Interruption of triacylglycerol synthesis in the
endoplasmic reticulum is the initiating event for saturated
fatty acid-induced lipotoxicity in liver cells
Michalis D. Mantzaris
1
, Epameinondas V. Tsianos
2
and Dimitrios Galaris
1
1 Laboratory of Biological Chemistry, University of Ioannina Medical School, Greece
2 First Division of Internal Medicine and Hepato-gastroenterology Unit, University of Ioannina Medical School, Greece
Introduction
Dietary habits in the Western world have changed dras-
tically during the last few decades, and this change cor-
relates with increasing levels of obesity, implying that
diet may be associated with the development of insulin
resistance, type 2 diabetes, cardiovascular disease and
other pathologies in the general population [1]. Con-
sumption of food rich in fat causes qualitative and
quantitative changes in serum free fatty acid (FFA) lev-
els, and increases the rate of uptake and accumulation
of lipids in nonadipose tissues such as the liver, which
is the main lipid-metabolizing organ. Inappropriate
accumulation of excess lipids in liver cells in the form
of lipid droplets has been proposed to lead to dysfunc-
tion of hepatocytes and, consequently, to serious path-
ological complications [2,3]. Nonalcoholic fatty liver
disease (NAFLD) is a term used to characterize a spec-
trum of pathological changes ranging from simple fatty
infiltration (steatosis) to hepatic steatosis accompanied
Keywords

ulum, apparently because of the formation of a pool of oversaturated inter-
mediates, represents the key initiating event in the mechanism of saturated
fatty acid-induced lipotoxicity.
Abbreviations
ACS, long-chain acyl-CoA synthetase; ATF4, activating transcription factor 4; BrdU, bromodeoxyuridine; CHOP, CCAAT ⁄ enhancer-binding
protein homologous protein; DAG, diacylglycerol; ER, endoplasmic reticulum; eIF2a, eukaryotic translation initiation factor 2a;
FITC, fluorescein isothiocyanate; FFA, free fatty acid; JNK, c-Jun N-terminal kinase; NAFLD, nonalcoholic fatty liver disease; PERK,
RNA-dependent protein kinase-like endoplasmic reticulum eukaryotic initiation factor-2a kinase; PI, propidium iodide; SD, standard deviation;
SFA, saturated fatty acid; TAG, triacylglycerol; TrC, triacsin C; UFA, unsaturated fatty acid.
FEBS Journal 278 (2011) 519–530 ª 2010 The Authors Journal compilation ª 2010 FEBS 519
by inflammation, fibrosis, and cirrhosis (nonalcoholic
steatohepatitis) [4,5]. Despite the high prevalence of
NAFLD and its potential for serious complications,
the underlying molecular mechanisms that determine
the progression to liver damage remain poorly under-
stood and need further investigation.
A number of recent in vitro and in vivo studies have
shown that different forms of fatty acids exert remark-
ably different effects. Exposure of a variety of cell
types, including hepatocytes, to long-chain saturated
fatty acids (SFAs) led to increased expression of proin-
flammatory cytokines, inhibition of insulin signaling,
induction of endoplasmic reticulum (ER) stress, and
promotion of cell death, mainly by apoptosis [6–12].
On the other hand, unsaturated fatty acids (UFAs)
were not toxic at the same concentrations and, in addi-
tion, their presence protected cells from SFA-induced
effects [6,13–16]. A protective role for endogenously
generated UFAs was also indicated by in vivo experi-
ments using genetically modified mice bearing an inacti-

oleate (18:1, cis), induced a transient inhibition of cell
proliferation during the first 24 h (Fig. 1A). This
observation was also confirmed by analysis of bromo-
deoxyuridine (BrdU) incorporation into DNA, which
decreased by more than 50% after 24 h of stearate
treatment. However, cells regained their normal prolif-
eration capacity at longer incubation periods, whereas
coadministration of oleate (0.3 mm) prevented the
AB
CD
EF
Fig. 1. Oleate prevents (SA) stearate-induced cytotoxicity. (A)
HepG2 cells (1.0 · 10
5
) were seeded in 24-well plates and cultured
for 24 h before being treated with vehicle (
), 300 lM stearate ( ),
300 l
M oleate (OA) ( ), or a combination of the two ( ). At the
indicated time points, cells were harvested, and viable cells were
counted (Trypan blue exclusion). (B) Cells were treated as above,
except that for the last 8 h of treatment they were supplemented
with BrdU (100 l
M). BrdU incorporation into DNA was detected by
using an antibody against BrdU as described in Experimental proce-
dures. Results are expressed as percentages of the respective con-
trols (Ctrl) (*P < 0.05). (C, D) The conditions were exactly as in (A)
and (B), respectively, except that 600 l
M of each fatty acid was
used. (E) Cells were treated with stearate (600 l

not shown).
Stearate treatment induces ER stress and
mitochondrial-mediated apoptosis
As shown in Fig. 2, treatment of HepG2 cells with
0.6 mm stearate led to the appearance of condensed
and fragmented nuclei (Fig. 2A,B), hypodiploid sub-
G
1
DNA (Fig. 2C), DNA internucleolar fragmentation
(Fig. 2D), caspase-3 cleavage (Fig. 2E), and the
appearance of cytochrome c in the cytosol (Fig. 2F),
which are clear characteristics of mitochondrial-medi-
ated apoptotic cell death. In all cases, oleate was not
toxic by itself, and its coadministration with stearate
prevented the appearance of these apoptotic markers.
Different Bcl-2 family members serve as proapop-
totic and antiapoptotic mitochondrial regulators under
certain circumstances [21,22]. As shown in Fig. 3A, the
relative amount of the antiapoptotic protein Bcl-2 was
gradually increased in HepG2 cells during the initial
22 h of stearate treatment, but this increase was inter-
rupted thereafter, and the Bcl-2 concentration stabi-
lized at a somewhat lower level. On the other hand,
the proapoptotic Bcl-2-like protein Bax was activated,
as indicated by its translocation from the cytosolic
fraction to the mitochondrial fraction following stea-
rate administration (Fig. 3B). The presence of oleate
inhibited this translocation, indicating the involvement
of Bax activation in stearate-induced mitochondrial
destabilization and apoptosis.

prevented stearate-induced phosphorylation of JNK
when the two agents were coadministered (Fig. 4C).
Taken together, these results indicate that the location
of the protective action of oleate was upstream of ER
stress activation.
Whether the protection offered by oleate was specific
for SFA-induced ER stress or represented a more gen-
eral phenomenon was also investigated. The presence
of oleate was unable to prevent the toxic effects
induced by thapsigargin or tunicamycin (two classical
ER stress inducers), indicating that oleate is not a gen-
eral inhibitor of the ER stress response (results not
shown).
Stearate treatment interrupts TAG synthesis and
lipid droplet accumulation
In addition to its role in proper protein folding, the
ER is responsible for lipid synthesis. In particular,
excess availability of FFAs, as is the case in the pres-
ent experimental model, leads to increased formation
of TAGs, which are either released from the cells as
very low density lipoprotein or stored in the cytosol as
lipid droplets. It was observed that the accumulation
of lipid droplets was efficient in oleate-treated cells,
whereas stearate-treated cells contained fewer and
smaller lipid droplets after 24 h of treatment (Fig. 5A).
Moreover, coadministration of oleate restored the
capacity of stearate-treated cells to accumulate lipid
droplets. This observation was further confirmed by
M. D. Mantzaris et al. Saturated fatty acid-induced lipotoxicity
FEBS Journal 278 (2011) 519–530 ª 2010 The Authors Journal compilation ª 2010 FEBS 521

with FFAs for time periods of 3, 6, 12, 24 and 36 h,
before staining of the accumulated neutral lipids with
Nile red and analysis of the fluorescence intensity of
individual cells by flow cytometry. Cell fluorescence
increased progressively in oleate-treated and oleate
plus stearate-treated cells, whereas it was significantly
lower in stearate-treated cells during the first 3 and 6 h
of treatment (Fig. 5B). Interestingly, the fluorescence
in stearate-treated cells started to decrease gradually at
exposure times longer than 6 h, giving rise to a distinct
cell population with basal levels of fluorescence
(Fig. 5B). After 36 h, almost the entire cell population
was devoid of lipid droplets.
It is obvious from these results that TAG synthesis
was initially hindered following stearate administration
and was completely interrupted at longer incubation
periods. Interestingly, the interruption of TAG synthe-
sis preceded the appearance of toxic effects, supporting
the notion that it constitutes the initiating event in the
process of lipotoxicity.
Stearate has to be activated in order to be toxic
The first enzyme involved in metabolism of FFAs after
their uptake into liver cells is the long-chain acyl-CoA
synthetase (ACS), which activates fatty acids by link-
ing them to coenzyme A. As shown in Fig. 6, triac-
sin C (TrC), a specific competitive inhibitor of ACS
[24,25], was not toxic by itself, whereas it inhibited the
accumulation of lipid droplets following exposure of
cells to either stearate or oleate (Fig. 6A,B). At the
same time, TrC protected cells from stearate-induced

the phosphorylation of eIF2a and the expression of ATF4 and
CHOP proteins were examined by western blotting with specific
antibodies. (B) Western blot analysis of the expression of CHOP in
total cell extracts prepared from cells treated with 600 l
M stearate,
600 l
M oleate (OA) or a combination of the two (SA ⁄ OA) for 36 h.
(C) The intensity of phosphorylation of JNK (p-JNK) and the amount
of the total protein was examined by western blot analysis in total
cell extracts derived from cells treated as in (A). Where indicated
thapsigargin (Thap)-treated cells (2 l
M for 24 h) were used as posi-
tive controls (Ctrl).
M. D. Mantzaris et al. Saturated fatty acid-induced lipotoxicity
FEBS Journal 278 (2011) 519–530 ª 2010 The Authors Journal compilation ª 2010 FEBS 523
TAG synthesis. In addition, these results show that the
properties of the metabolic intermediates of stearate
and oleate must be fundamentally different.
Discussion
The results presented in this investigation, in agree-
ment with previously reported observations, revealed
fundamentally different effects of SFAs and UFAs on
liver cells [8,11–13]. In an attempt to identify the key
event(s) responsible for these differences, we examined
the main steps involved, following the uptake of satu-
rated and unsaturated FFAs into the cells. After their
internalization, FFAs are converted to fatty acyl-CoA,
a reaction catalyzed by ACS. Fatty acyl-CoAs are acti-
vated forms of fatty acids that can be either oxidized
in mitochondria or utilized in the ER as substrates for

Ctrl
Ctrl
Ctrl
Ctrl
SA OA SA/OA
A
B
Fig. 5. Stearate (SA) supplementation interrupts lipid droplet accumulation. HepG2 cells were exposed to 600 lM stearate, 600 lM oleate
(OA) or a combination of the two at 600 l
M each. (A) To identify lipid droplets, cells treated with FFA media for 24 h were stained with Nile
red and analyzed by confocal microscopy. (B) Graphs showing the distribution of Nile red fluorescence intensity of individual cells at 3, 6, 12,
24 and 36 h were obtained by flow cytometric analysis in the FL1 channel (logarithmic scale). Control (Ctrl) cells (blue line), stearate-treated
cells (green line), oleate (OA)-treated cells (black line) and cells supplemented with both fatty acids (red line) are shown. These experiments
were repeated two more times, with essentially the same results.
Fig. 6. Acyl-CoA formation is necessary for stearate-induced toxicity. HepG2 cells were exposed for 48 h to vehicle (Ctrl), 600 lM of stearate
(SA), 600 l
M oleate (OA) or a combination of the two (SA ⁄ OA) in the absence or presence of 0.5 lM TrC, a specific inhibitor of ACS. (A)
Cells were stained with Nile red and analyzed by confocal microscopy. Representative photographs show inhibition of FFA-induced lipid drop-
let formation by TrC. (B) Quantitation of Nile red fluorescence by flow cytometry. Bars represent the mean fluorescence value of each distri-
bution ± SD of duplicate measurements from two independent experiments (*P < 0.05 versus control;
#
P < 0.05 versus TrC-untreated
cells). (C) Cells were harvested, and cell numbers were assessed by Trypan blue exclusion. Each bar represents the mean ± sd from tripli-
cate measurements (*P < 0.05). (D) Annexin V–FITC binding and PI staining were performed in order to assess cell death. Fluorescence
was analyzed by flow cytometry in 10
4
cells per sample. (E) Cells were exposed to typical ER stressors, thapsigargin (Tg, 2 lM) or tunicamy-
cin (Tm, 3 l
M), for 24 h, in the presence or absence of 0.5 lM TrC. Total cell extracts were isolated, and CHOP expression was examined
by western blot analysis.

C
M. D. Mantzaris et al. Saturated fatty acid-induced lipotoxicity
FEBS Journal 278 (2011) 519–530 ª 2010 The Authors Journal compilation ª 2010 FEBS 525
or tunicamycin-induced ER stress (Fig. 6E), thus
excluding the possibility of nonspecific inhibition of
ER stress. When the available acyl-CoAs are in excess,
they are channeled towards TAG synthesis.
The main findings of the present investigation were
the observations that TAG synthesis in liver cells
exposed to excess stearate was interrupted, and that
this interruption preceded the appearance of toxic
effects (Fig. 5A,B). In sharp contrast, oleate-treated
cells, which continued to proliferate normally, were
able to produce TAGs continuously and accumulate
them in the form of lipid droplets. Moreover, coad-
ministration of oleate restored the ability of stearate-
treated cells to synthesize TAGs and prevented cell
toxicity. These findings are in agreement with previous
observations from the Schaffer group, indicating
increased incorporation of palmitate (16:0) into the
TAG pool only in the presence of oleate [18].
The above results raise two main questions: (a) what
is the cause of TAG synthesis inhibition, and (b) what
is the exact nature of the events that ultimately lead to
cell toxicity?
Regarding the first question, it is obvious that one
or more steps (following acyl-CoA formation) in the
cascade of TAG formation that take place in ER
membranes are defective in SFA-treated but not in
UFA-treated cells. It has been previously shown that

[9,20,35]. Moffitt et al. [20] suggested that accumula-
tion in the ER lumen of oversaturated TAGs, which
cannot be further processed, because of their inappro-
priate physicochemical properties (high melting point),
is the main cause of toxicity. This proposal, however,
is not consistent with the disappearance of lipid drop-
lets from stearate-treated cells, as observed in this
investigation (Fig. 5B).
The ER is the site of synthesis of all secretory pro-
teins and resident proteins of the membrane system,
and any perturbation that compromises the protein-
folding capacity of the organelle can lead to ER stress
[36–38]. ER stress is a general, integrated stress
response displayed by mammalian cells. This response
can be divided in two phases according to the intensity
and the duration of the stress. An initial adaptive
response culminates in the temporary inhibition of
protein synthesis, providing cells with the opportunity
to recover and restore normal homeostasis. The data
presented in this work demonstrate that cells exposed
to stearate are moved initially towards such an adap-
tive state, as indicated by the transient inhibition of
cell proliferation (Fig. 1A,B) and the early phosphory-
lation of eIF2a (Fig. 4A). When the stress is more
intensive and prolonged, secondary events, such as
ATF4 and CHOP protein expression and JNK activa-
tion, were induced, leading ultimately to cell death by
apoptosis (Fig. 4A,C). Prolonged ER stress and JNK
activation, as observed in this study, usually stimulate
apoptosis by several pathways, including the transloca-

Experimental procedures
Cell culture and treatment
Human hepatocellular HepG2 (ATCC, HB-8065) and
Huh7 (Health Science Research Resources Bank,
JCRB0403, Osaka, Japan) carcinoma cells were grown in
DMEM containing 10% heat-inactivated fetal bovine
serum, 2 mm glutamine, 100 UÆmL
)1
penicillin, and
100 ngÆmL
)1
streptomycin, at 37 °C in air with 5% CO
2
.
Cells were seeded and left under normal conditions for 24 h
before any further treatment. Stock solutions of FFAs
(100 mm) were prepared in isopropanol by heating to
70 °C, and the desired concentrations were prepared in
growth medium supplemented with BSA, as described pre-
viously [41]. Briefly, a 5% (w ⁄ v) BSA solution in DMEM
was filtered and mixed with the fatty acid stock solution,
giving a concentration of 5 mm FFA (FFA ⁄ BSA molar
ratio of 6.6 : 1). The solution was left for 30 min at 50 ° C,
and diluted with DMEM, giving the desired concentrations.
Estimation of cell viability
Following FFA treatment, cell numbers were assessed by
Trypan blue exclusion. Floating and attached cells were
Fig. 7. Schematic representation of the molecular events that take place following exposure of liver cells to FFAs. Interruption of TAG syn-
thesis in conditions of excess availability of SFAs is the key point in the molecular mechanism of SFA-induced lipotoxicity. It is suggested
that creation of a pool of oversaturated lipid intermediates determines whether TAG formation will proceed normally or whether the process

expressed as a percentage of the total cell number.
Detection of lipid accumulation
Lipid droplet accumulation was detected by Nile red stain-
ing as previously described [42]. Cell imaging for Nile red
staining was performed by confocal microscopy. Quantifica-
tion of lipid droplets was performed by flow cytometric
analysis of the distribution of Nile red fluorescence in indi-
vidual cells. Briefly, cells were seeded in 24-well plates onto
11-mm glass coverslips for confocal microscopy, or in six-
well plates for flow cytometry. After FFA treatment, cells
were fixed with 3.7% paraformaldehyde for 10 min, washed
twice, and stained with Nile red (Sigma) solution (final con-
centration, 2 lgÆmL
)1
). Samples were kept for 45 min in
the dark at 37 °C to allow equilibrium with the dye. Cover-
slips were mounted in Mowiol, and viewed with a
Leica TCS-SP scanning confocal microscope, equipped with
an argon ⁄ krypton laser and Leica TCS software. Flow
cytometric analysis (15 000 events per sample) was carried
out with a CyFlow ML (Partec) equipped with an argon
laser, in the FL1 channel (logarithmic scale).
Determination of TAG levels
Cells seeded in six-well plates were treated with FFA med-
ium for 24 h. Cells were harvested, and lipids were
extracted twice with CHCl
3
⁄ MeOH ⁄ ddH
2
O (1 : 1 : 0.9).

PI
for 20 min, cells were analyzed by flow cytometry.
Preparation and analysis of DNA and protein
extracts
After the appropriate treatment, cellular DNA was isolated
from 3 · 10
6
cells per sample, and analyzed for internucleo-
lar fragmentation by agarose gel electrophoresis. Prepara-
tion of mitochondrial and cytosolic fractions was achieved
by differential centrifugation, as described previously [45].
For western blot analysis, cell lysates (40–50 lg of protein)
were subjected to SDS ⁄ PAGE, and the separated proteins
were demonstrated by immunoblotting after being trans-
ferred to nitrocellulose membranes. Antibodies against
cytochrome c (sc-13156), Bax (2D2, sc-20067), Bcl-2(100)
(sc-509), ATF4 (sc-200) and CHOP ⁄ GADD153 (R-20,
sc-793) were from Santa Cruz Biotechnology. Antibodies
against phospho-JNK (#9251), total JNK (#9252) and
phospho-eIF2a (#9721) were from Cell Signaling. Horse-
radish peroxidase-conjugated antibody against caspase-3
(#610325) was from BD Pharmigen, and antibody against
b-actin (A5441) was from Sigma.
Statistical analysis
All data are expressed as the mean ± standard deviation
(SD). Differences between groups were compared by one-
way ANOVA followed by a post hoc Bonferroni correction
test for multiple comparisons, using originpro 8 software
(OriginLab). Differences were considered to be statistically
significant at P < 0.05.

8 Akazawa Y, Cazanave S, Mott JL, Elmi N, Bronk SF,
Kohno S, Charlton MR & Gores GJ (2010) Palmitol-
eate attenuates palmitate-induced Bim and PUMA
up-regulation and hepatocyte lipoapoptosis. J Hepatol
52, 586–593.
9 Borradaile NM, Han X, Harp JD, Gale SE, Ory DS &
Schaffer JE (2006) Disruption of endoplasmic reticulum
structure and integrity in lipotoxic cell death. J Lipid
Res 47, 2726–2737.
10 Wang D, Wei Y & Pagliassotti MJ (2006) Saturated
fatty acids promote endoplasmic reticulum stress and
liver injury in rats with hepatic steatosis. Endocrinology
147, 943–951.
11 Li Z, Berk M, McIntyre TM, Gores GJ & Feldstein AE
(2008) The lysosomal–mitochondrial axis in free fatty
acid-induced hepatic lipotoxicity. Hepatology 47, 1495–
1503.
12 Jung TW, Lee YJ, Lee MW & Kim SM (2009) Full-
length adiponectin protects hepatocytes from palmitate-
induced apoptosis via inhibition of c-Jun NH2 terminal
kinase. FEBS J 276, 2278–2284.
13 Wei Y, Wang D, Topczewski F & Pagliassotti MJ
(2006) Saturated fatty acids induce endoplasmic reticu-
lum stress and apoptosis independently of ceramide in
liver cells. Am J Physiol Endocrinol Metab 291, E275–
E281.
14 Lai E, Bikopoulos G, Wheeler MB, Rozakis-Adcock M
& Volchuk A (2008) Differential activation of ER stress
and apoptosis in response to chronically elevated free
fatty acids in pancreatic beta-cells. Am J Physiol Endo-

1829.
21 Chipuk JE & Green DR (2008) How do BCL-2 proteins
induce mitochondrial outer membrane permeabilization?
Trends Cell Biol 18, 157–164.
22 Youle RJ & Strasser A (2008) The BCL-2 protein fam-
ily: opposing activities that mediate cell death. Nat Rev
Mol Cell Biol 9, 47–59.
23 Malhi H & Gores GJ (2008) Cellular and molecular
mechanisms of liver injury. Gastroenterology 134, 1641–
1654.
24 Vessey DA, Kelley M & Warren RS (2004) Character-
ization of triacsin C inhibition of short-, medium-,
and long-chain fatty acid:CoA ligases of human liver.
J Biochem Mol Toxicol 18, 100–106.
25 Tomoda H, Igarashi K & Omura S (1987) Inhibition of
acyl-CoA synthetase by triacsins. Biochim Biophys Acta
921, 595–598.
26 Yen CL, Stone SJ, Koliwad S, Harris C & Farese RV
Jr (2008) Thematic review series: glycerolipids. DGAT
enzymes and triacylglycerol biosynthesis. J Lipid Res
49, 2283–2301.
M. D. Mantzaris et al. Saturated fatty acid-induced lipotoxicity
FEBS Journal 278 (2011) 519–530 ª 2010 The Authors Journal compilation ª 2010 FEBS 529
27 Guo W, Huang N, Cai J, Xie W & Hamilton JA (2006)
Fatty acid transport and metabolism in HepG2 cells.
Am J Physiol Gastrointest Liver Physiol 290, G528–
G534.
28 Bruce JS & Salter AM (1996) Metabolic fate of oleic
acid, palmitic acid and stearic acid in cultured hamster
hepatocytes. Biochem J 316 (Pt 3), 847–852.

Rev Mol Cell Biol 8, 519–529.
37 Zhang K & Kaufman RJ (2008) From endoplasmic-
reticulum stress to the inflammatory response. Nature
454, 455–462.
38 Yoshida H (2007) ER stress and diseases. FEBS J 274,
630–658.
39 McCullough KD, Martindale JL, Klotz LO, Aw TY &
Holbrook NJ (2001) Gadd153 sensitizes cells to
endoplasmic reticulum stress by down-regulating Bcl2
and perturbing the cellular redox state. Mol Cell Biol
21, 1249–1259.
40 Puri P, Mirshahi F, Cheung O, Natarajan R, Maher
JW, Kellum JM & Sanyal AJ (2008) Activation and
dysregulation of the unfolded protein response in non-
alcoholic fatty liver disease. Gastroenterology 134,
568–576.
41 Karaskov E, Scott C, Zhang L, Teodoro T, Ravazzola
M & Volchuk A (2006) Chronic palmitate but not
oleate exposure induces endoplasmic reticulum stress,
which may contribute to INS-1 pancreatic beta-cell
apoptosis. Endocrinology 147, 3398–3407.
42 Gubern A, Casas J, Barcelo-Torns M, Barneda D, de la
Rosa X, Masgrau R, Picatoste F, Balsinde J, Balboa
MA & Claro E (2008) Group IVA phospholipase A2 is
necessary for the biogenesis of lipid droplets. J Biol
Chem 283, 27369–27382.
43 Darzynkiewicz Z, Bruno S, Del Bino G, Gorczyca W,
Hotz MA, Lassota P & Traganos F (1992) Features of
apoptotic cells measured by flow cytometry. Cytometry
13, 795–808.