Research Article
3485
Hypoxia causes triglyceride accumulation by
HIF-1-mediated stimulation of lipin 1 expression
Ilias Mylonis1, Hiroshi Sembongi2, Christina Befani1, Panagiotis Liakos1, Symeon Siniossoglou2 and
George Simos1,3,*
1
Laboratory of Biochemistry, Medical School, University of Thessaly, BIOPOLIS, Larissa 41110, Greece
Cambridge Institute for Medical Research, University of Cambridge, Wellcome Trust/Medical Research Council Building, Hills Road, Cambridge,
CB2 0XY, UK
3
Institute of Biomedical Research and Technology (BIOMED), 51 Papanastasiou Street, Larissa 41222, Greece
2
*Author for correspondence ([email protected])
Journal of Cell Science
Accepted 6 March 2012
Journal of Cell Science 125, 3485–3493
ß 2012. Published by The Company of Biologists Ltd
doi: 10.1242/jcs.106682
Summary
Adaptation to hypoxia involves hypoxia-inducible transcription factors (HIFs) and requires reprogramming of cellular metabolism that is
essential 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 by
induction of lipin 1, a phosphatidate phosphatase isoform that catalyzes the penultimate step in triglyceride biosynthesis, whereas lipin 2
remains unaffected. Hypoxic upregulation of lipin 1 expression involves predominantly HIF-1, which binds to a single distal hypoxiaresponsive element in the lipin 1 gene promoter and causes its activation under low oxygen conditions. Accumulation of hypoxic
triglycerides or lipid droplets can be blocked by siRNA-mediated silencing of lipin 1 expression or kaempferol-mediated inhibition of
HIF-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
Introduction
Exposure 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), VHLmediated 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 hypoxiaresponsive 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-1a
these 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 (Flügel et al.,
2007) or protein–protein interactions (Liu et al., 2007), MAPKdependent 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 (Boström 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-2a
in 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 fatty
liver dystrophic (fld) mouse and, conversely, overexpression of
lipin 1 in the adipose tissue of transgenic mice promotes obesity
(Péterfy et al., 2001; Phan and Reue, 2005). Moreover, lipin 1 as
well as lipin 2, a liver-enriched isoform, interact with and/or
modulate the activity of several transcription factors, including
members of the peroxisome proliferator-activated receptor
(PPAR) family and SREBP that control the expression of genes
involved in fatty acid and lipid metabolism (Donkor et al., 2009;
Finck et al., 2006; Koh et al., 2008; Peterson et al., 2011). Apart
of its apparently important role in adipogenesis, relatively little is
known about the function of lipin 1 in cells other than adipocytes.
Various stimuli such as glucocorticoids, insulin or fasting
alter lipin 1 expression and/or activity, regulation of which is
incompletely understood and involves transcriptional control,
post-translational modifications and compartmentalization (reviewed
by 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 accumulation
in cells of non-adipocyte origin, suggesting that lipin 1 is an
important mediator of the metabolic adaptation to hypoxia at the
cellular level. Moreover, inhibition of HIF-1 by pharmacological
or genetic means can suppress the hypoxic induction of lipin 1
and TG or LD production.
Results
Hypoxia induces lipin 1-dependent triglyceride
accumulation
To investigate the effect of hypoxia on lipid biosynthesis
we analysed human hepatoblastoma (Huh7) and cervical
adenocarcinoma (HeLaM) cells exposed to normoxia or
hypoxia (1% O2). As expected, hypoxia caused a robust
expression 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 types
hypoxia 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 by
determining the cellular TG content using a biochemical assay
(Fig. 1C). Also in this case, TG accumulation under hypoxia was
similar to the accumulation triggered by oleic acid.
Analysis of Huh7 cells by western blotting showed that in
addition 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 hepatic
isoform, remained unaffected indicating an isoform-specific effect
of 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 the
HIFs (Fig. 2B). Induction of lipin 1 by hypoxia was also confirmed
using immunofluorescence microscopy in both Huh7 (Fig. 2C) and
HeLaM (Fig. 2D) cells. The overexpression of endogenous lipin 1
under 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). Similar
images were obtained in HeLaM cells, in which, however,
cytoplasmic spots and nuclei were stained more intensely at early
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.
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, noncancer 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
Journal of Cell Science
HIF-1 regulates lipin 1
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.
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 siRNAmediated 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
3487
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.
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 HIF1a 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
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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).
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-1a
activity 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 gainof-function and loss-of-function experiments strongly suggest that
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 GFPpositive (+) 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.
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
Journal of Cell Science
HIF-1 regulates lipin 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).
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 39truncated 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.
3489
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.
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
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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 DNAprotein complexes isolated from hypoxically treated cells in
comparison to rabbit IgG immunoprecipitates or anti-HIF-1a
immunoprecipitates from normoxic cells (Fig. 8C,D). Therefore,
HIF-1a not only activates but also physically associates with the
Lpin1 promoter.
Discussion
We 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
Journal of Cell Science
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.
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, GFPHIF-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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HIF-1 regulates lipin 1
was also shown to involve HIF-1a (Nath et al., 2011). On the
other hand, in double knockout studies, hepatic steatosis caused
by liver-specific deficiency of pVHL could be avoided by
simultaneous inactivation of HIF-2a (Qu et al., 2011; Rankin
et al., 2009). However, in these animal models HIF-2 was mainly
associated with suppression of fatty acid oxidation, increased
lipid storage capacity, serum TG levels and inflammation. In our
study with human cells, HIF-2 also seems to be involved in lipin
1 regulation and LD accumulation but its role is minor compared
to HIF-1 and probably indirect since it requires more complex
regulatory elements. It, therefore, appears that HIF-1a and HIF2a may have overlapping or complementary but distinct roles in
the regulation of hepatic lipid metabolism and the development
of related pathologies such as alcoholic and non-alcoholic fattyliver disease (AFLD and NAFLD, respectively), with HIF-1
predominantly stimulating lipin 1 and triglyceride synthesis
under hypoxic or other stress conditions. HIF-1 has also been
implicated in cardiac steatosis by controlling expression of
PPARc, a principal transcriptional activator of lipid anabolism
(Krishnan et al., 2009). Lipin 1 also co-activates PPARc (Koh
et al., 2008), so the HIF-1–lipin-1 axis may serve to amplify not
only PAP1 activity but also the non-catalytic transcriptional lipin
1 functions.
In order for lipin 1 to exert its transcriptional role, it has to
translocate inside the nucleus. Although lipin 1 is predominantly
cytoplasmic in our tested cell lines under normoxia, hypoxic
treatment increases its nuclear pool especially after long-term
induction. A recent report has shown that nuclear localization of
lipin 1 is regulated by nutrient- and growth factor-responsive
kinase mTOR complex 1 (mTORC1)-mediated phosphorylation
(Peterson et al., 2011). Inhibition of mTORC1 causes normoxic
redistribution of lipin 1 from the cytoplasm to the nucleus, which
in turn reduces the nuclear protein levels of SREBP-1 (Peterson
et al., 2011), a transcription factor regulating many lipogenic
genes, including lipin 1 (Ishimoto et al., 2009). Therefore, our
data showing nuclear accumulation of lipin 1 and a decrease of
lipin 1 mRNA expression at later points of hypoxic treatment
could be due to inhibition of mTORC1 that is known to occur
after prolonged exposure to hypoxia (Wouters and Koritzinsky,
2008). The physiological relevance of lipin 1 hypoxic nuclear
migration could thus be justified by the operation of the
following negative feedback mechanism. After the initial HIF1-mediated induction of lipin 1 and the subsequent TG
accumulation, inhibition of mTOR by prolonged hypoxia
causes translocation of lipin 1 inside the nucleus, where, by
suppressing SREBP-1 function, it reduces expression of its own
and other lipogenic genes and blocks further TG de novo
synthesis.
The idea that cells in a stressful situation, such as oxygen
deprivation, respond by stimulating, even transiently, an anabolic
process, e.g. lipid biosynthesis, is counterintuitive but not without
precedent. Hypoxia also stimulates glycogen synthesis in a HIF1-dependent way (Brahimi-Horn et al., 2011; Fakas et al., 2011;
Pescador et al., 2010). Apparently, exposure of cells to mild
hypoxia may serve as a way of sensing ensuing ischemia/nutrient
limitation and, while it is still available, glucose is stored for later
use as anaerobic source of energy. However, this may not be the
case for lipids since they can only produce energy through
oxidative phosphorylation, a process drastically inhibited by
severe hypoxia. A more likely explanation for storing
triglycerides in the form of lipid droplets under hypoxia could
3491
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 Methods
Plasmids
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 introduced
into pUC19 vector, sequenced and subcloned into pGL3-basic vector (Promega,
Madison, WI, USA) to construct pGL3-HRE1–8. Truncated promoter forms, pGL3HRE2–8 (22198), pGL3-HRE5–8 (21297), pGL3-HRE7–8 (2602) and pGL3HRE8 (2445) were constructed by using as template the pUC19-HRE1–8 plasmid,
and an appropriate set of primers listed in supplementary material Table S1. The
occurring PCR fragments were then subcloned into the pGL3-basic vector and
sequenced. To construct HRE mutant plasmid pGL3-mHRE1–8, three nucleotide
substitutions (CCGTG to CAACG) were introduced by PCR using hL1pHRE1mF1
and hL1pHRE1mR1 primers listed in supplementary material Table S1. Plasmids
pGL3-HRE1 (23298 to 22588), pGL3-HRE1–4 (23298 to 21585) and their
respective mutant forms, pGL3-mHRE1 and pGL3-mHRE1–4, were constructed by
digesting pGL3-HRE1–8 with BglII, to yield fragment HRE1, or HindIII, to yield
HRE1–4, and subcloning to the empty pGL3-vector. pEGFP-HIF-1a, pCMVFLAG-MTD and pCMV-FLAG-MTD-SA plasmids were previously described
(Mylonis et al., 2008). To construct pEGFP-HIF-2a, full-length HIF-2a cDNA was
obtained from pcDNA-HIF-2a, kindly provided by S. L. McKnight [University of
Texas Southwestern Medical Center (Tian et al., 1997)].
Cell culture, transfection and reporter gene assays
Human HeLaM and Huh7 cells were cultured in DMEM (Biosera) containing
10% FCS and 100 U/ml penicillin-streptomycin (Gibco BRL). Human bronchial
smooth muscle cells (Lonza, Switzerland) were cultured in DMEM F-12
containing 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) or
with kaempferol (10–50 mM; Sigma). For oleic acid treatment, the medium was
supplemented with 0.4 mM oleic acid (Sigma) bound to 1.5% bovine serum
albumin. 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 Life
Sciences, Pencoed, UK). Transient transfections and reporter gene assays were
performed as described previously (Mylonis et al., 2008).
Western blot and immunofluorescence
Antibodies used for western blotting, immunofluorescence and chromatin
immunoprecipitation were: affinity purified rabbit polyclonal antibodies against
HIF-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 or
tubulin (Millipore, Billerica, MA, USA). Analysis of total cellular proteins by
immunoblotting and immunofluorescence microscopy were carried out as
3492
Journal of Cell Science 125 (14)
previously described (Grimsey et al., 2008; Lyberopoulou et al., 2007; Mylonis
et al., 2008; Mylonis et al., 2006). To visualize lipid droplets after usual
immunofluorescence 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 for
5 minutes to facilitate the formation of an even suspension (Al-Anzi and
Zinn, 2010) and 20 ml aliquots were tested using a biochemical triglyceride
determination 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 sequences
shown 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-1a
was 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).
Journal of Cell Science
RNA extraction and Real-Time PCR
Total RNA from Huh7 cells was isolated using the Trizol reagent (Invitrogen) and
cDNA was synthesized with the High Capacity Reverse Transcription kit (Applied
Biosystems, Carlsbad, CA). Real-time PCR was performed with SYBR GreenER
qPCR SuperMix Universal (Invitrogen) in a MiniOpticon instrument (Bio-Rad).
The mRNAs encoding lipin 1, lipin 2 and VEGF were amplified using primers
listed in supplementary material Table S1. Each sample was assayed in triplicate
for both target and internal control. Relative quantitative gene expression was
calculated using the DDCt method and presented as relative units.
Chromatin immunoprecipitation
Huh7 cells were treated with 1.1% formaldehyde for 30 minutes, quenched with
125 mM glycine for 5 min and collected by centrifugation. Following steps were
performed as previously described (Braliou et al., 2008) using a polyclonal antiHIF-1a antibody (Lyberopoulou et al., 2007) or rabbit IgG. The region 22916 to
22686 of the hLPIN1 promoter was amplified from immunoprecipitated
chromatin or input samples using primers listed in supplementary material Table
S1. The 230 bp product was analyzed by agarose gel electrophoresis or quantified
by 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 Ct
of 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.
Acknowledgements
We 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 providing
hBMS cells and instructions for their culture.
Funding
This study was supported by the National Strategic Reference
Framework/National Action ‘‘Cooperation’’ [grant number 09SYN12-682] co-funded by the Greek Government and the European
Regional Development Fund (G.S.); A Medical Research Council
Senior Fellowship [grant number G0701446 to S.S.]; and a European
Molecular 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
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