Technology (BIOMED), 51 Papanastasiou Street, Larissa 41222, Greece

*Author for correspondence (simos@med.uth.gr)

Accepted 6 March 2012Journal of Cell Science 125, 3485–3493� 2012. Published by The Company of Biologists Ltddoi: 10.1242/jcs.106682

SummaryAdaptation to hypoxia involves hypoxia-inducible transcription factors (HIFs) and requires reprogramming of cellular metabolism that isessential during both physiological and pathological processes. In contrast to the established role of HIF-1 in glucose metabolism, the

involvement of HIFs and the molecular mechanisms concerning the effects of hypoxia on lipid metabolism are poorly characterized.Here, we report that exposure of human cells to hypoxia causes accumulation of triglycerides and lipid droplets. This is accompanied byinduction of lipin 1, a phosphatidate phosphatase isoform that catalyzes the penultimate step in triglyceride biosynthesis, whereas lipin 2remains unaffected. Hypoxic upregulation of lipin 1 expression involves predominantly HIF-1, which binds to a single distal hypoxia-

responsive element in the lipin 1 gene promoter and causes its activation under low oxygen conditions. Accumulation of hypoxictriglycerides or lipid droplets can be blocked by siRNA-mediated silencing of lipin 1 expression or kaempferol-mediated inhibition ofHIF-1. We conclude that direct control of lipin 1 transcription by HIF-1 is an important regulatory feature of lipid metabolism and its

adaptation to hypoxia.

Key words: Hypoxia, Lipid metabolism, HIF, Lipin, Phosphatidate phosphatase, PAP1

IntroductionExposure of human tissues or cells to reduced oxygen

concentration (hypoxia) is encountered both during physiological

and pathological processes (Semenza, 2011) and causes dramatic

changes in gene expression. Essential to this response are the

hypoxia-inducible transcription factors HIF-1 and HIF-2 (EPAS1),

recent major anti-cancer targets (Majmundar et al., 2010). Under

normal oxygen conditions the regulatory alpha subunit (HIFa) of

HIFs is continuously produced and destroyed, in a process

involving modification by prolyl hydroxylases (PHDs), VHL-

mediated polyubiquitination and subsequent proteasomal

degradation (Epstein et al., 2001; Ivan et al., 2001; Kallio et al.,

1999; Kamura et al., 2000; Maxwell et al., 1999; Turkish and

Sturley, 2007). When oxygen concentration is low, hydroxylation

is impaired and HIFa is stabilized, translocates to the nucleus,

dimerizes with ARNT (HIFb) to form HIF and binds to hypoxia-

responsive elements (HREs) in the promoters/enhancers of its

target genes.

Tissue-specific and differential control of HIFs also involves

several oxygen-independent molecular mechanisms. For HIF-1athese include regulation of its mRNA synthesis by NF-kB (Belaiba

et al., 2007; Rius et al., 2008) or the PKR–Stat3 axis (Papadakis

et al., 2010), control of its stability by phosphorylation (Flugel et al.,

2007) or protein–protein interactions (Liu et al., 2007), MAPK-

dependent regulation of its nuclear transport (Mylonis et al., 2008)

and CK1-mediated control of its heterodimerization (Kalousi et al.,

2010).

One of the major adaptive changes in response to hypoxia is

HIF-mediated reprogramming of cellular metabolism. HIF-1

activates genes responsible for switching from oxidative to

glycolytic metabolism such as the ones coding for glucose

transporters, glycolytic enzymes, lactate dehydrogenase and

pyruvate dehydrogenase kinase 1 (Majmundar et al., 2010).

Furthermore, hypoxia has been shown to stimulate lipid storage

and inhibit lipid catabolism in cultured cardiac myocytes and

macrophages (Bostrom et al., 2006; Huss et al., 2001), but the

involvement of HIFs in these processes was not investigated. More

recent studies using transgenic mice have suggested that artificial

or diet induced activation of liver HIF signalling can augment lipid

accumulation by modifying hepatocyte lipid metabolism (Kim

et al., 2006; Nath et al., 2011; Qu et al., 2011; Rankin et al., 2009).

Although these studies have implicated both HIF-1a and HIF-2ain liver fat accumulation and the subsequent maladaptive

pathologies, their particular roles have remained largely unclear

or controversial and no direct HIF targets have been so far

identified among the major lipidogenic enzymes.

A key role in lipid biosynthesis is played by lipins, a family of

proteins with Mg2+-dependent phosphatidate phosphatase (PAP1)

activity that catalyze the conversion of phosphatidic acid (PA) to

diacylglycerol (DAG) in the penultimate step of triglyceride (TG)

synthesis (Donkor et al., 2007; Han et al., 2006). TGs form the

core of lipid droplets (LD), the major neutral lipid stores in

adipocytes as well as other cell types. Mutations in the lipin 1

gene (Lpin1), which is expressed predominantly in adipose tissue

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and skeletal muscle, are the cause of lipodystrophy in the fattyliver dystrophic (fld) mouse and, conversely, overexpression of

lipin 1 in the adipose tissue of transgenic mice promotes obesity(Peterfy et al., 2001; Phan and Reue, 2005). Moreover, lipin 1 aswell as lipin 2, a liver-enriched isoform, interact with and/or

modulate the activity of several transcription factors, includingmembers of the peroxisome proliferator-activated receptor(PPAR) family and SREBP that control the expression of genesinvolved in fatty acid and lipid metabolism (Donkor et al., 2009;

Finck et al., 2006; Koh et al., 2008; Peterson et al., 2011). Apartof its apparently important role in adipogenesis, relatively little isknown about the function of lipin 1 in cells other than adipocytes.

Various stimuli such as glucocorticoids, insulin or fastingalter lipin 1 expression and/or activity, regulation of which isincompletely understood and involves transcriptional control,

post-translational modifications and compartmentalization (reviewedby Harris and Finck, 2011).

We now report that the lipin 1 gene is directly controlled by

HIF-1 and its up-regulation is linked to TG and LD accumulationin cells of non-adipocyte origin, suggesting that lipin 1 is animportant mediator of the metabolic adaptation to hypoxia at the

cellular level. Moreover, inhibition of HIF-1 by pharmacologicalor genetic means can suppress the hypoxic induction of lipin 1and TG or LD production.

ResultsHypoxia induces lipin 1-dependent triglycerideaccumulation

To investigate the effect of hypoxia on lipid biosynthesis

we analysed human hepatoblastoma (Huh7) and cervicaladenocarcinoma (HeLaM) cells exposed to normoxia orhypoxia (1% O2). As expected, hypoxia caused a robustexpression of nuclear HIF-1a that could be detected by

immunofluorescence microscopy (Fig. 1A,B, left panels).Furthermore, Nile Red staining revealed that in both cell typeshypoxia caused a significant and time-dependent increase in LD

accumulation, which was comparable to that caused by oleic acid(OA) treatment used as positive control under normoxia(Fig. 1A,B, middle panels). The induction of neutral lipid

accumulation by hypoxia was confirmed in Huh7 cells bydetermining the cellular TG content using a biochemical assay(Fig. 1C). Also in this case, TG accumulation under hypoxia wassimilar to the accumulation triggered by oleic acid.

Analysis of Huh7 cells by western blotting showed that inaddition to HIF-1a, the protein levels of lipin 1, which is considered

to be the minor liver isoform, also increased in response to hypoxia(Fig. 2A). On the contrary, lipin 2, normally the major hepaticisoform, remained unaffected indicating an isoform-specific effectof hypoxia on lipin expression. Similar results were also obtained

with HeLaM cells treated with hypoxia or dimethyloxalyl glycine(DMOG), a PHD inhibitor that stabilizes HIFa and activates theHIFs (Fig. 2B). Induction of lipin 1 by hypoxia was also confirmed

using immunofluorescence microscopy in both Huh7 (Fig. 2C) andHeLaM (Fig. 2D) cells. The overexpression of endogenous lipin 1under low oxygen conditions allowed its visualization, which was

previously not easily attainable in this type of cells. In Huh7 cells,hypoxically induced lipin 1 displayed a predominant cytoplasmic,both diffuse and reticular, localization. Nuclear staining was weak

but became more evident at later time points (Fig. 2C). Similarimages were obtained in HeLaM cells, in which, however,cytoplasmic spots and nuclei were stained more intensely at early

and late time points, respectively (Fig. 2D). Therefore, hypoxia

causes an increase in the cytoplasmic protein levels of lipin 1,

which, under prolonged treatment, also appears to migrate inside

the nucleus.

To test whether the effects of hypoxia on lipin 1 expression and

lipid droplet accumulation can also be reproduced in normal, non-

cancer cells we used primary, non-transformed human bronchial

smooth muscle (hBSM) cells. Analysis of these cells by western

blotting showed that both HIF-1a and lipin 1 increased in response

to hypoxia, especially after 48 hours of treatment, whereas lipin 2

expression was not detectable (Fig. 3A). Hypoxic induction was

confirmed using immunofluorescence microscopy (Fig. 3B), which

additionally showed that lipin 1 is predominantly cytoplasmic in

hBSM cells under normoxia but also translocates inside the nucleus

under hypoxia. Finally, Nile Red staining revealed that 48 hours of

hypoxic treatment causes significant LD accumulation also in hBSM

cells (Fig. 3C), suggesting that hypoxia affects lipin 1 expression

and neutral lipid content in both cancer and non-cancer cells.

The analysis was continued using Huh7 cells, which, as derived

by hepatocytes, are more appropriate for studying the relationship

between lipin 1 induction and lipid accumulation. First, to

investigate the increase in lipin 1 protein levels, we determined

Fig. 1. Hypoxia induces neutral lipid accumulation in Huh7 and HeLaM

cells. (A,B) Fluorescence microscopy analysis. Huh7 (A) or HeLaM (B) cells

incubated at 1% O2 for 0–48 hours or with 0.4 mM oleic acid (OA) for

24 hours were processed for immunodetection of HIF-1a, stained with Nile

Red to visualize lipid droplets and DAPI to show the nuclei and observed by

fluorescence microscopy. (C) Determination of triglyceride content in Huh7

cells incubated as in A. Results are mean 6 s.e.m. *P,0.05.

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its mRNA levels using qPCR. Lipin 1 mRNA significantly

increased in Huh7 cells subjected to hypoxia for 8 hours, whereas

lipin 2 mRNA was little affected (Fig. 4A). In the same

experiment, hypoxia caused robust mRNA expression of VEGF,

a well-known HIF target. Interestingly, lipin 1 mRNA levels

decreased after 24 hours of hypoxic treatment (although still

remaining higher than normoxia) suggesting a transient effect or

the operation of a negative feedback loop, possibly related to the

nuclear entry of lipin 1 that was also observed at later time points.

Since Lpin1 appears to be an early hypoxia-inducible gene, we

addressed its involvement in hypoxic TG accumulation by siRNA-

mediated silencing. Knocking down of lipin 1 was effective both

under normoxia and hypoxia as judged by western blotting

(Fig. 4B, upper panel). Depletion of lipin 1 under normoxia did not

significantly affect the TG content of the cells (Fig. 4B, lower

panel). In contrast, suppression of lipin 1 expression completely

blocked hypoxia-inducible TG accumulation. To a lesser extent,

lipin 1 knockdown also diminished TG accumulation triggered by

OA treatment, in agreement with a general role of lipin 1 in

response to lipidogenic stimuli in hepatocytes. Therefore, lipin 1 is

not only transcriptionally controlled by hypoxia but is also

required for the ensuing overproduction of triglycerides.

HIF-1 mediates hypoxic up-regulation of lipin 1 and TG

synthesis

We then investigated the role of HIFs in the hypoxic induction of

lipin 1 and TG synthesis. Overexpression of either HIF-1a or

HIF-2a in Huh7 cells increased the protein levels of lipin 1, with

HIF-1a being considerably more effective (Fig. 5A). In contrast,

neither HIF-1a nor HIF-2a overexpression affected lipin 2

expression. Quantification of the fluorescent signal of Huh7 cells

overexpressing HIFa and stained with Nile Red showed that

HIFa overexpression also causes LD accumulation (Fig. 5B). In

accordance to its effect on lipin 1 expression, HIF-1a exerted a

stronger effect on LD content than HIF-2a. Therefore, HIF-1 and,

to lesser extent, HIF-2 activation is sufficient to induce both lipin

1 and LD production even under normoxic conditions.

The involvement of HIFs in the hypoxic induction of lipin 1

was further tested by siRNA-mediated repression of HIF-1a or

HIF-2a in Huh7 cells treated with DMOG. Knockdown of HIF-

1a greatly reduced lipin 1 expression (Fig. 5C). HIF-2a silencing

also had a negative, but considerably weaker, effect on lipin 1.

Neither knockdowns affected lipin 2. These data imply that

hypoxic lipin 1 induction and subsequent TG and LD

accumulation is predominantly mediated by HIF-1, with HIF-2

playing a possible minor role. Knocking down HIF-1a using a

shRNA plasmid also largely eliminated the capacity of hypoxia

Fig. 2. Hypoxia induces lipin 1 expression in Huh7 and HeLaM cells.

(A) Western blotting analysis of Huh7 cells incubated at 1% O2 for 0–

48 hours. (B) Western blotting analysis of HeLaM cells incubated at 1% O2 or

with 1 mM DMOG for 0–24 hours. (C) Immunofluorescence microscopy

analysis. Huh7 cells were incubated at 1% O2 for 0–48 hours, analyzed by

immunofluorescence microscopy for HIF-1a and lipin 1 and stained with

DAPI to show the nuclei. (D) Immunofluorescence microscopy analysis.

HeLaM cells were incubated at 1% O2 for 0–24 hours and analyzed as in C. Fig. 3. Hypoxia induces lipin 1 expression and lipid droplet accumulation

in hBSM cells. (A) Western blotting analysis of hBSM cells incubated at 1%

O2 for 0–48 hours. (B) Immunofluorescence microscopy analysis. hBSM

cells were incubated at 1% O2 for 0–48 hours, analyzed by

immunofluorescence microscopy for HIF-1a and lipin 1 and stained with

DAPI to show the nuclei. (C) Fluorescence microscopy analysis. hBSMC

cells were incubated at 1% O2 for 0–48 hours, processed for immunodetection

of HIF-1a, stained with Nile Red to visualize lipid droplets and DAPI to show

the nuclei and observed by fluorescence microscopy.

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to induce lipin 1 (Fig. 5D) and, moreover, caused a modest but

significant reduction in hypoxic TG accumulation (Fig. 5E),

while it did not affect TG accumulation caused by OA treatment.

Oxygen tension and hydroxylation regulate HIF-1a protein

stability. However, the ability of HIF-1a to enter the nucleus and

activate transcription is additionally controlled by phosphorylation.

We have previously shown that CK1d targets HIF-1a and inhibits

its activity by impairing its binding to ARNT (Kalousi et al., 2010),

while phosphorylation by p44/42 MAPK stimulates HIF-1aactivity by blocking its nuclear export (Mylonis et al., 2008). To

correlate lipin 1 expression to HIF-1a activity rather than just its

protein levels, we used kinase inhibitors. Treatment of Huh7 cells

with the CK1d inhibitor IC261, which stimulates HIF-1a activity

(Kalousi et al., 2010), potentiated hypoxic induction of lipin 1

without significantly affecting lipin 2 or HIF-1a protein levels

(Fig. 5F). Inversely, treatment of Huh7 cells with the flavonoid

kaempferol, which suppresses HIF-1a activity by impairing its

nuclear accumulation through MAPK inhibition (Mylonis et al.,

2010), diminished lipin 1 expression under hypoxia (Fig. 6A,B).

The same treatment also significantly reduced the amount of LDs in

Huh7 cells grown under hypoxia (Fig. 6B,C). Therefore, both gain-

of-function and loss-of-function experiments strongly suggest that

hypoxic induction of lipin 1 and neutral lipid accumulation involves

predominantly HIF-1.

The human Lpin1 promoter contains a conserved

functional HRE

A direct role of HIF-1 in Lpin1 regulation requires the

contribution of an HRE. Examination of the 3.5 kb promoter

region of Lpin1 revealed the presence of 8 potential HRE sites

conforming to the consensus XCGTG (X5A or G or C). To test

Fig. 4. Hypoxic triglyceride accumulation is lipin 1 dependent. (A) Lipin

1 is transcriptionally upregulated by hypoxia. Determination of lipin 1, lipin 2

and VEGF mRNA levels in Huh7 cells incubated at 1% O2 for 0–24 hours.

(B) Lipin 1 is required for TG accumulation under hypoxia. Top panel: Huh7

cells transfected with lipin 1 or control siRNAs and incubated at 20% or 1%

O2 for 48 hours were analyzed by western blotting. Bottom panel:

Determination of triglyceride content in cells treated as above or with 0.4 mM

oleic acid (OA) for the same period. Values in charts represent the mean 6

s.e.m. of two independent experiments performed in triplicate (*P,0.05;

***P,0.001).

Fig. 5. HIF-1 mediates hypoxic upregulation of lipin 1 and TG synthesis.

(A) HIF overexpression induces lipin 1. Western blot analysis of Huh7 cells

overexpressing GFP, GFP-HIF-1a or GFP-HIF-2a. (B) HIF overexpression

induces LD accumulation. Huh7 cells overexpressing GFP, GFP-HIF-1a or

GFP-HIF-2a were stained with Nile Red and their fluorescent signal was

monitored by microscopy and quantified using ImageJ software. Values

represent the mean area of Nile Red staining of 50 GFP-negative (2) or GFP-

positive (+) cells (6 s.e.m.) from two independent experiments (*P,0.05).

(C) HIFs are required for induction of lipin 1 by DMOG. Western blot

analysis of Huh7 cells transfected with siRNA against HIF-1a or HIF-2a and

treated with 1 mM DMOG for 8 hours. (D) HIF-1 is required for hypoxic

induction of lipin 1. Western blot analysis of Huh7 cells transfected with

pSUPERshHIF1 or a control plasmid and incubated under 1% or 20% O2 for 8

hours. (E) HIF-1 is involved in the hypoxic induction of TG accumulation.

Determination of triglyceride content in Huh7 cells transfected with

pSUPERshHIF1 or a control plasmid and incubated for 48 hours under 20%

or 1% O2 or treated with 0.4 mM oleic acid (OA) for 48 hours. Data represent

the mean (6 s.e.m.) of two independent experiments performed in

quadruplicate (**P,0.01). (F) Activation of HIF-1 by CK-1d inhibition

enhances hypoxic induction of lipin 1. Western blot analysis of Huh7 cells

incubated for 8 hours under 20% or 1% O2 with or without 2 mM IC261.

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the functionality of these potential HREs, we constructed

luciferase reporter plasmids containing the whole region

(3298 bp; HRE1–8) or 59 truncated forms containing 7

(HRE2–8), 4 (HRE5–8), 2 (HRE7–8) or 1 (HRE8) proximal

potential HREs (Fig. 7A, upper panel) and introduced them in

Huh7 cells. The full-length region (HRE1–8) exhibited the highest

promoter activity under normoxia and, more importantly, it was

the only one responding to and activated by hypoxia (Fig. 7A,

upper panel), suggesting that only the most distal HRE (HRE1) is

functional. This was also positively shown by using two 39-

truncated forms of the Lpin1 promoter region containing only

HRE1 or four distal HRE sites (HRE1–4): both of these promoter

fragments could also be activated by hypoxia (Fig. 7A, lower

panel). HRE1 is conserved (Fig. 7B) as it is perfectly matched by

a potential distal HRE in the mouse Lpin1 promoter region and is

found close to the only other known distal regulatory elements,

the SRE and NF-Y sites (Ishimoto et al., 2009). Finally, mutation

of HRE1 (Fig. 7B), in the context of both full-length and

truncated Lpin1 promoter regions (producing mHRE1–8, mHRE1

and mHRE1–4), led to complete loss of responsiveness to hypoxia

(Fig. 7C). Therefore, transcriptional activation of Lpin1 under

hypoxia is mediated by a single distal HRE site in its promoter.

HIF-1 directly interacts with the Lpin1 promoter

Interaction of HIFs with HRE1 in the Lpin1 promoter was tested by

introducing wild-type or mutant reporter constructs HRE1–8, HRE1

and HRE2–8 in Huh7 cells overexpressing HIF-1a or HIF-2a. HIF-1

Fig. 6. Inhibition of HIF-1 by kaempferol impairs hypoxic induction of

lipin 1 expression and LD accumulation. (A) Treatment with kaempferol

inhibits hypoxic induction of lipin 1. Western blot analysis of Huh7 cells

incubated for 8 hours under 20% or 1% O2 with or without or 10 mM

kaempferol (Kae). (B,C) Treatment with kaempferol inhibits hypoxic LD

accumulation. Huh7 cells incubated for 24 hours under 1% O2 with or without

50 mM kaempferol (Kae) were processed for immunodetection of HIF-1a,

stained with Nile Red and monitored with fluorescence microscopy (B). Nile

Red staining was quantified using ImageJ software and the results represent

the mean area of Nile Red staining of 50 cells grown in the absence (2) or

presence (+) of kaempferol (6 s.e.m.) (*P,0.05).Fig. 7. The human Lpin1 promoter contains a conserved functional distal

HRE. (A) A single distal HRE is both necessary and sufficient for the

hypoxic induction of the Lpin1 promoter. Luciferase activity in Huh7 cells

transfected with pGL3 constructs of the human Lpin1 promoter or its

fragments and incubated under 20% or 1% O2 for 16 hours. Values are

expressed in relation to the values obtained for the empty pGL3 vector and

represent the mean (6 s.e.m.) of three independent experiments performed in

triplicate. The potential HREs in the promoter region are schematically

depicted as inverted triangles. (B) The distal HRE in Lpin1 promoter is

conserved. Top panel: Schematic representation of the human and mouse

Lpin1 promoter regions (23300 to 21) depicting potential HRE sites

(inverted triangles), conserved SRE and NF-Y elements (grey polygon and

circle, respectively) and the distal functional HRE (boxed). Bottom panel:

Sequence alignment of the region of the human and mouse Lpin1 promoters

surrounding HRE1. Asterisks indicate conserved nucleotides. The sequence of

the HRE1 and its mutated form are depicted in boxes. (C) Mutation of the

distal HRE inhibits hypoxic induction of the Lpin1 promoter. Luciferase

activity in Huh7 cells transfected with pGL3 constructs containing wild-type

or mutant fragments of the human Lpin1 promoter and incubated for 16 hours

under normoxia or hypoxia. Values are expressed as percentage activity in

relation to normoxia and represent the mean (6 s.e.m.) of three independent

experiments performed in triplicate. The potential HREs are schematically

depicted as inverted triangles and a grey X indicates the mutation.

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could activate transcription from wild-type full-length or single

HRE1 promoter constructs but not from constructs lacking HRE1

(HRE2–8) or containing its mutant form (mHRE1–8 and mHRE1;

Fig. 8A). HIF-2 also activated the wild-type full-length promoter,

albeit weaker than HIF-1, but failed to activate transcription of the

single distal HRE construct (HRE1) suggesting that its role in Lpin1

activation is minor and involves additional regulatory elements

aside of HRE1.

Involvement of endogenous HIF-1 in Lpin1 regulation was

tested in cells transformed with the full-length Lpin1 promoter

construct and overexpressing the wild-type or SA mutant form of

the HIF-1a MTD under hypoxia. Overexpression of wild-type

MTD (MAPK-target domain), a 42 amino acid long fragment of

HIF-1a containing its MAPK modification sites, specifically

inhibits HIF-1a by blocking its phosphorylation and triggering its

nuclear export, while the mutant SA form of MTD lacks the

MAPK sites and is inactive (Mylonis et al., 2008). In cells

expressing wild-type MTD the hypoxic activation of Lpin1

promoter was virtually abolished but persisted in those cells

expressing the mutant SA MTD form (Fig. 8B), providing further

evidence that HIF-1 is the major transcriptional activator of

Lpin1 gene under hypoxia.

Direct binding of HIF-1a to HRE1 was tested by chromatin

immunoprecipitation using Huh7 cells cultured under hypoxia or

normoxia. The Lpin1 promoter region containing HRE1 was

greatly enriched in anti-HIF-1a immunoprecipitates of DNA-

protein complexes isolated from hypoxically treated cells in

comparison to rabbit IgG immunoprecipitates or anti-HIF-1aimmunoprecipitates from normoxic cells (Fig. 8C,D). Therefore,

HIF-1a not only activates but also physically associates with the

Lpin1 promoter.

DiscussionWe have revealed a direct link between HIF-1a and lipin 1 or the

cellular response to hypoxia and the metabolic pathway that leads

to synthesis of triglycerides and formation of lipid droplets

(Fig. 8E). Although this connection was identified in cultured

human cells, it can mechanistically explain previous observations

made with whole transgenic animals. Over-expression of a

constitutively active form of HIF-1a in mouse liver led to hepatic

fat accumulation and liver steatosis (Kim et al., 2006). In

contrast, similar over-expression of HIF-2a did not result in liver

steatosis but rather caused vascular lesions. Furthermore, lipid

accumulation in mouse hepatocytes caused by alcohol feeding

Fig. 8. HIF-1 directly interacts with and activates the Lpin1

promoter. (A) HIF-1 activates the Lpin1 promoter via the distal

HRE. Luciferase activity in Huh7 cells overexpressing GFP, GFP-

HIF-1a or GFP-HIF-2a and transfected with pGL3 constructs

containing wild-type or mutant fragments of the human Lpin1

promoter as indicated. Values are expressed as a percentage of

activity in relation to that of the empty GFP vector and represent

the mean (6 s.e.m.) of three independent experiments performed

in triplicate. (B) Inhibition of HIF-1 by MTD overexpression

inhibits hypoxic induction of the Lpin1 promoter. Luciferase

activity in Huh7 cells co-transfected with pGL3 constructs of the

human Lpin1 promoter and plasmids expressing wild-type or

mutant (SA) MTD and incubated for 16 hours under 20% or 1%

O2. Values are expressed in relation to the values obtained for the

empty pGL3 vector and represent the mean (6 s.e.m.) of three

independent experiments performed in triplicate. (C) HIF-1a

associates with the distal HRE in the Lpin1 promoter. Gel

electrophoresis of PCR products amplified from anti-HIF-1a or

rabbit IgG chromatin immunoprecipitates of Huh7 cells, incubated

for 8 hours under 20% or 1% O2, using primers specific for the

distal HRE of the Lpin1 promoter (schematically indicated).

(D) Quantification of results in C by real-time PCR. The data

represent the mean (6 s.e.m.) of two independent experiments

performed in triplicate (***P,0.001). (E) Schematic model

describing a HIF-1- and lipin-1-dependent mechanism of hypoxic

induction of TG accumulation.

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was also shown to involve HIF-1a (Nath et al., 2011). On theother hand, in double knockout studies, hepatic steatosis caused

by liver-specific deficiency of pVHL could be avoided bysimultaneous inactivation of HIF-2a (Qu et al., 2011; Rankinet al., 2009). However, in these animal models HIF-2 was mainlyassociated with suppression of fatty acid oxidation, increased

lipid storage capacity, serum TG levels and inflammation. In ourstudy with human cells, HIF-2 also seems to be involved in lipin1 regulation and LD accumulation but its role is minor compared

to HIF-1 and probably indirect since it requires more complexregulatory elements. It, therefore, appears that HIF-1a and HIF-2a may have overlapping or complementary but distinct roles in

the regulation of hepatic lipid metabolism and the developmentof related pathologies such as alcoholic and non-alcoholic fatty-liver disease (AFLD and NAFLD, respectively), with HIF-1predominantly stimulating lipin 1 and triglyceride synthesis

under hypoxic or other stress conditions. HIF-1 has also beenimplicated in cardiac steatosis by controlling expression ofPPARc, a principal transcriptional activator of lipid anabolism

(Krishnan et al., 2009). Lipin 1 also co-activates PPARc (Kohet al., 2008), so the HIF-1–lipin-1 axis may serve to amplify notonly PAP1 activity but also the non-catalytic transcriptional lipin

1 functions.

In order for lipin 1 to exert its transcriptional role, it has totranslocate inside the nucleus. Although lipin 1 is predominantlycytoplasmic in our tested cell lines under normoxia, hypoxic

treatment increases its nuclear pool especially after long-terminduction. A recent report has shown that nuclear localization oflipin 1 is regulated by nutrient- and growth factor-responsive

kinase mTOR complex 1 (mTORC1)-mediated phosphorylation(Peterson et al., 2011). Inhibition of mTORC1 causes normoxicredistribution of lipin 1 from the cytoplasm to the nucleus, which

in turn reduces the nuclear protein levels of SREBP-1 (Petersonet al., 2011), a transcription factor regulating many lipogenicgenes, including lipin 1 (Ishimoto et al., 2009). Therefore, our

data showing nuclear accumulation of lipin 1 and a decrease oflipin 1 mRNA expression at later points of hypoxic treatmentcould be due to inhibition of mTORC1 that is known to occurafter prolonged exposure to hypoxia (Wouters and Koritzinsky,

2008). The physiological relevance of lipin 1 hypoxic nuclearmigration could thus be justified by the operation of thefollowing negative feedback mechanism. After the initial HIF-

1-mediated induction of lipin 1 and the subsequent TGaccumulation, inhibition of mTOR by prolonged hypoxiacauses translocation of lipin 1 inside the nucleus, where, by

suppressing SREBP-1 function, it reduces expression of its ownand other lipogenic genes and blocks further TG de novosynthesis.

The idea that cells in a stressful situation, such as oxygen

deprivation, respond by stimulating, even transiently, an anabolicprocess, e.g. lipid biosynthesis, is counterintuitive but not withoutprecedent. Hypoxia also stimulates glycogen synthesis in a HIF-

1-dependent way (Brahimi-Horn et al., 2011; Fakas et al., 2011;Pescador et al., 2010). Apparently, exposure of cells to mildhypoxia may serve as a way of sensing ensuing ischemia/nutrient

limitation and, while it is still available, glucose is stored for lateruse as anaerobic source of energy. However, this may not be thecase for lipids since they can only produce energy through

oxidative phosphorylation, a process drastically inhibited bysevere hypoxia. A more likely explanation for storingtriglycerides in the form of lipid droplets under hypoxia could

be that, in this way, cells buffer the toxicity caused by free fatty

acids, which build up because of suppressed respiratory chain

activity. Increased PAP1 activity and formation of lipid droplets

can, thus, be a protective response that prevents intracellular

lipotoxicity (Fakas et al., 2011; Turkish and Sturley, 2007)

although stored triglycerides could still serve as a source of

lipotoxic intermediates if the cells remain for long unable to

handle them metabolically (Neuschwander-Tetri, 2010).

Irrespective of its cellular role, the direct involvement of HIF-1

in triglyceride formation may open new avenues for therapeutic

interventions in lipid disorders leading to NAFLD and

steatohepatitis. Furthermore, LD accumulation in non-adipose

tissue (ectopic fat) is a hallmark of the metabolic syndrome and is

linked to the pathogenesis of insulin resistance and diabetes

(Unger et al., 2010). The extensive recent pharmacological

targeting of HIF-1 as means of anti-cancer or anti-ischemic

treatment, has led to the identification and availability of many

agents that inhibit or activate HIF-1, respectively (Semenza,

2011). Our results with HIF-1 inhibitors, such as kaempferol and

MTD, provide proof-of-principle that such agents may also be

useful as effective modulators of intracellular lipid metabolism

and can form the basis of novel molecular therapies targeting

lipin 1.

Materials and MethodsPlasmids

The human Lpin1 promoter region (nucleotides 23298 to 21; Acc. No.:NT_005334.16) was amplified by PCR using HeLaM total DNA as template,primers hL1prmF1 and hL1prmR1 and KOD Hot start DNA polymerase(TOYOBO Life Science, Tokyo, Japan). The PCR fragment was then introducedinto pUC19 vector, sequenced and subcloned into pGL3-basic vector (Promega,Madison, WI, USA) to construct pGL3-HRE1–8. Truncated promoter forms, pGL3-HRE2–8 (22198), pGL3-HRE5–8 (21297), pGL3-HRE7–8 (2602) and pGL3-HRE8 (2445) were constructed by using as template the pUC19-HRE1–8 plasmid,and an appropriate set of primers listed in supplementary material Table S1. Theoccurring PCR fragments were then subcloned into the pGL3-basic vector andsequenced. To construct HRE mutant plasmid pGL3-mHRE1–8, three nucleotidesubstitutions (CCGTG to CAACG) were introduced by PCR using hL1pHRE1mF1and hL1pHRE1mR1 primers listed in supplementary material Table S1. PlasmidspGL3-HRE1 (23298 to 22588), pGL3-HRE1–4 (23298 to 21585) and theirrespective mutant forms, pGL3-mHRE1 and pGL3-mHRE1–4, were constructed bydigesting pGL3-HRE1–8 with BglII, to yield fragment HRE1, or HindIII, to yieldHRE1–4, and subcloning to the empty pGL3-vector. pEGFP-HIF-1a, pCMV-FLAG-MTD and pCMV-FLAG-MTD-SA plasmids were previously described(Mylonis et al., 2008). To construct pEGFP-HIF-2a, full-length HIF-2a cDNA wasobtained from pcDNA-HIF-2a, kindly provided by S. L. McKnight [University ofTexas Southwestern Medical Center (Tian et al., 1997)].

Cell culture, transfection and reporter gene assays

Human HeLaM and Huh7 cells were cultured in DMEM (Biosera) containing10% FCS and 100 U/ml penicillin-streptomycin (Gibco BRL). Human bronchialsmooth muscle cells (Lonza, Switzerland) were cultured in DMEM F-12containing 10% FCS and 100 U/ml penicillin-streptomycin. When required,cells were treated for 4–24 hours with 1 mM dimethyloxalyl glycine (DMOG;Alexis Biochemicals), with CK1 inhibitor IC261 (2 mM; Sigma, St Louis, MO) orwith kaempferol (10–50 mM; Sigma). For oleic acid treatment, the medium wassupplemented with 0.4 mM oleic acid (Sigma) bound to 1.5% bovine serumalbumin. For hypoxic treatment, cells were exposed for 4–48 hours to 1% O2, 95%N2 and 5% CO2 in an IN VIVO2 200 hypoxia workstation (RUSKINN LifeSciences, Pencoed, UK). Transient transfections and reporter gene assays wereperformed as described previously (Mylonis et al., 2008).

Western blot and immunofluorescence

Antibodies used for western blotting, immunofluorescence and chromatinimmunoprecipitation were: affinity purified rabbit polyclonal antibodies againstHIF-1a (Lyberopoulou et al., 2007), lipin 1 and lipin 2 (Grimsey et al., 2008),rabbit polyclonal antibodies against GFP (Mylonis et al., 2008) or anti-HIF-2a(Abcam, Cambridge, UK) and mouse monoclonal antibodies against actin ortubulin (Millipore, Billerica, MA, USA). Analysis of total cellular proteins byimmunoblotting and immunofluorescence microscopy were carried out as

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previously described (Grimsey et al., 2008; Lyberopoulou et al., 2007; Mylonis

et al., 2008; Mylonis et al., 2006). To visualize lipid droplets after usualimmunofluorescence microscopy procedure cells were stained with Nile Red

(Sigma; 0.1 mg in PBS) for 15 minutes, washed with PBS and mounted on slides(Greenspan et al., 1985).

Triglyceride determination

Huh7 cells were lysed in 150 ml PBS with 0.1% Triton X-100 (Sigma). The

mixture was then sonicated (five 10 sec pulses at maximum intensity), boiled for5 minutes to facilitate the formation of an even suspension (Al-Anzi and

Zinn, 2010) and 20 ml aliquots were tested using a biochemical triglyceridedetermination kit (Biosys, Athens, Greece). Triglyceride values were normalized

to protein measured with Bio-Rad Protein Assay (Bio-Rad, Hercules, CA).

siRNA- and shRNA-mediated silencing

Huh7 cells were incubated in serum-free DMEM for 4 hours with siRNA (10 nM)

against HIF-1a or HIF-2a (Qiagen, Hilden, Germany) or Lpin1 (target sequencesshown in supplementary material Table S1) in the presence of Lipofectamine

RNAiMAX (Invitrogen, Carlsbad, CA). AllStars siRNA (Qiagen, Hilden,Germany) was used as negative control. shRNA-mediated silencing of HIF-1awas done using pSuper-shHIF-1a, kindly provided by I. Papandreou and N. C.Denko [Stanford University School of Medicine (Papandreou et al., 2006)] and

Lipofectamine 2000 (Invitrogen).

RNA extraction and Real-Time PCR

Total RNA from Huh7 cells was isolated using the Trizol reagent (Invitrogen) andcDNA was synthesized with the High Capacity Reverse Transcription kit (Applied

Biosystems, Carlsbad, CA). Real-time PCR was performed with SYBR GreenERqPCR SuperMix Universal (Invitrogen) in a MiniOpticon instrument (Bio-Rad).

The mRNAs encoding lipin 1, lipin 2 and VEGF were amplified using primerslisted in supplementary material Table S1. Each sample was assayed in triplicate

for both target and internal control. Relative quantitative gene expression wascalculated using the DDCt method and presented as relative units.

Chromatin immunoprecipitation

Huh7 cells were treated with 1.1% formaldehyde for 30 minutes, quenched with125 mM glycine for 5 min and collected by centrifugation. Following steps were

performed as previously described (Braliou et al., 2008) using a polyclonal anti-HIF-1a antibody (Lyberopoulou et al., 2007) or rabbit IgG. The region 22916 to

22686 of the hLPIN1 promoter was amplified from immunoprecipitatedchromatin or input samples using primers listed in supplementary material Table

S1. The 230 bp product was analyzed by agarose gel electrophoresis or quantifiedby real-time PCR. The Ct (threshold cycle) of immunoprecipitated samples (with

specific anti-HIF-1a antibody or with non-specific IgG) was normalized to the Ctof total input DNA (DCt). Results obtained with specific anti-HIF-1a antibody was

then normalized to those obtained with non-specific IgG antibodies (DDCt).

Densitometric and statistical analysis

Quantification of the surface covered by Nile Red fluorescence was performed

with ImageJ public domain software for image analysis and expressed as pixels/cell (Abramoff et al., 2004). Statistical differences between two groups of data

were assessed using the unpaired t-test in the SigmaPlot v. 9.0 software (Systat);P,0.05 was considered to be significant.

AcknowledgementsWe are grateful to Drs I. Papandreou, N. C. Denko and S. L.McKnight for generously providing plasmids and to Dr E. Paraskeva,Laboratory of Physiology, University of Thessaly, for providinghBMS cells and instructions for their culture.

FundingThis study was supported by the National Strategic ReferenceFramework/National Action ‘‘Cooperation’’ [grant number 09SYN-12-682] co-funded by the Greek Government and the EuropeanRegional Development Fund (G.S.); A Medical Research CouncilSenior Fellowship [grant number G0701446 to S.S.]; and a EuropeanMolecular Biology Organization short-term fellowship (I.M.).Deposited in PMC for release after 6 months.

Supplementary material available online at

http://jcs.biologists.org/lookup/suppl/doi:10.1242/jcs.106682/-/DC1

ReferencesAbramoff, M. D., Magelhaes, P. J. and Ram, S. J. (2004). Image Processing with

ImageJ. Biophot. Intern. 11, 36-42.

Al-Anzi, B. and Zinn, K. (2010). Colorimetric measurement of triglycerides cannotprovide an accurate measure of stored fat content in Drosophila. PLoS ONE 5,e12353.

Belaiba, R. S., Bonello, S., Zahringer, C., Schmidt, S., Hess, J., Kietzmann, T. andGorlach, A. (2007). Hypoxia up-regulates hypoxia-inducible factor-1alpha transcriptionby involving phosphatidylinositol 3-kinase and nuclear factor kappaB in pulmonaryartery smooth muscle cells. Mol. Biol. Cell 18, 4691-4697.

Bostrom, P., Magnusson, B., Svensson, P. A., Wiklund, O., Boren, J., Carlsson,

L. M., Stahlman, M., Olofsson, S. O. and Hulten, L. M. (2006). Hypoxia convertshuman macrophages into triglyceride-loaded foam cells. Arterioscler. Thromb. Vasc.

Biol. 26, 1871-1876.

Brahimi-Horn, M. C., Bellot, G. and Pouyssegur, J. (2011). Hypoxia and energetictumour metabolism. Curr. Opin. Genet. Dev. 21, 67-72.

Braliou, G. G., Verga Falzacappa, M. V., Chachami, G., Casanovas, G.,

Muckenthaler, M. U. and Simos, G. (2008). 2-Oxoglutarate-dependent oxygenasescontrol hepcidin gene expression. J. Hepatol. 48, 801-810.

Donkor, J., Sariahmetoglu, M., Dewald, J., Brindley, D. N. and Reue, K. (2007).Three mammalian lipins act as phosphatidate phosphatases with distinct tissueexpression patterns. J. Biol. Chem. 282, 3450-3457.

Donkor, J., Zhang, P., Wong, S., O’Loughlin, L., Dewald, J., Kok, B. P., Brindley,D. N. and Reue, K. (2009). A conserved serine residue is required for thephosphatidate phosphatase activity but not the transcriptional coactivator functions oflipin-1 and lipin-2. J. Biol. Chem. 284, 29968-29978.

Epstein, A. C., Gleadle, J. M., McNeill, L. A., Hewitson, K. S., O’Rourke, J., Mole,

D. R., Mukherji, M., Metzen, E., Wilson, M. I., Dhanda, A. et al. (2001). C.elegans EGL-9 and mammalian homologs define a family of dioxygenases thatregulate HIF by prolyl hydroxylation. Cell 107, 43-54.

Fakas, S., Qiu, Y., Dixon, J. L., Han, G. S., Ruggles, K. V., Garbarino, J., Sturley,S. L. and Carman, G. M. (2011). Phosphatidate phosphatase activity plays key rolein protection against fatty acid-induced toxicity in yeast. J. Biol. Chem. 286, 29074-29085.

Finck, B. N., Gropler, M. C., Chen, Z., Leone, T. C., Croce, M. A., Harris, T. E.,

Lawrence, J. C., Jr and Kelly, D. P. (2006). Lipin 1 is an inducible amplifier of thehepatic PGC-1alpha/PPARalpha regulatory pathway. Cell Metab. 4, 199-210.

Flugel, D., Gorlach, A., Michiels, C. and Kietzmann, T. (2007). Glycogen synthasekinase 3 phosphorylates hypoxia-inducible factor 1alpha and mediates its destabilizationin a VHL-independent manner. Mol. Cell. Biol. 27, 3253-3265.

Greenspan, P., Mayer, E. P. and Fowler, S. D. (1985). Nile red: a selective fluorescentstain for intracellular lipid droplets. J. Cell Biol. 100, 965-973.

Grimsey, N., Han, G. S., O’Hara, L., Rochford, J. J., Carman, G. M. and

Siniossoglou, S. (2008). Temporal and spatial regulation of the phosphatidatephosphatases lipin 1 and 2. J. Biol. Chem. 283, 29166-29174.

Han, G. S., Wu, W. I. and Carman, G. M. (2006). The Saccharomyces cerevisiaeLipin homolog is a Mg2+-dependent phosphatidate phosphatase enzyme. J. Biol.

Chem. 281, 9210-9218.

Harris, T. E. and Finck, B. N. (2011). Dual function lipin proteins and glycerolipidmetabolism. Trends Endocrinol. Metab. 22, 226-233.

Huss, J. M., Levy, F. H. and Kelly, D. P. (2001). Hypoxia inhibits the peroxisomeproliferator-activated receptor alpha/retinoid X receptor gene regulatory pathway incardiac myocytes: a mechanism for O2-dependent modulation of mitochondrial fattyacid oxidation. J. Biol. Chem. 276, 27605-27612.

Ishimoto, K., Nakamura, H., Tachibana, K., Yamasaki, D., Ota, A., Hirano, K.,Tanaka, T., Hamakubo, T., Sakai, J., Kodama, T. et al. (2009). Sterol-mediatedregulation of human lipin 1 gene expression in hepatoblastoma cells. J. Biol. Chem.

284, 22195-22205.

Ivan, M., Kondo, K., Yang, H., Kim, W., Valiando, J., Ohh, M., Salic, A., Asara, J. M.,

Lane, W. S. and Kaelin, W. G., Jr (2001). HIFalpha targeted for VHL-mediateddestruction by proline hydroxylation: implications for O2 sensing. Science 292, 464-468.

Kallio, P. J., Wilson, W. J., O’Brien, S., Makino, Y. and Poellinger, L. (1999).Regulation of the hypoxia-inducible transcription factor 1alpha by the ubiquitin-proteasome pathway. J. Biol. Chem. 274, 6519-6525.

Kalousi, A., Mylonis, I., Politou, A. S., Chachami, G., Paraskeva, E. and Simos, G.

(2010). Casein kinase 1 regulates human hypoxia-inducible factor HIF-1. J. Cell Sci.

123, 2976-2986.

Kamura, T., Sato, S., Iwai, K., Czyzyk-Krzeska, M., Conaway, R. C. and Conaway,

J. W. (2000). Activation of HIF1alpha ubiquitination by a reconstituted von Hippel-Lindau (VHL) tumor suppressor complex. Proc. Natl. Acad. Sci. USA 97, 10430-10435.

Kim, W. Y., Safran, M., Buckley, M. R., Ebert, B. L., Glickman, J., Bosenberg, M.,

Regan, M. and Kaelin, W. G., Jr (2006). Failure to prolyl hydroxylate hypoxia-inducible factor alpha phenocopies VHL inactivation in vivo. EMBO J. 25, 4650-4662.

Koh, Y. K., Lee, M. Y., Kim, J. W., Kim, M., Moon, J. S., Lee, Y. J., Ahn, Y. H. and

Kim, K. S. (2008). Lipin1 is a key factor for the maturation and maintenance ofadipocytes in the regulatory network with CCAAT/enhancer-binding protein alpha andperoxisome proliferator-activated receptor gamma 2. J. Biol. Chem. 283, 34896-34906.

Krishnan, J., Suter, M., Windak, R., Krebs, T., Felley, A., Montessuit, C.,Tokarska-Schlattner, M., Aasum, E., Bogdanova, A., Perriard, E. et al. (2009).Activation of a HIF1alpha-PPARgamma axis underlies the integration of glycolyticand lipid anabolic pathways in pathologic cardiac hypertrophy. Cell Metab. 9, 512-524.

Journal of Cell Science 125 (14)3492

Journ

alof

Cell

Scie

nce

Liu, Y. V., Baek, J. H., Zhang, H., Diez, R., Cole, R. N. and Semenza, G. L. (2007).RACK1 competes with HSP90 for binding to HIF-1alpha and is required for O(2)-independent and HSP90 inhibitor-induced degradation of HIF-1alpha. Mol. Cell 25,207-217.

Lyberopoulou, A., Venieris, E., Mylonis, I., Chachami, G., Pappas, I., Simos, G.,

Bonanou, S. and Georgatsou, E. (2007). MgcRacGAP interacts with HIF-1alpha andregulates its transcriptional activity. Cell. Physiol. Biochem. 20, 995-1006.

Majmundar, A. J., Wong, W. J. and Simon, M. C. (2010). Hypoxia-inducible factorsand the response to hypoxic stress. Mol. Cell 40, 294-309.

Maxwell, P. H., Wiesener, M. S., Chang, G. W., Clifford, S. C., Vaux, E. C.,

Cockman, M. E., Wykoff, C. C., Pugh, C. W., Maher, E. R. and Ratcliffe, P. J.(1999). The tumour suppressor protein VHL targets hypoxia-inducible factors foroxygen-dependent proteolysis. Nature 399, 271-275.

Mylonis, I., Chachami, G., Samiotaki, M., Panayotou, G., Paraskeva, E., Kalousi,

A., Georgatsou, E., Bonanou, S. and Simos, G. (2006). Identification of MAPKphosphorylation sites and their role in the localization and activity of hypoxia-inducible factor-1alpha. J. Biol. Chem. 281, 33095-33106.

Mylonis, I., Chachami, G., Paraskeva, E. and Simos, G. (2008). Atypical CRM1-dependent nuclear export signal mediates regulation of hypoxia-inducible factor-1alpha by MAPK. J. Biol. Chem. 283, 27620-27627.

Mylonis, I., Lakka, A., Tsakalof, A. and Simos, G. (2010). The dietary flavonoidkaempferol effectively inhibits HIF-1 activity and hepatoma cancer cell viabilityunder hypoxic conditions. Biochem. Biophys. Res. Commun. 398, 74-78.

Nath, B., Levin, I., Csak, T., Petrasek, J., Mueller, C., Kodys, K., Catalano, D.,

Mandrekar, P. and Szabo, G. (2011). Hepatocyte-specific hypoxia-inducible factor-1a is a determinant of lipid accumulation and liver injury in alcohol-induced steatosisin mice. Hepatology 53, 1526-1537.

Neuschwander-Tetri, B. A. (2010). Hepatic lipotoxicity and the pathogenesis ofnonalcoholic steatohepatitis: the central role of nontriglyceride fatty acid metabolites.Hepatology 52, 774-788.

Papadakis, A. I., Paraskeva, E., Peidis, P., Muaddi, H., Li, S., Raptis, L.,

Pantopoulos, K., Simos, G. and Koromilas, A. E. (2010). eIF2alpha Kinase PKRmodulates the hypoxic response by Stat3-dependent transcriptional suppression ofHIF-1alpha. Cancer Res. 70, 7820-7829.

Papandreou, I., Cairns, R. A., Fontana, L., Lim, A. L. and Denko, N. C. (2006). HIF-1 mediates adaptation to hypoxia by actively downregulating mitochondrial oxygenconsumption. Cell Metab. 3, 187-197.

Pescador, N., Villar, D., Cifuentes, D., Garcia-Rocha, M., Ortiz-Barahona, A.,

Vazquez, S., Ordonez, A., Cuevas, Y., Saez-Morales, D., Garcia-Bermejo, M. L.

et al. (2010). Hypoxia promotes glycogen accumulation through hypoxia inducible

factor (HIF)-mediated induction of glycogen synthase 1. PLoS ONE 5, e9644.

Peterfy, M., Phan, J., Xu, P. and Reue, K. (2001). Lipodystrophy in the fld mouse

results from mutation of a new gene encoding a nuclear protein, lipin. Nat. Genet. 27,

121-124.

Peterson, T. R., Sengupta, S. S., Harris, T. E., Carmack, A. E., Kang, S. A.,

Balderas, E., Guertin, D. A., Madden, K. L., Carpenter, A. E., Finck, B. N. et al.

(2011). mTOR complex 1 regulates lipin 1 localization to control the SREBP

pathway. Cell 146, 408-420.

Phan, J. and Reue, K. (2005). Lipin, a lipodystrophy and obesity gene. Cell Metab. 1,

73-83.

Qu, A., Taylor, M., Xue, X., Matsubara, T., Metzger, D., Chambon, P., Gonzalez,

F. J. and Shah, Y. M. (2011). Hypoxia-inducible transcription factor 2a promotes

steatohepatitis through augmenting lipid accumulation, inflammation, and fibrosis.

Hepatology 54, 472-483.

Rankin, E. B., Rha, J., Selak, M. A., Unger, T. L., Keith, B., Liu, Q. and Haase,

V. H. (2009). Hypoxia-inducible factor 2 regulates hepatic lipid metabolism. Mol.

Cell. Biol. 29, 4527-4538.

Rius, J., Guma, M., Schachtrup, C., Akassoglou, K., Zinkernagel, A. S., Nizet, V.,

Johnson, R. S., Haddad, G. G. and Karin, M. (2008). NF-kappaB links innate

immunity to the hypoxic response through transcriptional regulation of HIF-1alpha.

Nature 453, 807-811.

Semenza, G. L. (2011). Oxygen sensing, homeostasis, and disease. N. Engl. J. Med. 365,

537-547.

Tian, H., McKnight, S. L. and Russell, D. W. (1997). Endothelial PAS domain protein

1 (EPAS1), a transcription factor selectively expressed in endothelial cells. Genes

Dev. 11, 72-82.

Turkish, A. and Sturley, S. L. (2007). Regulation of triglyceride metabolism. I.

Eukaryotic neutral lipid synthesis: ‘‘Many ways to skin ACAT or a DGAT’’. Am. J.

Physiol. Gastrointest. Liver Physiol. 292, G953-G957.

Unger, R. H., Clark, G. O., Scherer, P. E. and Orci, L. (2010). Lipid homeostasis,

lipotoxicity and the metabolic syndrome. Biochim. Biophys. Acta 1801, 209-214.

Wouters, B. G. and Koritzinsky, M. (2008). Hypoxia signalling through mTOR and

the unfolded protein response in cancer. Nat. Rev. Cancer 8, 851-864.

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