Download Hepatic triacylglycerol synthesis and secretion: DGAT2 as the link

Biochem. J. (2013) 451, 1–12 (Printed in Great Britain)
1
doi:10.1042/BJ20121689
REVIEW ARTICLE
Hepatic triacylglycerol synthesis and secretion: DGAT2 as the link between
glycaemia and triglyceridaemia
Victor A. ZAMMIT1
Division of Metabolic and Vascular Health, Warwick Medical School, University of Warwick, Coventry KA4 7AL, U.K.
The liver regulates both glycaemia and triglyceridaemia. Hyperglycaemia and hypertriglyceridaemia are both characteristic
of (pre)diabetes. Recent observations on the specialised role of
DGAT2 (diacylglycerol acyltransferase 2) in catalysing the de
novo synthesis of triacylglycerols from newly synthesized fatty
acids and nascent diacylglycerols identifies this enzyme as
the link between the two. This places DGAT2 at the centre
of carbohydrate-induced hypertriglyceridaemia and hepatic
steatosis. This function is complemented, but not substituted for,
by the ability of DGAT1 to rescue partial glycerides from complete
hydrolysis. In peripheral tissues not normally considered to be
lipogenic, synthesis of triacylgycerols may largely bypass DGAT2
except in hyperglycaemic/hyperinsulinaemic conditions, when
induction of de novo fatty acid synthesis in these tissues may
contribute towards increased triacylglycerol secretion (intestine)
or insulin resistance (adipose tissue, and cardiac and skeletal
muscle).
INTRODUCTION
of skeletal muscle, is the greatest contributor towards wholebody insulin resistance [6–9]. The term ‘lipotoxicity’ has
been applied to this phenomenon, not because TAG itself is
deleterious, but because the intermediates of TAG synthesis
and/or lipolysis are highly biologically active. For example,
long-chain acyl-CoA are membrane-active, and can affect gene
expression and ion channel activity (see [10]). Together with
DAGs (diacylgycerols), they have been suggested to cause
insulin resistance by activating PKC (protein kinase C) isoforms
whose serine phosphorylation of insulin signalling cascade
components results in the attenuation of insulin signalling [11–
15]. Similarly, intracellular levels of ceramide, which can be
synthesized de novo from palmitoyl-CoA, are also known to
correlate with increased insulin resistance in muscle [16]. Fatty
acids and ceramides also both feed into the synthesis of proinflammatory pathways [e.g. NF-κB (nuclear factor κB)] and the
synthesis of pro-insulin resistance factors [e.g. TNFα (tumour
necrosis factor α)]. Therefore the extent to which ectopic TAG
content in tissues is associated with an excessive supply of fatty
acids tends to be a feature of metabolic pathological conditions,
with one notable exception (see below). Lipotoxicity may not be
the only mechanism that is involved in the aetiology of insulin
resistance, but multiple other mechanisms {e.g. generation of
ROS (reactive oxygen species) and ER (endoplasmic reticulum)
stress [17]} may be related to, or overlap with, it [15].
TAGs (triacylglycerols; also known as triglycerides) perform
multiple functions in mammalian cells and in the traffic of
substrates between tissues. They are a major source of dietary
energy, and the intricate mechanisms that have been developed
to enable their digestion, uptake, distribution and storage attest to
their importance as the ultimate high-capacity form of energy
storage in vertebrates. White adipose tissue is specialized for
their storage, and it is therefore not surprising that it should
be the origin of multiple signals (adipokines) that feedback to
the brain the longer-term energy status of the organism, and
also act directly on other tissues. However, lipid droplets (of
which TAG is a major component) exist in most cell types where
they reflect the difference between the rate at which fatty acids
reach tissues, or are synthesized therein, and the rate at which
they are metabolized. This may occur through oxidation (e.g.
in muscle), or, as in the liver and intestine, through packaging
within TRLs (TAG-rich lipoproteins) which are secreted. TAG
that is accumulated in tissues other than adipose is referred to as
‘ectopic’ and can be used as a local readily available source of
energy for oxidative tissues (e.g. cardiac muscle, type-1 skeletal
muscle fibres). But expansion of ectopic lipid levels tends to be
associated with metabolic dysregulation and, in particular, altered
glucose–fatty acid interactions. Thus hepatic steatosis results
in the deterioration of liver function, the possibility of hepatic
insulin resistance, and the development of inflammation, nonalcoholic steatohepatitis and associated morbidities [1]. Increased
cardiac intramyocytic TAG is associated with cardiomyopathy
and heart failure [2–5]. Elevated IMTG (intramyocellular TAG)
in skeletal muscle, particularly in type-2 fibres, is associated
with muscle insulin resistance which, owing to the large mass
Key words: de novo fatty acid synthesis, diacylglycerol acyltransferase, heart, hypertriglyceridaemia, intestine, lipogenesis, liver,
muscle.
TAG SYNTHESIS AND SECRETION
Most TAG formation can occur in one of two ways. First,
through re-esterification of partial glycerides [DAG and MAG
(monoacylglycerol)] that themselves arise from partial hydrolysis
Abbreviations used: apoB, apolipoprotein B; C/EBP, CCAAT/enhancer-binding protein; ChREBP, carbohydrate-responsive-element-binding protein;
DAG, diacylglycerol; DGAT, DAG acyltransferase; ER, endoplasmic reticulum; ERM, ER membrane; FAS, fatty acid synthase; FATP1, fatty acid transport
protein 1; GPAT, glycerol-3-phosphate acytransferase; IMTG, intramyocellular triacylglycerol; LD, lipid droplet; LDL, low-density lipoprotein; LPL, lipoprotein
lipase; MAG, monoacylglycerol; MGAT, MAG acyltransferase; MTP, microsomal transfer protein; SCD1, stearoyl-CoA desaturase 1; SREBP-1c, sterolregulatory-element-binding protein 1c; TAG, triacylglycerol; TM, transmembrane; TRL, TAG-rich lipoprotein; VLDL, very-LDL.
1
email [email protected]
c The Authors Journal compilation c 2013 Biochemical Society
2
V. A. Zammit
of pre-existing TAGs. In this instance, no net genuinely new
synthesis of glyceride occurs. Secondly through the de novo
incorporation of glycerol 3-phosphate into the glyceride entity
by sequential esterification of its hydroxy groups, followed by
the formation of DAG, and its final esterification to TAG. This
involves genuine de novo glyceride synthesis. Hepatocytes have
the capacity to do both. Together with enterocytes, they are
specialized for the secretion of TRLs, VLDLs [very-LDLs (lowdensity lipoproteins)] and chylomicrons respectively. However,
in enterocytes, most TAG formation represents a reassembly of
the TAG molecule from the products of dietary TAG digestion
by pancreatic lipase (i.e. 2-MAG and fatty acids). In the liver,
TAG synthesis is an integral part of the mechanisms through
which the liver orchestrates lipid metabolism for the whole
body, whereby fatty acids mobilized from adipose tissue, or
released into the circulation during lipoprotein lipase action
on TRLs, are re-packaged into VLDLs and recirculated to the
periphery.
However, the liver is also the tissue that maintains glucose
homoeostasis, with glucose uptake and release modulated continually to maintain the plasma glucose concentration within relatively narrow limits. Therefore the liver would be expected to have
the function of integrating glycaemia with triglyceridaemia, and
to have mechanisms that link the availability of glucose
and other saccharides (e.g. fructose) with its output of VLDL–
TAG, which largely determines triglyceridaemia in obesity and
Type 2 diabetes [18–20]. It is of note that high-carbohydrate
diets in humans and animal models are associated with
hypertriglyceridaemia, adipose weight gain and their associated
metabolic pathologies [21–23]. This ability of the liver to link
carbohydrate with TAG metabolism may account for the marked
lack of success of the ‘low-fat’ dietary advice to diminish
the burden of dyslipidaemia and associated cardiometabolic
disease in industrialized populations over the last several
decades.
SECRETION OF TRLs
In hepatocytes and enterocytes, the secretion of TRLs occurs
through the assembly of particles containing a hydrophobic
core of TAG and cholesteryl esters, within a hydrophilic coat
of phospholipids, cholesterol and associated apoproteins. ApoB
(apolipoprotein B) is the molecule that keeps the particle together
by virtue of its repeated highly hydrophobic sequences. The
mechanism of this assembly is similar in the two cell types,
although that in hepatocytes has been much more extensively
studied [24,25]. At the outset it needs to be emphasized
that, although terms including ‘cytosolic’ and ‘luminal’ are
used routinely, the hydrophobic nature of TAG necessitates
that its synthesis and intracellular compartmentation are highly
membrane-associated/dependent, although TAG metabolism still
appears to distinguish on which aspect of the membrane it occurs
or to which compartment the resulting TAG is destined for.
Central to the assembly of nascent pre-VLDL particles is the
co-translational insertion of the apoB molecule through the ERM
(ER membrane). During this process, apoB abstracts TAG from
within an inter-leaflet location of the ERM where it is synthesized
into the lumen of the ER. There it surrounds this relatively small
quantity of TAG to form lipid-poor nascent LDL-type particles
which can be observed in the ER lumen [24]. This process requires
the involvement of the MTP (microsomal transfer protein). It
is the first step in the formation of small VLDL particles (VLDL2)
[24]. However, this accounts for only approximately 30 % of the
TAG synthesized on the ERM. Most of the TAG is enveloped
c The Authors Journal compilation c 2013 Biochemical Society
by the cytosolic leaflet of the ERM to form ‘cytosolic’ lipid
droplets which are bounded by a single phospholipid layer, the
proteome of which consists, although not exclusively, of many
ER membrane proteins, in addition to others that are acquired
independently and reversibly [26–28].
There is a third morphologically and compartmentally distinct
pool of TAG within hepatocytes that resides within the lumen of
the smooth ER. Although having other proteins associated with
them, these lipid droplets are devoid of apoB [29,30]. It is an
important pool of TAG because it serves to enlarge or ‘mature’
the nascent lipoprotein particles into large VLDL, as the two meet
within the proximal Golgi to undergo the second lipidation step;
this step also appears to be MTP-dependent [31]. The extent to
which this second-step lipidation occurs is presumed to determine
the size distribution of the VLDL secreted (larger VLDL1
compared with smaller VLDL2). Larger VLDLs (which are
more prevalent in diabetes [18,19]) determine the atherogenicity
of the LDL that is formed after their TAG is hydrolysed
by lipoprotein lipase in peripheral tissues and undergoes lipid
exchange with HDL (high-density lipoprotein) in the circulation
[19,32]. Therefore the ability to effect the second-step lipidation of
nascent VLDLs through the transfer of TAG from ER luminal LDs
(lipid droplets) to nascent lipoprotein particles is a process with
important (patho)physiological consequences, and which may be
amenable to pharmacological manipulation if we can understand
the mechanism(s) through which this apoB-free pool of TAG
originates and is utilized for VLDL enlargement and maturation.
The origin of this intra-ER luminal pool of TAG is thought to
be the TAG within the cytosolic LDs, but only after hydrolysis
followed by resynthesis [33]. Communication between TAG in
the cytosolic lipid droplets and the ER luminal lipid droplets is,
therefore, an important process which accounts for approximately
70 % of overall TAG secretion by the liver. However, hydrolysis
re-esterification occurs at a much higher rate than the rate of
secretion (approximately 20-fold faster) [34]. The observations
that TAG hydrolysis does not proceed all the way to glycerol and
fatty acids, but to partial glycerides (DAG and MAG) [34–36],
gave rise to the concept [37,38] that the communication between
the cytosolic and luminal pools of TAG is performed by DAG
which, unlike TAG, is highly permeable through the lipid bilayer
[39]. The corollary of this concept was that there would be DGAT
(DAG acyltransferase) activity expressed on both sides of the
ERM (i.e. across the barrier represented by the impermeability of
the lipid bilayer to acyl-CoA) to enable TAG synthesis to occur
on both aspects of the membrane.
Early observations showed that, indeed, DAG esterifying
activity (DGAT activity) is expressed on both aspects of the
membrane, i.e. that there are ‘overt’ and ‘latent’ DGAT activities
of DGAT [37,38]. It was subsequently shown that acyl-CoA
availability for TAG synthesis on the luminal aspect of the
membrane may be completed by the transfer of acyl moieties
as acylcarnitine esters [40,41]. Although the enzyme mechanism
for acylcarnitine formation and transfer across the ER membrane
is not as well studied as for the mitochondria, there are longstanding observations which suggest that an analogous system
to that in mitochondria exists in the ERM [42]. This model
accounted for the transfer of the glyceride entity across the
ERM, while keeping separate the TAG in the cytosolic and
ER luminal LDs [43]. The racemization of the stereoisomers
of TAG between that in the liver and that of secreted VLDLs
indicated that lipolysis–re-esterifiaction cycling occurred to DAG
and MAG [35,36], but it did not exclude the possibility that
cycling occurs also within the ER lumen, and that DAG
moves in both directions and equilibrates across the ERM
(see Figure 1).
DGAT2 integrates glycaemia and triglyceridaemia
Figure 1
3
DGAT2 and DGAT1 act in series
Pathway showing how DGAT2 acts upstream of DGAT1, and the relationship between de novo fatty acid synthesis, TAG synthesis and secretion in hepatocytes. DGAT2 acts upstream of DGAT1 and
utilizes nascent DAG and de -novo -synthesized fatty acids as substrates. TAG formed by DGAT2 provides substrate for lipases which generate partial glycerides that, together with exogenous fatty
acids, are substrates for DGAT1. DGAT2 utilises de -novo -synthesized (and desaturated) fatty acids preferentially, possibly by virtue of its co-localization with enzymes of the glycerol 3-phosphate
pathway and SCD1 (see the text). The association with FATP1 facilitates deposition of newly synthesized TAG into small cytosolic LDs. A minority of the TAG formed is partly sequestered for secretion
during the co-translational insertion of the apoB molecule through the ERM, and partly stored in cytosolic LDs, without undergoing lipolysis. Most of the newly DGAT2-synthesized TAG is hydrolysed
by lipase(s) located in the cytosolic compartment of the cell (or structures exposed to it) to give partial glycerides that are re-esterified to TAG by DGAT1, using preformed exogenously derived fatty
acids. DAG equilibrates across the ERM. DGAT1 activity expressed on the luminal aspect of the ERM uses DAG (derived from TAG lipolysis either in the cytosol or from TAG–DAG cycling within the
ER lumen) to synthesize TAG within the ER lumen. This luminal (apoB-free) TAG is used for the second-step lipidation of nascent VLDL, before the mature VLDL particle is secreted. LP, lipoprotein.
Modified from Wurie, H.R., Buckett, L. and Zammit, V.A. (2012) Diacylglycerol acyltransferase 2 acts upstream of diacylglycerol acyltransferase 1 and utilizes nascent diglycerides and de novo
synthesized fatty acids in HepG2 cells. FEBS J. 279, 3033–3047, with permission.
TWO DISTINCT DGATs
The above observations were followed by the cloning of the cDNA
of two unrelated proteins that have DGAT activity, which were
termed DGAT1 and DGAT2, in the order in which they were
described [44–46]. A third enzyme present in human enterocytes
{MGAT (MAG acyltransferase) 3 [47]} also has substantial
DGAT activity, but will not be considered further in the present
review. In mammals, DGAT1 is expressed in skeletal muscle,
skin, intestine (ileum, colon) and testis, with lower levels of
expression in liver and adipose tissue [44]. DGAT2 is similarly
widely expressed with high expression levels in hepatocytes and
adipocytes [45]. Both of the different tissue distributions, and the
fact that the two major proteins with DGAT activity have been
evolved from two separate protein families [48], indicate that these
enzymes have specialized functions even though they catalyse
the same reaction and are co-expressed, to different extents, in
individual cell types. In addition, their respective overexpression
in hepatoma-derived MzcArdle cells [49] results in markedly
different intracellular patterns of TAG droplet accumulation
within the cells, again suggesting that the properties of TAG
synthesized by DGAT1 and DGAT2 may be distinct.
The salient physiological feature of DGAT1 and DGAT2 is that
although they catalyse the same reaction, and are co-expressed in
most cells, they are non-redundant, i.e. when the expression of one
is disrupted, the other cannot substitute for it (although it has been
claimed that DGAT1 can substitute for the absence of DGAT2
[49,50], those experiments only measured fatty acid incorporation
into TAG, which can proceed without net new synthesis of TAG,
through hydrolysis–re-esterification cycling, see above). This
phenomenon is most apparent in the very different phenotypes of
the respective global knockout mice (Dgat1 − / − and Dgat2 − / − ).
Dgat1 − / − mice have a very mild phenotype and are resistant to
obesity, and the development of diet-induced obesity and insulin
resistance [51]. They have slightly lower hepatic TAG levels when
fed a low-fat diet, but, importantly (see below) when maintained
on a high-fat diet they have a significantly lowered hepatic TAG
content [52,53]. In contrast Dgat2 − / − mice do not survive for
more that 24 h after birth and are severely lipopaenic, with waterbarrier deficiencies in their skin [49]. The inability of DGAT1
(the expression of which is not increased) to rescue the Dgat2 − / −
phenotype suggested that DGAT2 has a very specialized function.
The other important feature that emerged was a key difference
between DGAT1 and DGAT2, namely the multi-functionality of
c The Authors Journal compilation c 2013 Biochemical Society
4
V. A. Zammit
DGAT1 (but not DGAT2) in being able to esterify both DAG and
MAG [54,55]. This is particularly important not only because
it allows DGAT1 to esterify both the glyceride products of
TAG hydrolysis, but also because in non-hepatic tissues, MAG
may be the major substrate for TAG synthesis (see below).
Otherwise, there are only minor differences in the substrate
specificities between the two enzymes in vitro, which in vivo
are likely to be overwhelmed by the specialization arising from
compartmentalization and protein–protein interactions. Thus
when Tung Tree epitope-tagged DGAT1 and DGAT2 were
expressed in cultured cells, they occupied very discrete nonoverlapping regions of the ER [56]. Moreover, plant DGAT2
has been found to interact with GPAT (glycerol-3-phosphate
acytransferase) [57], and in animal cells DGAT2 (but not
DGAT1) co-localizes with SCD1 (stearoyl-CoA desaturase 1)
[58] and a protein with acyl-CoA synthase activity [FATP1
(fatty acid transport protein 1)] [59]. These observations suggest
that DGAT2 exists in a multiprotein micro-environment within
which substrates and products of fatty acid acylation, desaturation
and esterification are channelled sequentially among the closely
interacting constituent enzymes.
INTRACELLULAR LOCATION AND MEMBRANE TOPOLOGY OF
DGAT1 AND DGAT2 CATALYTIC SITES
The discovery that DGAT activity is expressed on both aspects
of the ERM [37] suggested that one of the newly described
DGAT proteins would have its active site exposed on the cytosolic
aspect of the ERM, and the other on the luminal aspect. However,
this turned out to be an oversimplification of the situation. The
exclusively overt expression of the DGAT2 active site is now well
established through the use of several different approaches. In
particular, epitope tagging of heterologously expressed protein
from both plant and mammalian sources demonstrated that
DGAT2 is a polytopic protein with two TM (transmembrane)
domains separated by a short loop, with both the N-terminal and
the larger and catalytic site-bearing C-terminal domain exposed
on the cytosolic aspect of the ERM [60]. DGAT2 is present not
only on the ERM, but also on LDs and mitochondria-associated
ER [61]. Its function on LDs appears to be to enable their
expansion through its interaction with the ER protein FATP1 [59].
Therefore all DGAT2 activity is ‘overt’, as it can be accessed by
cytosolic acyl-CoA esters, although its location is heterogeneous,
having been described on the surface of cytosolic LDs and
mitochondria. This has been ascertained through inhibitor
and transgenic mouse studies in which it was found that activity
corresponding to that of DGAT2 is expressed entirely overtly (i.e.
is fully accessible to cytosolic acyl-CoA substrates) [62,63].
The topology of DGAT1 is more complex. Whereas epitopetagging and partial proteolysis approaches concluded that the
DGAT1 catalytic site is expressed exclusively on the latent aspect
of the ERM [64], several other lines of evidence suggest that
the enzyme has a dual topology [62,63], with its active site
expressed on both the overt and latent aspects of the membrane.
Thus microsomes isolated from Dgat1 − / − mice have no latent
DGAT activity, which suggests that the latent activity is due
entirely to DGAT1 [62,63]. However, in these microsomes there
is also a diminution of a substantial fraction of the overt DGAT
activity, indicating that DGAT1 activity is also expressed on the
overt aspect of the ERM [63]. Moreover, experiments conducted
on HepG2 cells with two specific inhibitors developed against
human DGAT1, and belonging to different classes of chemicals,
showed that overt and latent DGAT1 activities are differentially
sensitive to these two classes of pharmacological agents [63].
c The Authors Journal compilation c 2013 Biochemical Society
These observations indicated that DGAT1 has dual membrane
topology in the ERM of HepG2 cells, with DGAT1 molecules
being orientated in one of two possible conformations within the
membrane, one with its active site available to cytosolic substrates,
and the other to luminal substrate pools (specifically acyl-CoA).
This agreed with the data in [65] where it was observed that
there was an increase in both overt and latent DGAT activity
when DGAT1 was expressed adenovirally in the liver of mice.
Dual topology of DGAT1 in the ERM has also been found in the
enterocyte-derived cell line Caco-2 (H.R.Wurie, L.Buckett and
V.A. Zammit, unpublished work).
The molecular mechanism through which DGAT1 achieves
its dual topology is still being elucidated. However, it may be
pertinent that DGAT1 exists as a tetramer [66], because the
existence of oligomeric membrane proteins with dual topology
is now well established [67–70]. Dual topology of oligomeric
membrane proteins is associated with weak topographical motifs
flanking the boundaries of their TM domains. This allows each
individual protomer to adopt an orientation independently of
the other protomers within the overall oligomeric structure,
resulting in their active sites being expressed on both aspects
of the membrane [69]. The ability of at least one enzyme that,
in yeast, is able to vary the relative proportion of number of
protomers facing one or the other aspect of the membrane
depending on the nutrient status of the organism suggests that
the balance between the protomers adopting each of the two
topologies is regulated by co- or post-translational mechanisms
[67,71]. DGAT1 exists as a dimer/tetramer in vivo, and the
partitioning of individual protomers between the two possible
membrane orientations may be the mechanism through which
the dual topology of its active site in the ERM is achieved.
Interestingly, changes in the relative magnitudes of overt and
latent DGAT activities in response to physiological, dietary
and pharmacological (fibrate, statin) treatments have been
described [72], suggesting that the relative proportions of the
number of protomers in each orientation is under metabolic
control. In particular, there is a 5-fold increase in the overt/latent
ratio of DGAT activity in rat liver microsomal membranes
following fibrate treatment of rats [72]. Considering that DGAT1
only constitutes a fraction of overt DGAT activity in the control
state, this increase in overt DGAT activity may be the result of
an even greater re-configuration of DGAT1 orientation across
the ERM after fibrate treatment. Such redistribution of DGAT1
topology across the ERM to favour cytosolic expression of its
active site may contribute towards the beneficial hypolipidaemic
effect of fibrates [72] especially as lumen-facing DGAT1 may
determine the size of VLDL particles (see below).
Recently, a mechanism has been described for the physiological
control/modulation of the relative proportions of protomers of
dual-topology proteins that face either aspect of eukaryotic
membranes [73,74]. This model proposes that physiologically
induced changes in membrane lipid composition may affect the
topology of individual conformers both during co-translational
insertion and by enabling re-orientation of the protein within
the membrane even after its insertion. This would provide the
basis for a mechanism through which the topology of such
proteins can be affected physiologically by nutritionally and
hormonally mediated changes in membrane lipid composition.
Such a mechanism may be harnessed pharmacologically as a
strategy of altering the fraction of TAG that is secreted by the
liver and the TAG content (size distribution) of VLDL.
The ability of DGAT1 to vary its topology in the ERM may
also enable it to adapt its orientation to the physiology of the
different cell types in which it occurs. Thus although having a
topology that results in partial latency of its active site within the
DGAT2 integrates glycaemia and triglyceridaemia
ERM is important for the function of cell types that secrete TRLs
(hepatocytes and enterocytes), it would be inappropriate for TAG
formation in non TRL-secreting cell types (e.g. myocytes and
adipocytes) in which TAG-containing LDs are exclusively located
in the cytosolic compartment. Therefore, for DGAT1, possessing
membrane-insertion motifs that enable it to be expressed with
either of the two possible topologies in the same membrane would
ensure that DGAT1 activity is expressed appropriately on either
aspect of the ERM in different cell types, depending on their acute
or long-term physiological requirements.
5
the carbohydrate content of the diet is increased substantially
[83,84]. Therefore the liver superficially appears to depend on a
‘catalytic’ amount of de-novo-synthesized fatty acid to enable
hepatic TAG synthesis and secretion to occur [35,36,85,86].
Whereas the link between carbohydrate intake and hepatic fatty
acid synthesis could be rationalized by the insulinaemic effects
of carbohydrate-rich diets on lipogenesis, the identity of the link
to TAG synthesis and secretion remained unresolved. The answer
appears to centre around the specialized and rate-limiting function
of DGAT2 [63].
PRE-REQUISITES FOR HEPATIC TAG SYNTHESIS AND SECRETION
The importance of the dual topology of DGAT1 within the ERM of
hepatocytes is that it enables it to maintain distinct TAG pools on
both sides of the membrane in the presence of lipase action. The
liver contains several lipases that hydrolyse TAG at high rates
on both the cytosolic and ER luminal LDs [75]. Therefore the
expression of DGAT1 activity on both aspects of the membrane
is important to maintain both TAG pools, especially as DGAT1
is the only one of the two DGATs that can utilize both DAG and
MAG (and long-chain fatty acyl-CoAs) as substrates [55] (see
below). These properties enable DGAT1 to rescue both of these
partial glycerides from complete hydrolysis. The question arises
however, as to why DGAT2 cannot perform this function on the
cytosolic aspect of the ERM, i.e. why DGAT1 activity is required
also on the cytosolic aspect of the ERM. The answer appears to
lie in the specialized role that DGAT2 plays in the pathway of
TAG synthesis.
There are two essential features of TAG synthesis and secretion
in the liver that have been recognized for a considerable amount
of time, but that appear to be paradoxical. These are (i) their
dependence on de novo lipogenesis of fatty acids [76–78], and
(ii) their dependence on the endogenous desaturation of the
products of de novo fatty acid synthesis [79]. The ability of highcarbohydrate diets to activate de novo fatty acid synthesis through
the activation of the relevant transcription factors [e.g. SREBP1c (sterol-regulatory-element-binding protein 1c) and ChREBP
(carbohydrate-responsive-element-binding protein)] is well
established [80]. This link between de novo fatty acid synthesis
and TAG synthesis/secretion was experimentally demonstrated
in vivo in mice bearing a hepatocyte-specific deletion of the
insulin receptor (LIRKO mice) [81]. These mice had increased
apoB secretion, but lower VLDL–TAG secretion. These changes
were accompanied by a down-regulation of FAS (fatty acid
synthase) and SCD1, but a marked increase in DGAT1 expression
(DGAT2 was not measured) [81]. The observation of the downregulation of SCD1 induced by the absence of insulin signalling
is significant, as hepatic SCD1 deficiency results in a markedly
impaired ability of the liver to synthesize and secrete TAG [79,82].
Therefore there appears to be a requirement by the liver for fatty
acids synthesized and desaturated in situ in order for it to be able
to synthesize and secrete TAG. The requirement for desaturation
of de-novo-synthesized fatty acids for hepatic lipogenesis and
TAG secretion to occur is likely to be due to the effects that the
unsaturated products of SCD1 have on the expression of lipogenic
enzymes, through their effects on transcription factors (SREBP1c and ChREBP) involved in carbohydrate-induced activation of
de novo fatty acid synthesis [22,79].
However, the dependence of TAG secretion on de novo
lipogenesis is not what would be expected from the very minor
contribution that de-novo-synthesized fatty acids make to secreted TAG. They constitute a small fraction (<5 %) of those secreted
within VLDL–TAG (both in animal models and humans) unless
DGAT2 AND DGAT1 ACT IN SERIES AND NOT IN PARALLEL
As mentioned above, DGAT2 activity is entirely ‘overt’, i.e. it is
accessible to cytosolic substrates. This agrees with its membrane
topology in the ERM [60] and its detection on the cytosolic
LDs and mitochondria-associated microsomes [61]. It accounts
for approximately 30 % of overt DGAT activity [63], and less
than 15 % of overall cellular DGAT activity [63,87]. And yet,
inhibition of such a small fraction of cellular DGAT activity has
an almost total inhibitory effect on the de novo synthesis of TAG
(defined as the de novo incorporation of the glyceroyl moiety into
TAG) [63]. By contrast, inhibition of DGAT2 does not affect the
incorporation of exogenously supplied oleate into TAG (which
can occur independently of any net synthesis of glyceride due
to hydrolysis–re-esterification cycling between TAG and partial
glycerides) [63].
Specific inhibition of DGAT1 results in a totally different
pattern of loss of incorporation of glycerol and preformed fatty
acids into TAG. There is a strictly proportional loss of labelling
of both the glyceride and acyl parts of the TAG molecule
[63]. Therefore there is an extreme specialization of the two
DGATs. DGAT2 is specialized for the incorporation of glycerol
3-phosphate, but not of exogenous preformed fatty acid into TAG.
These observations were corroborated by the observations of
[87] which showed, using stable isotopes, that DGAT2 inhibition
results in the loss of incorporation of glycerol, but not of oleate,
into TAG, both in hepatocytes and in vivo. By contrast, DGAT1
appeared to be specialized for incorporation of oleate, but not
glycerol, into TAG [87].
These observations raise questions as to (i) the identity of
the fatty acid substrates for DGAT2 (since it does not use
preformed exogenously supplied fatty acid), and (ii) how the
glyceroyl moiety newly incorporated into DAG is brought
together with long-chain fatty acids into the same TAG molecule.
The possibility emerged that if DGAT2 does not utilize preformed
exogenous fatty acids it may be specialized for the utilization
of de-novo-synthesized fatty acids that arise endogenously. This
was found to be the case [63]. Down-regulation of the relatively
low DGAT2 activity [by a selective inhibitor or by siRNA
(small interfering RNA)-mediated knockdown] in HepG2 cells
had the same disproportionately large (near total) inhibition
of the incorporation of acetate-derived fatty acids into TAG.
Therefore inhibition of DGAT2 affected glycerol and acetate
incorporation into TAG in parallel, indicating that DGAT2 utilizes
nascent diglycerides (newly formed and containing de-novosynthesized fatty acids) and de-novo-synthesized long-chain acylCoA as substrates [63]. (‘Nascent diglycerides’ refers to those
DAG molecules that arise from de novo incorporation of the
glyceroyl moiety, i.e. not those arising from the hydrolysis of
TAG.) By contrast, loss of DGAT1 inhibited glycerol and oleate
incorporation equally and stoichiometrically (at a 1:3 ratio),
suggesting that when DGAT1 activity is inhibited, the loss of
c The Authors Journal compilation c 2013 Biochemical Society
6
V. A. Zammit
any acyl chain from a TAG molecule, due to lipolysis, results in
the loss of the entire glyceride entity from the TAG fraction [63].
The parallel effects on (i) glycerol and acetate incorporation by
DGAT2 inhibition, and (ii) on glycerol and oleate incorporation
by DGAT1 inhibition indicate that the product of DGAT2
generates (after lipolysis) the DAG substrate for DGAT1, i.e. that
DGAT2 acts upstream of DGAT1 (Figure 1) [63].
IMPLICATIONS OF THE SEQUENTIAL ACTION OF DGAT2 AND DGAT1
There are very important implications of the observations
described above, namely (i) that DGAT1 and DGAT2 act in series
rather than in parallel in hepatocytes; (ii) that DGAT2 is primarily
responsible for the initial synthesis of TAG, and thus acts upstream
of DGAT1; and (iii) that DGAT1 functions primarily to retain
acyl groups and the glyceride moiety within the TAG fraction,
by catalysing the re-esterification of DAG and MAG formed
after lipase-mediated hydrolysis of TAG. The rapid TAG–partial
glyceride cycling that occurs in hepatocytes [33,34], a process
during which 70 % of TAG becomes racemized [35], results in
the TAG secreted in VLDL having a different stereo-isomeric
composition of acyl chains than of that in cytosolic LDs [35]. The
action of DGAT2 upstream of DGAT1, indicates that it has access
to pools of de-novo-synthesized diglyceride and fatty acids that are
not available to DGAT1. This explains the apparently paradoxical
and long-standing observations that VLDL–TAG formation is
strictly linked to the rate of de novo fatty acid synthesis in the
liver [76–78]. Such a rate-limiting role of DGAT2 would not
have been discerned from the modest contribution that de-novosynthesized fatty acids make towards secreted TAG under normal
carbohydrate diets [83]. Thus although in both humans and animal
models fed a normal carbohydrate diet the absolute contribution
of de-novo-synthesized fatty acids to VLDL–TAG is very low
(∼ 5 %) [83,88], inhibition of fatty acid synthesis results in an
almost complete cessation of TAG synthesis and VLDL–TAG
secretion [76–78]. Similarly, inhibition of SCD1 or disruption of
its gene have an almost complete inhibitory effect on hepatic TAG
synthesis in vivo [89] which cannot be rescued by dietary triolein
feeding [89], suggesting that the DGAT enzyme that utilizes denovo-synthesized fatty acids exerts a rate-limiting effect on overall
TAG synthesis and secretion. This is in agreement with the ratelimiting role predicted from the upstream function of DGAT2
suggested in [63] (Figure 1).
However, a minor, but substantial, proportion of TAG
synthesized by DGAT2 does not appear to undergo hydrolysis–reesterification cycling [63]. This suggests that this portion of newly
synthesized TAG is sequestered into a pool(s) that bypasses the
hydrolysis–re-esterification cycling and is incorporated directly
(i.e. used intact) for incorporation into LDs or for secretion. It
is well-established that a sizeable minority of TAG is secreted
without prior hydrolysis–re-esterification [33,34]. In this respect,
in [35,36] it was shown that in vivo only approximately 70 % of
newly synthesized TAG is racemized before secretion by the liver.
Similarly, studies on LD formation have suggested that different
LDs may have differential access to lipogenic enzymes [90], and
that DGAT2, but not DGAT1, is specifically associated with LDs
[90] and may form part of a bridge to the ERM in association
with FATP1 [59]. In this context, it is of interest that others have
observed that small ‘new’ and large ‘old’ LDs in the cytosolic
compartment of McArdle cells have different dynamics, and that
whereas there is transfer of TAG from small nascent droplets to
larger LDs, this does not happen in the reverse direction [30].
A role for DGAT1 in ‘rescuing’ partial glycerides (DAG and
MAG) after their formation through TAG hydrolysis, means that
c The Authors Journal compilation c 2013 Biochemical Society
this enzyme is very important in the retention of glycerides within
the TAG fraction of the liver (and other tissues, see below). This
may explain why overexpression of DGAT1 in mouse liver in vivo
increases hepatic TAG content [91]. DGAT1 has been shown to
have considerable MGAT activity in its own right [55], and may,
therefore, re-esterify both DAG and MAG to TAG. Therefore
DGAT1 is expected to play an essential role in the accumulation
of TAG either for retention in the cell or as a source of TAG
for secretion. Without its action, DAG and/or MAG generated
by lipases could be diverted to phospholipid synthesis (from
DAG) or total hydrolysis (from MAG) to glycerol and fatty acids.
DGAT1 activity may need to be much higher compared with that
of DGAT2 in hepatocytes, as its catalytic activity would need
to accommodate the very rapid cycling flux between TAG and
partial glycerides [33,34]. Because of the extent of this cycling,
the de-novo-synthesized fatty acids incorporated into TAG during
initial (DGAT2-mediated) synthesis may be rapidly replaced by
preformed fatty acids in the re-esterification reactions catalysed
by DGAT1, hence explaining the low content of de-novosynthesized acyl chains in secreted TAG, unless the rate of de
novo fatty acid synthesis is increased substantially, e.g. under
conditions of carbohydrate-diet-induced lipogenesis [83].
The model in Figure 1 may also explain the observation,
obtained with liver-specific DGAT1-knockout mice, that DGAT1
is required for the development of steatosis due to excess exogenous (preformed) fatty acid supply to the liver, whereas steatosis
associated with increased hepatic de novo fatty acid synthesis is
not DGAT1-dependent [92]. These data are fully consistent with
the conclusion that DGAT2 utilizes substrates that are, or are
composed of, de-novo-synthesized (‘new’) fatty acids, whereas
DGAT1 is primarily involved in esterifying ‘old’ preformed fatty
acids to DAG. More generally, these considerations also explain
the extreme phenotype and lethality (soon after birth) of global
Dgat2 gene disruption in mice [49], as this would prevent primary
TAG synthesis (de novo incorporation of glycerol into TAG). By
contrast, global disruption of the Dgat1 gene only leads to a
mild phenotype characterized by lower TAG contents of tissues
(including the liver) and mild hypotriglyceridaemia [52] as would
be expected from a lower rate of ‘rescue’ of DAG and MAG after
lipase-mediated hydrolysis of TAG.
The rate-limiting role of DGAT2 for de novo TAG synthesis
may explain why niacin is a good hypolipidaemic agent [32].
Niacin inhibits DGAT2 non-competitively, but not DGAT1
[93]. As expected from the relatively low activity of DGAT2
compared with DGAT1 [62], niacin only affects a relatively
small percentage of overall DGAT activity [62,63]. It is now
evident that by inhibiting DGAT2, niacin would prevent hepatic
steatosis associated with endogenous synthesis of fatty acids.
Interestingly, the efficacy of niacin as an agent against fatty liver is
dependent on DGAT2 polymorphisms of the individuals studied
[94], confirming the central role of the enzyme in determining the
rate of de novo TAG synthesis and, in turn, of the absolute rates
of net de novo TAG synthesis by the liver.
INSULIN EFFECTS ON TAG SYNTHESIS AND SECRETION
Hyperinsulinaemia in vivo is associated with hypertriglyceridaemia [95]. But insulin promotes the degradation of apoB [96],
an effect that would act contrary to the lipogenic effect of the
hormone [80,97]. In the present context, insulin is expected
to increase the rate of DGAT2-mediated TAG synthesis by
increasing the rate of de novo lipogenesis through its induction of
SREBP-1c [80]. Importantly, this insulin-stimulated lipogenesis
is preserved in insulin-resistant states, owing to the phenomenon
DGAT2 integrates glycaemia and triglyceridaemia
of selective insulin resistance [81,97]. Moreover, because insulinresistant states are accompanied by hyperinsulinaemia, hepatic
lipogenesis is expected to be stimulated to an even greater extent
than normal, rather than being inhibited under these conditions.
Thus in mice in which the insulin receptor was specifically
knocked out in hepatocytes (LIRKO mice) to produce ‘pure’
insulin resistance, there was a marked decrease in SCD1 and FAS
expression and inhibition of VLDL–TAG secretion in spite of an
increase in DGAT1 (DGAT2 was not measured) [81]. Moreover,
enhanced insulin signalling [mediated by interference with the
action of PTEN (phosphatase and tensin homologue deleted on
chromosome 10)] increased TAG synthesis and the accumulation
of cellular TAG, thus overriding the effect of insulin on apoB
degradation, and resulting in increased VLDL–TAG secretion.
These observations indicated that the level of both apoB and TAG
secretion are determined by the effects of insulin on lipogenic
rate and TAG availability rather than by the stability of apoB
[98]. This corroborates previous data [88,99] showing that in vivo
and in perfused livers (but not in isolated primary hepatocytes)
insulin only inhibits TAG secretion by the liver in the fasted state
(in rats and humans respectively), and that in the fed insulinreplete state, when lipogenesis is higher, insulin stimulates TAG
secretion [100,101]. Therefore the effect of insulin on hepatic
TAG secretion is dependent on the prior insulinaemic state of
the liver [99]. These observations are reproducible using rat
isolated perfused livers [102], but not in isolated hepatocytes
unless they are cultured with insulin for a prolonged period
[103]. They provide a rationale for the conclusions reached
previously, through work on humans fed a high-carbohydrate diet,
that hyperinsuinaemic hypertriglyceridaemia is not due to failure
of insulin to inhibit TAG secretion [23]. We have previously
highlighted the possibility that, due to the stimulation of TAG
secretion that it is likely to cause, the possible deleterious effects of
premature treatment of newly diagnosed Type 2 diabetic patients
with insulin may outweigh any benefits of the strict glycaemic
control achieved [100,101], a prospect that has been reiterated
more recently [81].
This may now be explained by the link that DGAT2 provides
between de novo fatty acid synthesis (which does not become
resistant to the action of insulin in the pre-diabetic state [98])
and TAG synthesis. Thus, in the (pre)diabetic state, DGAT2mediated de novo TAG synthesis proceeds undiminished (if not
actually increased), and DGAT1 will have an increased level of
substrate available (from circulating fatty acids released from
adipose tissue) for re-esterification of partial glycerides. As
expected, this combination of increased lipogenesis and increased
availability of circulating fatty acids results in the steatosis and
hypertriglyceridaemia that accompany this condition, in spite of
the relatively increased partitioning of fatty acids towards βoxidation [104,105]. The effects of the increased glucose and
cholesterol output by the liver {owing to insulin resistance
of pathways regulated by Akt, but not through mTORC1
(mammalian target of rapamycin complex 1) [81,98,106]},
coupled to the enhanced lipogenesis and associated synthesis
of TAG, result in combined hyperglycaemia, hyperinsulinaemia
and hypertriglyceridaemia experienced in insulin-resistant states.
Therefore DGAT2 emerges as the link between these three
important markers of the metabolic syndrome.
7
of TAG from the MAG and fatty acid produced by pancreatic
lipase in the lumen of the gut [25]. Therefore there is likely to
be only a small amount of net TAG synthesis (from glycerol
3-phosphate and de-novo-synthesized fatty acids). Accordingly,
MGAT activity is high in enterocytes. DGAT1 may contribute
towards this activity owing to its ability to use MAG as a substrate
in addition to its role in catalysing the final reaction in TAG
synthesis [55]. In humans, MGAT3 may contribute towards reesterification in enterocytes owing to its DGAT [107] activity
[47,108].
The dual distribution of DGAT1 on the overt and latent aspects
of the ER membrane of Caco2 cells has been experimentally
demonstrated (H.R. Wurie, L. Buckett and V.A. Zammit,
unpublished work) indicating that the role of DGAT1 in reesterifying partial glycerides within the ER lumen also exists
in this cell type. Indeed, although it was at first suggested
that DGAT1 is not necessary for dietary TAG uptake [109], it
is now known that intestinal TAG absorption in mice lacking
DGAT1 specifically in the intestine is delayed, resulting in much
lower post-prandial excursions of TAG [110]. This suggests that,
although as in the liver the direct transfer of intact TAG during
co-translational translocation of apoB across the ERM occurs,
in the absence of DGAT1 the high rates of secretion of TAG
enabled by resynthesis of TAG within the ER lumen is not
possible. Specific expression of DGAT1 solely in enterocytes of
Dgat1 − / − mice reverses the resistance of the animals to high-fatdiet-induced obesity [111], indicating that the lowering of the
post-prandial excursions of TAG may be sufficient to relieve
the lipotoxicity of tissues such as skeletal muscle. Treatment
of mice with a specific DGAT1 inhibitor totally abolishes postprandial TAG excursions and the appearance of labelled acyl
groups from dietary triolein in chylomicrons [110]. Interestingly,
although they had similar effects on chylomicron TAG secretion,
respective inhibition of MTP and of DGAT1 had opposite effects
on TAG accumulation in enterocytes [110]. Thus MTP inhibition
resulted in the accumulation of cellular TAG, whereas DGAT1 inhibition resulted in a decrease in cellular TAG content [110]. This
is likely to have arisen from the fact that inhibition of MTP results
in the backing-up of TAG within the cell when the MTP-dependent
mechanisms for TAG incorporation into chylomicrons are lost,
especially since TAG continues to be absorbed normally [110]. In
contrast, when enterocyte DGAT1 is inhibited, the (re)synthesis
of TAG from partial glycerides is not possible, resulting in total
hydrolysis of the partial glycerides to fatty acids and glycerol.
In enterocytes, the predominant use of sn-2-monoglyceride for
intracellular TAG (re)synthesis suggests that DGAT2 would be
mostly bypassed, as only DGAT1 (and other enzymes with MGAT
activity) would be required; DGAT2 expression, like de novo fatty
acid synthesis, is low in enterocytes [109] (see Figure 2). However,
TAG secretion by the intestine is positively related to the rate of de
novo fatty acid synthesis, although it is not dependent on it [107].
This indicates that while the rate of de novo fatty acid synthesis
in enterocytes is likely to be significantly lower than that in the
liver, DGAT2 may still be able to play a role in linking the two
processes under conditions in which lipogenesis from glucose
(e.g. in diabetes) is elevated; it may be involved in promoting
the increased rate of chylomicron TAG secretion observed in the
fructose-fed hamster, an animal model of diet-induced insulin
resistance [107,112].
THE ROLES OF DGAT2 AND DGAT1 IN NON-HEPATIC TISSUES
Enterocytes
Myocytes and adipocytes
Like hepatocytes, enterocytes secrete TRLs, but the process of
TAG synthesis in this cell type is largely one of re-synthesis
Cardiac and skeletal muscle fibres can accumulate substantial
amounts of TAG [2–4,113]. In the heart, this is associated
c The Authors Journal compilation c 2013 Biochemical Society
8
Figure 2
V. A. Zammit
Comparison of the possible routes to TAG synthesis in hepatocytes, enterocytes and myocytes
In tissues in which, unlike the liver, de novo synthesis of fatty acids from carbohydrate sources is not a major process (intestine and muscle), DGAT2 may be bypassed through the formation of partial
glycerides by lipase-mediated hydrolysis of TAG (e.g. by pancreatic lipase and LPL). These glycerides can then be used by DGAT1 to resynthesize TAG within the respective cell types (enterocytes
and myocytes). When de novo fatty acid synthesis from glucose becomes more important (e.g. in hyperglycaemiac and hyperinsulinaemic states in muscle) it would provide substrates for DGAT2.
In adipocytes, the involvement of DGAT2 may be more significant, as lipogenesis from glucose may be more prevalent. FA, fatty acid.
with cardiomyopathy [113], and in skeletal muscle with insulin
resistance when it occurs under conditions of dietary excess
[7–9,114]. However, muscles of athletes (which can oxidize
fatty acids at high rates) also have a high intra-myocytic TAG
content [115,116], a phenomenon termed the ‘athletes paradox’.
Myocytes receive a substantial part of their fatty acids as the
products of LPL (lipoprotein lipase) action on VLDL– and
chylomicron–TAG. Because LPL activity may result in the
incomplete lipolysis to MAG [117] it is possible that, as in
enterocytes, when metabolizing the products of LPL action,
muscle DGAT2 is bypassed in the formation (resynthesis) of
TAG inside the cells (see Figure 2). No MAG is detected in the
circulation when chylomicrons are hydrolysed by a subcutaneous
depot of adipose tissue in vivo [118], suggesting that MAG
formed by LPL may be very efficiently taken up by tissues. Once
inside the cells, it is either further hydrolysed by monoglyceride
lipase or used directly to synthesize DAG and TAG, in a manner
analogous to the use of 2-monoglyceride in enterocytes [25]. In
this instance, the MGAT-like activity of DGAT1 may be important
in the ‘clearance’ of MAG, DAG and fatty acid (acyl-CoA) from
the cytosol of myocytes under dyslipidaemic conditions. DGAT1
would undertake the resynthesis of TAG intracellularly from these
products of LPL action. In the process, this would ensure that the
concentrations of acyl-CoA and other intermediates (DAG and
ceramide) that may induce insulin resistance [15,119] are kept
low. This may explain why overexpression of DGAT1 in type1 muscle fibres renders them more insulin sensitive [120], and
would be in agreement with the observation in some studies
that exercise acutely increases DGAT1 expression in muscle
[8]. However, the correlation of DAG with insulin resistance
in muscle is still uncertain. Thus whereas chronic [121,122]
or acute [8] exercise lowers cellular ceramide, athletes tend to
have a higher intra-myocytic DAG content than sedentary insulinresistant individuals [122].
c The Authors Journal compilation c 2013 Biochemical Society
Figure 3 Proposed roles of the specialized functions of DGAT1 and DGAT2
in the alleviation and promotion respectively, of insulin resistance in skeletal
and cardiac muscle
Both insulin-resistant (e.g. in Type 2 diabetic) and highly insulin-sensitive (e.g. in athletes)
muscle cells contain high levels of TAG. Besides differences in the rate at which these muscles
can oxidize fatty acids, the DGAT enzyme responsible for the synthesis of TAG may be involved.
(A) In exercised muscle, increased DGAT1 activity clears acyl-CoA and DAG to form TAG
from preformed fatty acids. (B) In obese/diabetic individuals, increased lipogenesis induced by
hyperglycaemia and hyperinsulinaemia provides substrates for DGAT2. The TAG thus synthesized
is a source of acyl-CoA and DAG generated by intracellular lipases. ATGL, adipose TAG lipase;
FA, fatty acid; HSL, hormone-sensitive lipase; IS, insulin sensitivity; NEFA, non-esterified fatty
acid.
Interestingly, overexpression of DGAT2 in type-2 muscle fibres
has the opposite effect than that of the expression of DGAT1 in
type-1 fibres; it makes muscle more insulin resistant [113], even
though the accumulation of TAG in these fibres occurs to the same
DGAT2 integrates glycaemia and triglyceridaemia
Figure 4
9
The role of DGAT2 as the link between glycaemia and triglyceridaemia
Hyperglycaemia and attendant hyperinsulinaemia stimulate de novo fatty acid synthesis through the activation of transcription factors including SREBP-1c and ChREBP. DGAT2 utilizes the products
of de novo fatty acid synthesis and of de novo DAG formation for the synthesis of TAG which is used (after hydrolysis and re-esterification by DGAT1) for TAG secretion within VLDLs. Therefore
DGAT2 links hyperglycaemia (and attendant hyperinsulinaemia) and hypertriglyceridaemia.
extent as in type-1 fibres overexpressing DGAT1. Although the
targeting of a different fibre type in these experiments complicates
the interpretation of these data, they raise the interesting prospect
that DGAT2 activity is linked to lipogenesis and the net new
synthesis of glycerides also in muscle cells. TAG formed
by DGAT2 would be metabolically distinct from that formed by
DGAT1. Rather than clearing acyl-CoA, de novo synthesis of
new TAG by DGAT2 would provide substrate for lipases that
generate the intermediates (acyl-CoA and DAG) that promote
insulin resistance (see Figure 3). It is well-established that the
intramyocellular content of TAG (IMTG) is high in obese/diabetic
individuals, in whom muscle is insulin resistant, but also in trained
athletes, in whom it is very insulin sensitive. In view of the
specialized functions of DGAT2 and DGAT1, it is possible that
this ‘athletes paradox’ [115,116] may be related to the respective
properties of the distinct pools of TAG synthesized by DGAT1
(with its ability to remove lipid intermediates) or DGAT2 (linked
to lipogenesis-driven synthesis of new TAG). Although muscle
is not normally thought to be a lipogenic tissue, even under
basal conditions it uses a considerable fraction of glucose uptake
(∼ 5 %) to synthesize fatty acids, and this proportion increases
markedly when myocytes are cultured under hyperglycaemic
conditions in vitro [123,124]. Moreover, in myotubes cultured
from human muscle satellite cells, hyperglycaemia induces
SREBP-1c expression and lipogenesis, with a concomitant
increase in TAG content [123,124]. Therefore the specialization
and sequential nature of DGAT2 and DGAT1 activities may
explain why both insulin-resistant and insulin-sensitive muscle
can have equivalent amounts of overall cellular TAG, but have
opposing phenotypes (and different LD morphology). It is of note
that overexpression of DGAT1 in hearts of mice transgenic for
PPARγ (peroxisome-proliferator-activated receptor γ ) rescues
the myopathic phenotype of the animals without decreasing the
overall TAG content of the cardiomyocytes, the only observable
difference between the two phenotypes being the change in morphology of the LDs in cardiac myocytes from small to large [5].
In adipocytes, de novo fatty acid synthesis from glucose may
be an important process depending on the species and the fat
content of the diet. Therefore DGAT2 would be expected to be
important in the function of the tissue. Accordingly, both DGAT1
and DGAT2 were found to be important for TAG synthesis and
LD formation in adipocytes [50]. In differentiating adipocytes, the
induction of DGAT2 was found to be dependent on the activation
of C/EBP (CCAAT/enhancer-binding protein) β [125] and to be
maintained by C/EBPα, after initial differentiation, indicating a
central role for the enzyme in adipose tissue function.
HEPATIC DGAT2 AS THE LINK BETWEEN GLYCAEMIA AND
TRIGLYCERIDAEMIA
It is well established that a crucial factor in driving
hypertriglyceridaemia is insulin-stimulated hepatic de novo
lipogenesis mediated by increased nuclear SREBP-1c [80,97]. In
this respect, DGAT2 emerges as the link between glycaemia (and
its attendant insulinaemia) and the net new formation of TAG. It
occupies a fundamentally important node at which carbohydrate,
fatty acid and TAG metabolism intersect (see Figure 4). Its
relatively low activity within hepatocytes is rate-limiting for the
de novo synthesis of TAG. The interaction of DGAT2 with an
expanding list of other proteins [SCD1, GPAT and ACS (acyl-CoA
synthetase)–FATP1, see above] is probably required to create its
own microcompartment in the cell, within which newly synthesized fatty acids are channelled towards the de novo synthesis of
DAG and of DGAT2-catalysed TAG synthesis. Therefore, through
DGAT2 action, the lipogenesis that is stimulated by insulin (via
SREBP-1c) and carbohydrate (via ChREBP) is linked to de novo
TAG synthesis. Together with the function of DGAT1 of salvaging
partial glycerides from complete hydrolysis, DGAT2 action in the
liver is expected to result in hepatic steatosis, hypersecretion of
VLDL–TAG and the provision of substrate for the formation of
molecular species that are associated with insulin resistance.
Therefore DGAT2 plays a crucial role in integrating
carbohydrate and lipid metabolism in the liver owing to the ability
of the tissue to synthesize both fatty acids and TAG de novo, and to
use the products of both of these processes as substrates for the enzyme. Inhibition of DGAT2 pharmacologically in the liver would
be expected to break this link and to prevent the carbohydratemediated steatosis, insulin resistance and hypertriglyceridaemia
observed under hyperinsulinaemic conditions. The selective
insulin resistance [80,81] which enables both increased glucose
output from the liver concomitantly with increased lipogenesis
places DGAT2 at the centre of the glucose and fatty acid
disturbances of the metabolic syndrome and Type 2 diabetes.
ACKNOWLEDGEMENTS
I thank the many research colleagues who have contributed to the work described in the
present review.
FUNDING
The work of the author is supported by the Medical Research Council (UK), Diabetes UK,
the British Heart Foundation and the Biotechnology and Biological Sciences Research
Council (UK).
c The Authors Journal compilation c 2013 Biochemical Society
10
V. A. Zammit
REFERENCES
1 White, D. L., Kanwal, F. and El-Serag, H. B. (2012) Association between nonalcoholic
fatty liver disease and risk for hepatocellular cancer, based on systematic review. Clin.
Gastroenterol. Hepatol. 10, 1342–1359
2 Finck, B. N., Lehman, J. J., Leone, T. C., Welch, M. J., Bennett, M. J., Kovacs, A., Han,
X., Gross, R. W., Kozak, R., Lopaschuk, G. D. and Kelly, D. P. (2002) The cardiac
phenotype induced by PPARα overexpression mimics that caused by diabetes mellitus.
J. Clin. Invest. 109, 121–130
3 Schaffer, J. E. (2003) Lipotoxicity: when tissues overeat. Curr. Opin. Lipidol. 14,
281–287
4 Unger, R. H. (2002) Lipotoxic diseases. Annu. Rev. Med. 53, 319–336
5 Son, N. H., Yu, S., Tuinei, J., Arai, K., Hamai, H., Homma, S., Shulman, G. I., Abel, E. D.
and Goldberg, I. J. (2010) PPARγ -induced cardiolipotoxicity in mice is ameliorated by
PPARα deficiency despite increases in fatty acid oxidation. J. Clin. Invest. 120,
3443–3454
6 Pan, D. A., Lillioja, S., Kriketos, A. D., Milner, M. R., Baur, L. A., Bogardus, C., Jenkins,
A. B. and Storlien, L. H. (1997) Skeletal muscle triglyceride levels are inversely related to
insulin action. Diabetes 46, 983–988
7 Kelley, D. E., Goodpaster, B. H. and Storlien, L. (2002) Muscle triglyceride and insulin
resistance. Annu. Rev. Nutr. 22, 325–346
8 Schenk, S. and Horowitz, J. F. (2007) Acute exercise increases triglyceride synthesis in
skeletal muscle and prevents fatty acid-induced insulin resistance. J. Clin. Invest. 117,
1690–1698
9 van Loon, L. J., Manders, R. J., Koopman, R., Kaastra, B., Stegen, J. H., Gijsen, A. P.,
Saris, W. H. and Keizer, H. A. (2005) Inhibition of adipose tissue lipolysis increases
intramuscular lipid use in type 2 diabetic patients. Diabetologia 48, 2097–2107
10 Zammit, V. A. (1999) The malonyl-CoA-long-chain acyl-CoA axis in the maintenance of
mammalian cell function. Biochem. J. 343, 505–515
11 Summers, S. A., Garza, L. A., Zhou, H. and Birnbaum, M. J. (1998) Regulation of
insulin-stimulated glucose transporter GLUT4 translocation and Akt kinase activity by
ceramide. Mol. Cell. Biol. 18, 5457–5464
12 Liu, L., Zhang, Y., Chen, N., Shi, X., Tsang, B. and Yu, Y. H. (2007) Upregulation of
myocellular DGAT1 augments triglyceride synthesis in skeletal muscle and protects
against fat-induced insulin resistance. J. Clin. Invest. 117, 1679–1689
13 Yu, C., Chen, Y., Cline, G. W., Zhang, D., Zong, H., Wang, Y., Bergeron, R., Kim, J. K.,
Cushman, S. W., Cooney, G. J. et al. (2002) Mechanism by which fatty acids inhibit
insulin activation of insulin receptor substrate-1 (IRS-1)-associated
phosphatidylinositol 3-kinase activity in muscle. J. Biol. Chem. 277, 50230–50236
14 Griffin, M. E., Marcucci, M. J., Cline, G. W., Bell, K., Barucci, N., Lee, D., Goodyear, L.
J., Kraegen, E. W., White, M. F. and Shulman, G. I. (1999) Free fatty acid-induced insulin
resistance is associated with activation of protein kinase Cθ and alterations in the
insulin signaling cascade. Diabetes 48, 1270–1274
15 Samuel, V. T. and Shulman, G. I. (2012) Mechanisms for insulin resistance: common
threads and missing links. Cell 148, 852–871
16 Chavez, J. A. and Summers, S. A. (2012) A ceramide-centric view of insulin resistance.
Cell Metab. 15, 585–594
17 Lark, D. S., Fisher-Wellman, K. H. and Neufer, P. D. (2012) High-fat load: mechanism(s)
of insulin resistance in skeletal muscle. Int. J. Obes. Suppl. 2, S31–S36
18 Adiels, M., Boren, J., Caslake, M. J., Stewart, P., Soro, A., Westerbacka, J., Wennberg,
B., Olofsson, S. O., Packard, C. and Taskinen, M. R. (2005) Overproduction of VLDL1
driven by hyperglycemia is a dominant feature of diabetic dyslipidemia. Arterioscler.,
Thromb., Vasc. Biol. 25, 1697–1703
19 Taskinen, M. R. (2001) Pathogenesis of dyslipidemia in type 2 diabetes. Exp. Clin.
Endocrinol. Diabetes 109 Suppl. 2, S180–S188
20 Malmstrom, R., Packard, C. J., Caslake, M., Bedford, D., Stewart, P., Yki-Jarvinen, H.,
Shepherd, J. and Taskinen, M. R. (1997) Defective regulation of triglyceride metabolism
by insulin in the liver in NIDDM. Diabetologia 40, 454–462
21 Abbasi, F., McLaughlin, T., Lamendola, C. and Reaven, G. M. (2000) Insulin regulation
of plasma free fatty acid concentrations is abnormal in healthy subjects with muscle
insulin resistance. Metabolism 49, 151–154
22 Flowers, M. T. and Ntambi, J. M. (2009) Stearoyl-CoA desaturase and its relation to
high-carbohydrate diets and obesity. Biochim. Biophys. Acta 1791, 85–91
23 McLaughlin, T., Abbasi, F., Lamendola, C., Yeni-Komshian, H. and Reaven, G. (2000)
Carbohydrate-induced hypertriglyceridemia: an insight into the link between plasma
insulin and triglyceride concentrations. J. Clin. Endocrinol. Metab. 85, 3085–3088
24 Olofsson, S. O., Asp, L. and Boren, J. (1999) The assembly and secretion of
apolipoprotein B-containing lipoproteins. Curr. Opin. Lipidol. 10, 341–346
25 Cartwright, I. J. and Higgins, J. A. (2001) Direct evidence for a two step assembly of
apo-B48 containing lipoproteins in the lumen of the smooth endoplasmic reticulum of
rabbit enterocytes. J. Biol. Chem. 276, 48048–48057
c The Authors Journal compilation c 2013 Biochemical Society
26 Liu, P., Ying, Y., Zhao, Y., Mundy, D. I., Zhu, M. and Anderson, R. G. (2004) Chinese
hamster ovary K2 cell lipid droplets appear to be metabolic organelles involved in
membrane traffic. J. Biol. Chem. 279, 3787–3792
27 Brasaemle, D. L. and Wolins, N. E. (2012) Packaging of fat: an evolving model of lipid
droplet assembly and expansion. J. Biol. Chem. 287, 2273–2279
28 Wolins, N. E., Brasaemle, D. L. and Bickel, P. E. (2006) A proposed model of fat
packaging by exchangeable lipid droplet proteins. FEBS Lett. 580, 5484–5491
29 Alexander, C. A., Hamilton, R. L. and Havel, R. J. (1976) Subcellular localization of B
apoprotein of plasma lipoproteins in rat liver. J. Cell Biol. 69, 241–263
30 Wang, H., Wei, E., Quiroga, A. D., Sun, X., Touret, N. and Lehner, R. (2010) Altered lipid
droplet dynamics in hepatocytes lacking triacylglycerol hydrolase expression. Mol. Biol.
Cell 21, 1991–2000
31 Kulinski, A., Rustaeus, S. and Vance, J. E. (2002) Microsomal triacylglycerol transfer
protein is required for lumenal accretion of triacylglycerol not associated with ApoB, as
well as for ApoB lipidation. J. Biol. Chem. 277, 31516–31525
32 Gotto, Jr, A. M. (1998) Triglyceride as a risk factor for coronary artery disease. Am. J.
Cardiol. 82, 22Q–25Q
33 Wiggins, D. and Gibbons, G. F. (1992) The lipolysis/esterification cycle of hepatic
triacylglycerol. Its role in the secretion of very-low-density-lipoprotein and its response
to hormones and sulphonylureas. Biochem. J. 284, 457–462
34 Lankester, D., Brown, A. and Zammit, V. (1998) Use of cytosolic triacylglycerol
hydrolysis products and of exogenous fatty cid for the synthesis of triacylglycerol
secreted by cultured hepatocytes. J. Lipid Res. 32, 1635–1645
35 Yang, L. Y., Kuksis, A., Myher, J. J. and Steiner, G. (1995) Origin of triacylglycerol
moiety of plasma very low density lipoproteins in the rat: structural studies. J. Lipid Res.
36, 125–136
36 Yang, L. Y., Kuksis, A., Myher, J. J. and Steiner, G. (1996) Contribution of de novo fatty
acid synthesis to very low density lipoprotein triacylglycerols: evidence from mass
isotopomer distribution analysis of fatty acids synthesized from [2 H6 ]ethanol. J. Lipid
Res. 37, 262–274
37 Owen, M., Corstorphine, C. G. and Zammit, V. A. (1997) Overt and latent activities of
diacylglycerol acyltransferase in rat liver microsomes: possible roles in very-low-density
lipoprotein triacylglycerol secretion. Biochem. J. 323, 17–21
38 Owen, M. and Zammit, V. A. (1997) Evidence for overt and latent forms of DGAT in rat
liver microsomes. Implications for the pathways of triacylglycerol incorporation into
VLDL. Biochem. Soc. Trans. 25, 21S
39 Allan, D. and Thomas, P. (1978) Rapid transbilayer diffusion of 1,2-diacylglycerol and its
relevance to control of membrane curvature. Nature 276, 289–290
40 Abo-Hashema, K. A., Cake, M. H., Power, G. W. and Clarke, D. (1999) Evidence for
triacylglycerol synthesis in the lumen of microsomes via a lipolysis-esterification
pathway involving carnitine acyltransferases. J. Biol. Chem. 274, 35577–35582
41 Broadway, N., Pease, R. and Saggerson, E. (1999) Carnitine acyltransferases and
associated transport processes in the endoplasmic reticulum: missing links in the VLDL
story? In In Current Views of Fatty acid Oxidation and Ketogenesis: From Organelles to
Point Mutations (Quant, P. and Eaton, S., eds), pp. 59–68, Luwer Academic/Plenum
Publishers, New York
42 Murthy, M. S. R. and Pande, S. V. (1994) Malonyl-CoA-sensitive and -insensitive
carnitine palmitoyltransferase activities of microsomes are due to different proteins. J.
Biol. Chem. 269, 18283–18286
43 Zammit, V. A., Buckett, L. K., Turnbull, A. V., Wure, H. and Proven, A. (2008)
Diacylglycerol acyltransferases: potential roles as pharmacological targets. Pharmacol.
Ther. 118, 295–302
44 Cases, S., Smith, S. J., Zheng, Y. W., Myers, H. M., Lear, S. R., Sande, E., Novak, S.,
Collins, C., Welch, C. B., Lusis, A. J. et al. (1998) Identification of a gene encoding an
acyl CoA:diacylglycerol acyltransferase, a key enzyme in triacylglycerol synthesis. Proc.
Natl. Acad. Sci. U.S.A. 95, 13018–13023
45 Cases, S., Stone, S., Zhou, P., Yen, E., Tow, B., Lardizabal, K. D., Voelker, T. and Farese,
Jr, R. V. (2001) Cloning of DGAT2, a second mammalian diacylglycerol acyltransferase,
and related family members. J. Biol. Chem. 276, 38870–38876
46 Lardizabal, K. D., Mai, J. T., Wagner, N. W., Wyrick, A., Voelker, T. and Hawkins, D. J.
(2001) DGAT2: a new diacylglycerol acyltransferase gene family: purification, cloning
and expression in insect cells of two polypeptides from Mortierella ramannniana with
diacylglycerol acyltransferase activity. J. Biol. Chem. 276, 38862–38869
47 Cao, J., Cheng, L. and Shi, Y. (2007) Catalytic properties of MGAT3, a putative
triacylgycerol synthase. J. Lipid Res. 48, 583–591
48 Sturley, S. L. and Hussain, M. M. (2012) Lipid droplet formation on opposing sides of
the endoplasmic reticulum. J. Lipid Res. 53, 1800–1810
49 Stone, S. J., Myers, H. M., Watkins, S. M., Brown, B. E., Feingold, K. R., Elias, P. M. and
Farese, Jr, R. V. (2004) Lipopenia and skin barrier abnormalities in DGAT2-deficient
mice. J. Biol. Chem. 279, 11767–11776
DGAT2 integrates glycaemia and triglyceridaemia
50 Harris, C. A., Haas, J. T., Streeper, R. S., Stone, S. J., Kumari, M., Yang, K., Han, X.,
Brownell, N., Gross, R. W., Zechner, R. and Farese, Jr, R. V. (2011) DGAT enzymes are
required for triacylglycerol synthesis and lipid droplets in adipocytes. J. Lipid Res. 52,
657–667
51 Cases, S., Zhou, P., Shillingford, J. M., Wiseman, B. S., Fish, J. D., Angle, C. S.,
Hennighausen, L., Werb, Z. and Farese, Jr, R. V. (2004) Development of the mammary
gland requires DGAT1 expression in stromal and epithelial tissues. Development 131,
3047–3055
52 Chen, H. C., Smith, S. J., Ladha, Z., Jensen, D. R., Ferreira, L. D., Pulawa, L. K.,
McGuire, J. G., Pitas, R. E., Eckel, R. H. and Farese, Jr, R. V. (2002) Increased insulin
and leptin sensitivity in mice lacking acyl CoA:diacylglycerol acyltransferase 1. J. Clin.
Invest. 109, 1049–1055
53 Smith, S. J., Cases, S., Jensen, D. R., Chen, H. C., Sande, E., Tow, B., Sanan, D. A.,
Raber, J., Eckel, R. H. and Farese, Jr, R. V. (2000) Obesity resistance and multiple
mechanisms of triglyceride synthesis in mice lacking Dgat. Nat. Genet. 25, 87–90
54 Orland, M. D., Anwar, K., Cromley, D., Chu, C. H., Chen, L., Billheimer, J. T., Hussain,
M. M. and Cheng, D. (2005) Acyl coenzyme A dependent retinol esterification by acyl
coenzyme A: diacylglycerol acyltransferase 1. Biochim. Biophys. Acta 1737, 76–82
55 Yen, C. L., Monetti, M., Burri, B. J. and Farese, Jr, R. V. (2005) The triacylglycerol
synthesis enzyme DGAT1 also catalyzes the synthesis of diacylglycerols, waxes, and
retinyl esters. J. Lipid Res. 46, 1502–1511
56 Shockey, J. M., Gidda, S. K., Chapital, D. C., Kuan, J. C., Dhanoa, P. K., Bland, J. M.,
Rothstein, S. J., Mullen, R. T. and Dyer, J. M. (2006) Tung tree DGAT1 and DGAT2 have
nonredundant functions in triacylglycerol biosynthesis and are localized to different
subdomains of the endoplasmic reticulum. Plant Cell 18, 2294–2313
57 Gidda, S. K., Shockey, J. M., Falcone, M., Kim, P. K., Rothstein, S. J., Andrews, D. W.,
Dyer, J. M. and Mullen, R. T. (2011) Hydrophobic-domain-dependent protein-protein
interactions mediate the localization of GPAT enzymes to ER subdomains. Traffic 12,
452–472
58 Man, W. C., Miyazaki, M., Chu, K. and Ntambi, J. (2006) Colocalization of SCD1 and
DGAT2: implying preference for endogenous monounsaturated fatty acids in triglyceride
synthesis. J. Lipid Res. 47, 1928–1939
59 Xu, N., Zhang, S. O., Cole, R. A., McKinney, S. A., Guo, F., Haas, J. T., Bobba, S., Farese,
Jr, R. V. and Mak, H. Y. (2012) The FATP1-DGAT2 complex facilitates lipid droplet
expansion at the ER-lipid droplet interface. J. Cell Biol. 198, 895–911
60 Stone, S. J., Levin, M. C. and Farese, Jr, R. V. (2006) Membrane topology and
identification of key functional amino acid residues of murine acyl-CoA:diacylglycerol
acyltransferase-2. J. Biol. Chem. 281, 40273–40282
61 Stone, S. J., Levin, M. C., Zhou, P., Han, J., Walther, T. C. and Farese, Jr, R. V. (2009)
The endoplasmic reticulum enzyme DGAT2 is found in mitochondria-associated
membranes and has a mitochondrial targeting signal that promotes its association with
mitochondria. J. Biol. Chem. 284, 5352–5361
62 Wurie, H. R., Buckett, L. and Zammit, V. A. (2011) Evidence that diacylglycerol
acyltransferase 1 (DGAT1) has dual membrane topology in the endoplasmic reticulum of
HepG2 cells. J. Biol. Chem. 286, 36238–36247
63 Wurie, H. R., Buckett, L. and Zammit, V. A. (2012) Diacylglycerol acyltransferase 2 acts
upstream of diacylglycerol acyltransferase 1 and utilizes nascent diglycerides and de
novo synthesized fatty acids in HepG2 cells. FEBS J. 279, 3033–3047
64 McFie, P. J., Stone, S. L., Banman, S. L. and Stone, S. J. (2010) Topological orientation
of acyl-CoA:diacylglycerol acyltransferase-1 (DGAT1) and identification of a putative
active site histidine and the role of the N terminus in dimer/tetramer formation. J. Biol.
Chem. 285, 37377–37387
65 Yamazaki, T., Sasaki, E., Kakinuma, C., Yano, T., Miura, S. and Ezaki, O. (2005)
Increased very low density lipoprotein secretion and gonadal fat mass in mice
overexpressing liver DGAT1. J. Biol. Chem. 280, 21506–21514
66 Cheng, D., Meegalla, R. L., He, B., Cromley, D. A., Billheimer, J. T. and Young, P. R.
(2001) Human acyl-CoA:diacylglycerol acyltransferase is a tetrameric protein. Biochem.
J. 359, 707–714
67 Chen, Y. J., Pornillos, O., Lieu, S., Ma, C., Chen, A. P. and Chang, G. (2007) X-ray
structure of EmrE supports dual topology model. Proc. Natl. Acad. Sci. U.S.A. 104,
18999–19004
68 Lolkema, J. S., Dobrowolski, A. and Slotboom, D. J. (2008) Evolution of antiparallel
two-domain membrane proteins: tracing multiple gene duplication events in the DUF606
family. J. Mol. Biol. 378, 596–606
69 Rapp, M., Granseth, E., Seppala, S. and von Heijne, G. (2006) Identification and
evolution of dual-topology membrane proteins. Nat. Struct. Mol. Biol. 13, 112–116
70 Sebag, J. A. and Hinkle, P. M. (2009) Regions of melanocortin 2 (MC2) receptor
accessory protein necessary for dual topology and MC2 receptor trafficking and
signaling. J. Biol. Chem. 284, 610–618
71 Gouffi, K., Gerard, F., Santini, C. L. and Wu, L. F. (2004) Dual topology of the Escherichia
coli TatA protein. J. Biol. Chem. 279, 11608–11615
11
72 Waterman, I. J. and Zammit, V. A. (2002) Differential effects of fenofibrate or simvastatin
treatment of rats on hepatic microsomal overt and latent diacylglycerol acyltransferase
activities. Diabetes 51, 1708–1713
73 Bogdanov, M. and Dowhan, W. (2012) Lipid-dependent generation of dual topology for a
membrane protein. J. Biol. Chem. 287, 37939–37948
74 Dowhan, W. and Bogdanov, M. (2012) Molecular genetic and biochemical approaches
for defining lipid-dependent membrane protein folding. Biochim. Biophys. Acta 1818,
1097–1107
75 Quiroga, A. D. and Lehner, R. (2011) Role of endoplasmic reticulum neutral lipid
hydrolases. Trends Endocrinol. Metab. 22, 218–225
76 Azain, M. J., Fukuda, N., Chao, F. F., Yamamoto, M. and Ontko, J. A. (1985)
Contributions of fatty acid and sterol synthesis to triglyceride and cholesterol secretion
by the perfused rat liver in genetic hyperlipemia and obesity. J. Biol. Chem. 260,
174–181
77 Fukuda, N., Azain, M. J. and Ontko, J. A. (1982) Altered hepatic metabolism of free fatty
acids underlying hypersecretion of very low density lipoproteins in the genetically obese
Zucker rats. J. Biol. Chem. 257, 14066–14072
78 Fukuda, N. and Ontko, J. A. (1984) Interactions between fatty acid synthesis, oxidation,
and esterification in the production of triglyceride-rich lipoproteins by the liver. J. Lipid
Res. 25, 831–842
79 Miyazaki, M., Kim, Y. C., Gray-Keller, M. P., Attie, A. D. and Ntambi, J. M. (2000) The
biosynthesis of hepatic cholesterol esters and triglycerides is impaired in mice with a
disruption of the gene for stearoyl-CoA desaturase 1. J. Biol. Chem. 275, 30132–30138
80 Shimomura, I., Bashmakov, Y., Ikemoto, S., Horton, J. D., Brown, M. S. and Goldstein, J.
L. (1999) Insulin selectively increases SREBP-1c mRNA in the livers of rats with
streptozotocin-induced diabetes. Proc. Natl. Acad. Sci. U.S.A. 96, 13656–13661
81 Brown, M. S. and Goldstein, J. L. (2008) Selective versus total insulin resistance: a
pathogenic paradox. Cell Metab. 7, 95–96
82 Chu, K., Miyazaki, M., Man, W. C. and Ntambi, J. M. (2006) Stearoyl-coenzyme A
desaturase 1 deficiency protects against hypertriglyceridemia and increases plasma
high-density lipoprotein cholesterol induced by liver X receptor activation. Mol. Cell.
Biol. 26, 6786–6798
83 Hellerstein, M. K., Schwarz, J. M. and Neese, R. A. (1996) Regulation of hepatic de novo
lipogenesis in humans. Annu. Rev. Nutr. 16, 523–557
84 Hudgins, L. C., Hellerstein, M., Seidman, C., Neese, R., Diakun, J. and Hirsch, J. (1996)
Human fatty acid synthesis is stimulated by a eucaloric low fat, high carbohydrate diet. J.
Clin. Invest. 97, 2081–2091
85 Francone, O. L., Kalopissis, A. D. and Griffaton, G. (1989) Contribution of cytoplasmic
storage triacylglycerol to VLDL-triacylglycerol in isolated rat hepatocytes. Biochim.
Biophys. Acta 1002, 28–36
86 Zammit, V. A. (1996) Role of insulin in hepatic fatty acid partitioning: emerging
concepts. Biochem. J. 314, 1–14
87 Qi, J., Lang, W., Geisler, J. G., Wang, P., Petrounia, I., Mai, S., Smith, C., Askari, H.,
Struble, G. T., Williams, R. et al. (2012) The use of stable isotope-labeled glycerol and
oleic acid to differentiate the hepatic functions of diacylglycerol acyltransferase-1 and -2.
J. Lipid Res. 53, 1106–1116
88 Aarsland, A., Chinkes, D. and Wolfe, R. R. (1996) Contributions of de novo synthesis of
fatty acids to total VLDL-triglyceride secretion during prolonged hyperglycemia/
hyperinsulinemia in normal man. J. Clin. Invest. 98, 2008–2017
89 Miyazaki, M., Kim, Y. C. and Ntambi, J. M. (2001) A lipogenic diet in mice with a
disruption of the stearoyl-CoA desaturase 1 gene reveals a stringent requirement of
endogenous monounsaturated fatty acids for triglyceride synthesis. J. Lipid Res. 42,
1018–1024
90 Kuerschner, L., Moessinger, C. and Thiele, C. (2008) Imaging of lipid biosynthesis: how
a neutral lipid enters lipid droplets. Traffic 9, 338–352
91 Millar, J. S., Stone, S. J., Tietge, U. J., Tow, B., Billheimer, J. T., Wong, J. S., Hamilton,
R. L., Farese, Jr, R. V. and Rader, D. J. (2006) Short-term hepatic overexpression of
DGAT1 or DGAT2 in female mice increases hepatic triglyceride synthesis without
changing VLDL triglyceride or ApoB production. J. Lipid Res. 47, 2297–2305
92 Villanueva, C. J., Monetti, M., Shih, M., Zhou, P., Watkins, S. M., Bhanot, S. and Farese,
Jr, R. V. (2009) Specific role for acyl CoA:diacylglycerol acyltransferase 1 (Dgat1) in
hepatic steatosis due to exogenous fatty acids. Hepatology 50, 434–442
93 Ganji, S. H., Tavintharan, S., Zhu, D., Xing, Y., Kamanna, V. S. and Kashyap, M. L. (2004)
Niacin noncompetitively inhibits DGAT2 but not DGAT1 activity in HepG2 cells. J. Lipid
Res. 45, 1835–1845
94 Hu, M., Chu, W. C., Yamashita, S., Yeung, D. K., Shi, L., Wang, D., Masuda, D., Yang, Y.
and Tomlinson, B. (2012) Liver fat reduction with niacin is influenced by DGAT-2
polymorphisms in hypertriglyceridemic patients. J. Lipid Res. 53, 802–809
95 Tobey, T. A., Greenfield, M., Kraemer, F. and Reaven, G. M. (1981) Relationship between
insulin resistance, insulin secretion, very low density lipoprotein kinetics, and plasma
triglyceride levels in normotriglyceridemic man. Metabolism 30, 165–171
c The Authors Journal compilation c 2013 Biochemical Society
12
V. A. Zammit
96 Chirieac, D. V., Chirieac, L. R., Corsetti, J. P., Cianci, J., Sparks, C. E. and Sparks, J. D.
(2000) Glucose-stimulated insulin secretion suppresses hepatic triglyceride-rich
lipoprotein and apoB production. Am. J. Physiol. Endocrinol. Metab. 279, E1003–E1011
97 Shimomura, I., Matsuda, M., Hammer, R. E., Bashmakov, Y., Brown, M. S. and
Goldstein, J. L. (2000) Decreased IRS-2 and increased SREBP-1c lead to mixed insulin
resistance and sensitivity in livers of lipodystrophic and ob /ob mice. Mol. Cell 6, 77–86
98 Biddinger, S. B., Hernandez-Ono, A., Rask-Madsen, C., Haas, J. T., Aleman, J. O.,
Suzuki, R., Scapa, E. F., Agarwal, C., Carey, M. C., Stephanopoulos, G. et al. (2008)
Hepatic insulin resistance is sufficient to produce dyslipidemia and susceptibility to
atherosclerosis. Cell Metab. 7, 125–134
99 Rennie, S. M., Park, B. S. and Zammit, V. A. (2000) A switch in the direction of the effect
of insulin on the partitioning of hepatic fatty acids for the formation of secreted
triacylglycerol occurs in vivo , as predicted from studies with perfused livers. Eur. J.
Biochem. 267, 935–941
100 Zammit, V. A. (2002) Insulin stimulation of hepatic triacylglycerol secretion in the
insulin-replete state: implications for the etiology of peripheral insulin resistance. Ann.
N. Y. Acad. Sci. 967, 52–65
101 Zammit, V. A., Waterman, I. J., Topping, D. and McKay, G. (2001) Insulin stimulation of
hepatic triacylglycerol secretion and the etiology of insulin resistance. J. Nutr. 131,
2074–2077
102 Zammit, V. A., Lankester, D. J., Brown, A. M. and Park, B. S. (1999) Insulin stimulates
triacylglycerol secretion by perfused livers from fed rats but inhibits it in livers from
fasted or insulin-deficient rats implications for the relationship between
hyperinsulinaemia and hypertriglyceridaemia. Eur. J. Biochem. 263, 859–864
103 Wiggins, D., Hems, R. and Gibbons, G. F. (1995) Decreased sensitivity to the inhibitory
effect of insulin on the secretion of very-low-density lipoprotein in cultured hepatocytes
from fructose-fed rats. Metabolism 44, 841–847
104 Moir, A. M. and Zammit, V. A. (1994) Effects of insulin treatment of diabetic rats on
hepatic partitioning of fatty acids between oxidation and esterification, phospholipid and
acylglycerol synthesis, and on the fractional rate of secretion of triacylglycerol in vivo .
Biochem. J. 304, 177–182
105 Zammit, V. A. and Moir, A. M. (1994) Monitoring the partitioning of hepatic fatty acids
in vivo : keeping track of control. Trends Biochem. Sci. 19, 313–317
106 Li, S., Brown, M. S. and Goldstein, J. L. (2010) Bifurcation of insulin signaling pathway
in rat liver: mTORC1 required for stimulation of lipogenesis, but not inhibition of
gluconeogenesis. Proc. Natl. Acad. Sci. U.S.A. 107, 3441–3446
107 Hsieh, J., Hayashi, A. A., Webb, J. and Adeli, K. (2008) Postprandial dyslipidemia in
insulin resistance: mechanisms and role of intestinal insulin sensitivity. Atherosclerosis
Suppl. 9, 7–13
108 Cheng, D., Nelson, T. C., Chen, J., Walker, S. G., Wardwell-Swanson, J., Meegalla, R.,
Taub, R., Billheimer, J. T., Ramaker, M. and Feder, J. N. (2003) Identification of acyl
coenzyme A:monoacylglycerol acyltransferase 3, an intestinal specific enzyme
implicated in dietary fat absorption. J. Biol. Chem. 278, 13611–13614
109 Buhman, K. K., Smith, S. J., Stone, S. J., Repa, J. J., Wong, J. S., Knapp, Jr, F. F., Burri,
B. J., Hamilton, R. L., Abumrad, N. A. and Farese, Jr, R. V. (2002) DGAT1 is not essential
for intestinal triacylglycerol absorption or chylomicron synthesis. J. Biol. Chem. 277,
25474–25479
110 Ables, G. P., Yang, K. J., Vogel, S., Hernandez-Ono, A., Yu, S., Yuen, J. J., Birtles, S.,
Buckett, L. K., Turnbull, A. V., Goldberg, I. J. et al. (2012) Intestinal DGAT1 deficiency
reduces postprandial triglyceride and retinyl ester excursions by inhibiting chylomicron
secretion and delaying gastric emptying. J. Lipid Res. 53, 2364–2379
Received 6 November 2012/3 January 2013; accepted 25 January 2013
Published on the Internet 14 March 2013, doi:10.1042/BJ20121689
c The Authors Journal compilation c 2013 Biochemical Society
111 Lee, B., Fast, A. M., Zhu, J., Cheng, J. X. and Buhman, K. K. (2010) Intestine-specific
expression of acyl CoA:diacylglycerol acyltransferase 1 reverses resistance to
diet-induced hepatic steatosis and obesity in Dgat1 − / − mice. J. Lipid Res. 51,
1770–1780
112 Haidari, M., Leung, N., Mahbub, F., Uffelman, K. D., Kohen-Avramoglu, R., Lewis, G. F.
and Adeli, K. (2002) Fasting and postprandial overproduction of intestinally derived
lipoproteins in an animal model of insulin resistance. Evidence that chronic fructose
feeding in the hamster is accompanied by enhanced intestinal de novo lipogenesis
and ApoB48-containing lipoprotein overproduction. J. Biol. Chem. 277,
31646–31655
113 Levin, M. C., Monetti, M., Watt, M. J., Sajan, M. P., Stevens, R. D., Bain, J. R., Newgard,
C. B., Farese, Sr, R. V. and Farese, Jr, R. V. (2007) Increased lipid accumulation and
insulin resistance in transgenic mice expressing DGAT2 in glycolytic (type II) muscle.
Am. J. Physiol. Endocrinol. Metab. 293, E1772–E1781
114 Pan, D. A., Mater, M. K., Thelen, A. P., Peters, J. M., Gonzalez, F. J. and Jump, D. B.
(2000) Evidence against the peroxisome proliferator-activated receptor α (PPARα) as
the mediator for polyunsaturated fatty acid suppression of hepatic L-pyruvate kinase
gene transcription. J. Lipid Res. 41, 742–751
115 Goodpaster, B. H., He, J., Watkins, S. and Kelley, D. E. (2001) Skeletal muscle lipid
content and insulin resistance: evidence for a paradox in endurance-trained athletes. J.
Clin. Endocrinol. Metab. 86, 5755–5761
116 Russell, A. P. (2004) Lipotoxicity: the obese and endurance-trained paradox. Int. J. Obes.
Relat. Metab. Disord. 28 (Suppl. 4), S66–S71
117 Bengtsson, G. and Olivecrona, T. (1980) Lipoprotein lipase. Mechanism of product
inhibition. Eur. J. Biochem. 106, 557–562
118 Fielding, B. A. and Frayn, K. N. (1998) Lipoprotein lipase and the disposition of dietary
fatty acids. Br. J. Nutr. 80, 495–502
119 Erion, D. M. and Shulman, G. I. (2010) Diacylglycerol-mediated insulin resistance. Nat.
Med. 16, 400–402
120 Liu, L., Shi, X., Bharadwaj, K. G., Ikeda, S., Yamashita, H., Yagyu, H., Schaffer, J. E., Yu,
Y. H. and Goldberg, I. J. (2009) DGAT1 expression increases heart triglyceride content
but ameliorates lipotoxicity. J. Biol. Chem. 284, 36312–36323
121 Dube, J. J., Amati, F., Stefanovic-Racic, M., Toledo, F. G., Sauers, S. E. and Goodpaster,
B. H. (2008) Exercise-induced alterations in intramyocellular lipids and insulin
resistance: the athlete’s paradox revisited. Am. J. Physiol. Endocrinol. Metab. 294,
E882–E888
122 Amati, F., Dube, J. J., Alvarez-Carnero, E., Edreira, M. M., Chomentowski, P., Coen,
P. M., Switzer, G. E., Bickel, P. E., Stefanovic-Racic, M., Toledo, F. G. and Goodpaster, B.
H. (2011) Skeletal muscle triglycerides, diacylglycerols, and ceramides in insulin
resistance: another paradox in endurance-trained athletes? Diabetes 60,
2588–2597
123 Aas, V., Kase, E. T., Solberg, R., Jensen, J. and Rustan, A. C. (2004) Chronic
hyperglycaemia promotes lipogenesis and triacylglycerol accumulation in human
skeletal muscle cells. Diabetologia 47, 1452–1461
124 Turcotte, L. P., Swenberger, J. R. and Yee, A. J. (2002) High carbohydrate availability
increases LCFA uptake and decreases LCFA oxidation in perfused muscle. Am. J.
Physiol. Endocrinol. Metab. 282, E177–E183
125 Payne, V. A., Au, W. S., Gray, S. L., Nora, E. D., Rahman, S. M., Sanders, R., Hadaschik,
D., Friedman, J. E., O’Rahilly, S. and Rochford, J. J. (2007) Sequential regulation of
diacylglycerol acyltransferase 2 expression by CAAT/enhancer-binding protein β
(C/EBPβ) and C/EBPα during adipogenesis. J. Biol. Chem. 282, 21005–21014