biogenesis and catabolism of diacylglycerols - role of
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Thomas O. Eichmann
Biogenesis and Catabolism of Diacylglycerols
- Role of Stereochemistry -
DOCTORAL THESIS/Dissertation
submitted to the Faculty of Natural Sciences
at the University of Graz (Austria)
for attainment of the degree
Doctor of Natural Sciences
Doctor rerum naturalium
(Dr. rer. nat.)
October 2012
Sometimes the questions are complicated and the answers are simple.
Theodor Seuss Geisel, US author
Gewidmet jenen, die mich vorbehaltlos unterstützen.
- Meine Familie, Freundin und Freunde -
Preface
Herewith I declare that this doctoral thesis has been written independently and without any
assistance from third parties. Moreover, references employed for the composition of this manuscript
are cited in the bibliography and no other sources were used.
____________________________ _______________
Mag. rer. nat. Thomas O. Eichmann Date (22.10.2012)
Acknowledgement/Danksagung
I want to express my sincerest thanks to…
… Rudolf Zechner for giving me the great opportunity to work in this outstanding laboratory.
Moreover, I want to thank for his supervision, motivation and support.
… Achim Lass for all the discussions and his invaluable guidance and permanent
encouragement throughout all these years.
… Robert Zimmermann, Günter Hämmerle, Karin Preiss-Landl and Fritz Spener for their
encouragement.
… the Doktoratskolleg (DK) Molecular Enzymology for funding my Ph.D. work and giving me
the opportunity to exchange and discuss ideas and problems with other great scientists.
… Robert V. Farese Jr. and Günter Hämmerle for contributing to this work by providing
enzymes and mouse models, respectively.
… all members of our laboratories that made this institute a wonderful place to work. In
particular, I want to thank those people who became more than just colleagues, Martina
Schweiger, Gabriele Schoiswohl, Franz Radner, Christoph Heier, Manju Kumari,
Chandramohan Chitraju, Matthias Romauch, Tarek Moustafa, Harald Hofbauer, Martin
Kreim, Kathrin Zierler, Ulrike Taschler, and Renate Schreiber for a great time.
Mein größter Dank aber gilt meiner ganzen Familie und meiner Freundin. Meinen
Eltern Walter und Inge und meinen Geschwistern Michael, Katja, Jörg und Teresa, welche
mich in jeder Lebenslage vorbehaltlos unterstützen und immer hinter mir stehen. Großer
Dank gilt auch meiner Freundin Anja, die mir immer zur Seite steht und auch in schweren
Zeiten eine unschätzbare Stütze ist. Ich danke auch meinen Freunden, Florian, Patrick,
Manfred und Rupert, welche mich seit Langem begleiten und mir stets Rückhalt bieten.
Abstract of the Doctoral Thesis Submitted to the Faculty of Natural Sciences at the University of Graz for Attainment of the Degree Doctor of Natural Sciences
Biogenesis and Catabolism of Diacylglycerols
- Role of Stereochemistry -
Mag. rer. nat. Thomas O. Eichmann
Institute of Molecular Biosciences
University of Graz
In mammals, excessive energy is stored in form of energy-dense triacylglycerol (TAG). A tight balance
between TAG synthesis and degradation is instrumental for maintaining energy homeostasis.
Dysregulation of energy homeostasis is strongly associated with obesity and related disorders, like
insulin resistance (IR) and type 2 diabetes mellitus (T2DM). Adipose triglyceride lipase (ATGL) is rate-
limiting for the initial step of TAG hydrolysis, generating diacylglycerol (DAG) and fatty acids (FAs).
Accumulations of both lipids are correlated to defective insulin signaling, hence provoking IR and
T2DM. DAG exist in three different stereochemical isoforms (sn-1,2; sn-1,3; sn-2,3), of which
exclusively sn-1,2 DAG affects insulin signaling via activation of protein kinase C (PKC). So far the FA-
and positional- (stereo-) selectivity of ATGL-dependent TAG hydrolysis is unknown. Yet a direct link
between an imbalance in TAG hydrolysis and defective insulin signaling is likely. The objective of this
study focused on the eludication of the FA- and stereoselectivity of ATGL as well as the
stereoselectivity of enzymes involved in further DAG utilization. Results reveal that ATGL exhibits a
strong preference for the hydrolysis of long-chain FA esters at the sn-2 position of TAG. This
selectivity broadens to the sn-1 position upon stimulation of the enzyme by its co-activator
comparative gene identification-58. Furthermore, ATGL-derived sn-1,3 DAGs are the preferred
substrate for the consecutive hydrolysis by hormone-sensitive lipase. Interestingly, also
diacylglycerol-O-acyltransferase 2 (DGAT2), which is a key enzyme of TAG synthesis, preferentially
esterifies sn-1,3 DAG. This suggests, that ATGL and DGAT2 act coordinately in the hydrolysis/re-
esterification of TAG and that ATGL creates a distinct pool of sn-1,3 DAG. The inability of ATGL to
generate sn-1,2 DAG suggests that TAG-derived DAG cannot directly impair insulin signaling via PKC
activation.
Table of contents
Introduction ................................................................................................................................... 1
Stereochemistry of DAG ........................................................................................................... 3
Metabolic formation of DAG .................................................................................................. 6
A) Formation of DAG by intracellular lipases ............................................................................ 7
B) De novo synthesis of DAG ..................................................................................................... 12
Utilization of DAG ...................................................................................................................... 15
DAG and DAG-derived signals .............................................................................................. 20
Aim of the Thesis ..................................................................................................................... 29
Results.............................................................................................................................................. 31
I) ATGL selectivity ...................................................................................................................... 32
A) Stereo/regioselectivity ........................................................................................................... 32
B) FA selectivity ........................................................................................................................... 37
C) Substrate selectivity ............................................................................................................... 47
II) Selectivity of DAG hydrolysis and re-esterification ................................................ 51
A) Stereo/regioselectivity of DGAT enzymes ........................................................................... 51
B) Stereo/regioselectivity of HSL-dependent DAG hydrolysis ............................................... 60
Discussion ...................................................................................................................................... 64
Materials & Experimental Procedures ..................................................................... 76
I) Materials ................................................................................................................................... 77
II) Experimental Procedures .................................................................................................. 78
Publications ................................................................................................................................. 86
First author .................................................................................................................................. 87
Co-author ..................................................................................................................................... 88
Appendix ........................................................................................................................................ 90
Abbreviations & Acronyms ........................................................................................................ 91
Bibliography ................................................................................................................................ 95
Introduction
Introduction
2
Energy homeostasis constitutes a primary necessity of living organisms to ensure constant metabolic
fluxes and physiological flexibility. The storage of excessive energy metabolites displays an
evolutionary highly conserved strategy, to mobilize energy reserves and consequently facilitate
survival in times of increased energy demand or inadequate nutrient supply. In mammals, a dietary
surplus of either carbohydrates or fat is converted to inert and energy-dense triacylglycerol (TAG). In
TAG the fatty acids (FAs), which are esterified to the glycerol backbone constitute the main source of
energy substrates. In higher animals, the highly hydrophobic TAG molecules are embedded within
glycerophospholipid (PL)-coated lipid droplets (LDs) and stored in almost every cell type. In large
quantities such LDs are found in adipocytes of the white adipose tissue (WAT). An efficient
mobilization of these TAG stores is required to maintain a constant whole body energy supply. Thus
metabolic pathways which balance TAG synthesis and degradation need to be precisely regulated.
This evolutionary important ability to store excessive energy has evolved to a human burden. In the
western world countries times of caloric scarcity are rare and the oversupply of inexpensive, calorie-
dense food often goes along with little to no caloric demands. Metabolic alterations following
overnutrition lead to dysregulations of the energy equilibrium. The resulting obesity may eventually
also cause disorders such as non-alcoholic fatty liver disease, coronary heart diseases, and typ 2
diabetes mellitus (T2DM), all summarized in the term metabolic syndrome [1, 2].
The cellular metabolism of TAG is accomplished by a variety of enzymes including lipases and
acyltransferases. The intermediates diacylglycerol (DAG), FAs and coenzyme A-activated FAs (FA-CoA)
constitute indispensable, bioactive precursors that are involved in a variety of metabolic processes.
Adipose triglyceride lipase (ATGL) [3-5] represents the major lipase responsible for the initiation of
TAG mobilization in adipocytes. FAs of this reaction are thought to be released to circulation to
supply peripheral tissues either fuelling mitochondrial beta-oxidation, act as precursor for newly
synthesized lipids, or act as ligands for a variety of nuclear receptor transcription factors. The
hydrolysis of TAG by ATGL also generates DAG which may function as substrate for further
degradation, as precursor for TAG and PL synthesis, or as potential signaling molecule.
Dysregulations of ATGL, which lead to either blunted or excessive lipolysis, may also result in altered
levels of this metabolic-active lipid species in various tissues. Such fluctuations in bioactive lipid
content may have non-conceivable consequences in regard to cellular signaling and are often
interrelated with the development of diseases that are part of the metabolic syndrome.
For the understanding of the interrelation between altered energy metabolism and disturbed cellular
signaling, the biochemistry of lipid intermediates, namely DAGs and FAs, as well as their physiological
fate is of great importance.
Introduction
3
Stereochemistry of DAG
Lipolysis is the catabolic arm of TAG metabolism, which liberates FAs from inert TAG stored in LDs. It
is catalyzed by lipases, which represent a special class of serine hydrolases. Generally, hydrolases are
characterized as enzymes catalyzing the reversible, hydrolytic cleavage of various chemical bonds
including carboxylic ester, ether, amide and peptide bonds. Lipases exclusively catalyze the hydrolysis
of carboxylic esters in aqueous environment and the reverse reaction in organic milieu [6]. One
fundamental difference between hydrolases/esterases and lipases is the fact that esterases
encounter water-soluble substrates, whereas lipases are activated by oil/water interfaces (e.g. lipid
emulsions in water) [7]. This unique ability was recognized early in studies with pancreatic lipase [8,
9]. The structural factor determining interface-activation is a mobile element, called lid, which is
detectable in almost all lipases [10]. This mobile lid caps the substrate binding site in the absence of
an interface. The open form, which is thought to be stabilized in a hydrophobic environment, makes
the substrate binding site accessible and hence elevates enzymatic activity [11]. Follow-up studies
verified the fundamental difference of esterases and lipases in that esterases show normal Michaelis-
Menten kinetic whereas lipases sharply increase activity upon exceedance of the substrate solubility
limit, which yields in an interface formed by emulsified substrate molecules [12]. This privileges
lipases to release FAs during hydrolysis of lipids (e.g. glycerolipids, cholesterylesters), which are
usually stored in PL-coated micelles or are embedded within biological membranes. A second
characteristic, which distinguishes lipases from other esterases is the feature of particular
stereochemistry. Lipases can encounter both chiral and prochiral substrates. This ability makes them
unique compared to other hydrolases, like proteases, phospholipases, and nucleases, which can only
hydrolyze one optical form of their respective substrate [13, 14].
Isomers (from greek: isos – equal, mèros – part) are molecules sharing identical molecular formulas
but differ in structure. Isomerism can be divided into two main groups. On the one hand structural
isomers, which exhibit differentially linked atoms and functional groups. On the other hand spatial
isomers (stereoisomers), which display same linkage of atoms and functional groups but differ in
their geometrical position in space. A special group of stereoisomers, named enantiomers, is related
by reflection, which implies that two enantiomers are not superimposable. Furthermore,
enantiomers are characterized by an asymmetric or chiral carbon atom, featured by four different
ligands (Fig. 1).
Introduction
4
FIGURE 1. Compilation of the different forms of isomerism on the basis of DAG. DAGs feature different forms of
isomerism and can differ either in structural or spatial conformation.
DAGs arise during a variety of metabolic reactions and are important signaling molecules. They
illustrate a lipid class that exhibits different isomeric properties. TAG, one possible metabolic
precursor of DAG, contains three FAs esterified to a glycerol backbone. This implicates that TAG
provides three possible sites for lipase-dependent hydrolysis, which potentially lead to three
different DAG isoforms. Herein, the stereospecific numbering (sn) indicates the position of the FA at
the glycerol backbone (sn-1, sn-2, sn-3). Besides chirality, TAGs exhibit another important property of
lipase substrates, namely prochirality. Prochirality describes the condition that an achiral molecule
can be converted into a chiral molecule by a single step reaction. In case of TAG esterified with a
single FA species the achiral carbon atom at sn-2 position becomes a chiral center by removal of the
attached FA at either sn-1 or sn-3 position (Fig. 2).
FIGURE 2. TAG lipases can produce a new center of chirality during TAG hydrolysis. Sequence rule order: CH2-O-
COR>R2>R1 (modified after [15])
Introduction
5
Certain TAG species exhibit chemically identical but enantiotopic reactive groups (pro-R and pro-S;
e.g. oleic acid (C18:1) at sn-1 and sn-3 position), which are chirally discriminated during lipase-
dependent hydrolysis reaction, yielding a chiral DAG product. Lipase-dependent cleavage of a FA at
either sn-1 or sn-3 position of a TAG molecule leads to one of the two enantiomers, sn-2,3 or sn-1,2
DAG, respectively. These DAG enantiomers face themselves as reflection and are not
superimposable. Furthermore, enantiomeric DAGs can be classified in respect to the R/S
configuration nomenclature based on the Cahn-Ingold-Prelog (CIP) system [16, 17]. The CIP system is
used to uniquely specify enantiomeric molecules. Therefore, priorities are assigned to all groups
attached to the chiral center (CIP-rules) [16, 17]. Subsequently, the lowest ranked group is set below
the image plane and the other groups are counted starting at highest priority substituents. The
counted sequence can be either clockwise or counterclockwise and specifies the present molecule as
either R-configurated (from latin: rectus - right) or S-configurated (from latin: sinister - left). In case
of DAG, sn-1,2 DAG reflects the S-configuration whereas sn-2,3 DAG is R-configured (Fig. 3).
FIGURE 3. R/S nomenclature of DAG enantiomers according to Cahn-Ingold-Prelog convention. sn-1,2 DAG represents the
S-configuration whereas sn-2,3 DAG represents the R-configuration.
Upon hydrolysis of sn-2 bound FA esters the generated DAG exhibits sn-1,3 conformation and owns
no chiral center. In regard to the different region of hydrolysis (sn-1/sn-3: esters of a primary alcohol;
sn-2: ester of a secondary alcohol) sn-1,3 DAG is a so called regiomer (Fig. 1). Thus, lipases can
regioselectively differentiate between sn-2 and sn-1/sn-3 position or enantioselectively differ
between sn-1 and sn-3 position. Similarly, other enzymes, like acyltransferases can potentially
discriminate different isoforms of DAG.
These differences regarding isomerism of DAG as well as the selectively of metabolic DAG-
generating/consuming enzymes could be an important issue when lipids, like DAG, which display
intersections between lipid and signaling metabolism, are investigated. So, the selectivity of cellular
Introduction
6
lipases/acyltransferases as well as the impact of DAG isomerism on cellular metabolism is obviously
very important.
Metabolic formation of DAG
Intracellularly, several reactions which contribute to the generation of DAG are located at different
subcellular compartments including the endoplasmic reticulum (ER), LDs, and the plasma-membrane
(Fig. 4). Therefore, either TAGs stored in cytoplasmic or ER-associated LDs or PLs, which assemble
cellular membranes can act as source material for lipase-dependent generation of DAG (Fig. 5).
Additionally, DAGs can arise during de novo synthesis of TAG either generated by acyltransferases or
phosphohydrolases. The stereo/regioselectivity of enzymes involved and thus the isomerism of the
formed DAGs is widely unknown but might play a crucial role for subsequent cellular reactions. The
following sections describe known biochemical characteristics and stereo/regiochemical properties
of enzymes involved in the formation of DAG.
FIGURE 4. Catabolic and anabolic reactions leading to the formation of different DAG isoforms. DAG can be generated by
hydrolysis of either TAG or PLs, by dephosphorylation of phosphatidic acid, or by the esterification of monoacylglycerol
catalyzed by certain acyltransferases.
Introduction
7
A) Formation of DAG by intracellular lipases
Within most cell types, TAG turnover is crucial to balance energy storage and distribution. In whole
body energy metabolism this function is mainly achieved by specialized cells, named adipocytes,
present in WAT. In adipocytes, excessive energy is stored in form of TAG departed in cytoplasmic LDs.
Interestingly, the size of LDs is different in adipocytes as compared to other cell types. Whereas
adipocytes harbor usually a single LD in a size range about 100 µm, non-adipose tissue cells exhibit
usually multiple LDs with diameters of around 1 µm. Nevertheless, all LDs share basically the same
architecture. The core is strictly assembled of hydrophobic lipid esters, like TAG and cholesteryl
esters (CEs), and the surface is formed by a PL monolayer [18]. This monolayer harbors a variety of
anchored or embedded proteins and serves as an amphipathic shield against the aqueous milieu,
which is present in the cell [19]. The most important function of adipocyte LDs is the storage and
lipase-dependent release of energy metabolites, primarly in form of FAs. This tightly regulated
process of TAG degradation, known as lipolysis, generates stepwise lipid intermediates like FAs, DAG,
and monoacylglycerol (MAG) and is executed by a cascade of lipases. Therein, HSL was first identified
and thus thought to be essential for the initial step of TAG hydrolysis generating potential signaling
molecules, more precisely DAG and FAs.
FIGURE 5. Formation of DAG by intracellular lipases. DAG can be generated by hydrolysis of TAG or PLs. Intracellularly,
ATGL and HSL are the main TAG lipases. The generation of DAG from PLs is catalyzed by PLC. ATGL, adipose triglyceride
hydrolase; DAG, diacylglycerol; HSL, hormone-sensitive lipase; PLC, phospholipase C; TAG, triacylglycerol.
Hormone sensitive lipase (HSL)
Following the observation that WAT lipolysis is strongly inducible by hormonal stimulation, HSL was
the first characterized TAG lipase in WAT [20, 21]. HSL is known to exhibit a uniquely-broad substrate
spectrum, which includes TAG, DAG, MAG, CE, retinyl ester (RE) as well as water-soluble short chain
Introduction
8
esters [21-25]. However, in vitro studies showed that the specific activity of HSL is highest against
DAG, which exceeds those against TAG and MAG around 10-fold [23, 26]. Earlier studies described a
sn-1/(3) specificity of HSL for DAG [26], whereas most recently HSL was identified to be quite sn-3
specific [27]. Concerning substrate specificity, HSL exhibits preference for polyunsaturated FAs
(PUFAs; n-3/n-6), which was shown using crude preparations of rat HSL [28]. Although the preference
for the hydrolysis of DAG was noticed, HSL was long considered to be the rate-limiting lipase in TAG
mobilization of adipose and non-adipose tissues. Albeit HSL is expressed in many tissues, protein as
well as mRNA expression of HSL are highest in WAT and brown adipose tissue (BAT) [29]. The COOH-
terminal region of HSL harbors the α/β hydrolase fold domain, containing an active site serine (Ser423)
as part of a catalytic triad (Asp703, His733) responsible for hydrolytic activity [22, 30-32]. The lipid
binding site as well as the site responsible for protein dimerization was found at the NH2-terminal
region of HSL [33].
HSL activity is strongly regulated by hormones. A variety of phosphorylation events control HSL
activity by affecting both intracellular localization and protein-protein interactions. The major
positive stimulus is caused by catecholamines, which bind to β-adrenergic receptors during periods
of nutritional deprivation (fasting). The contrary nutritional condition (feeding) inhibits HSL action via
insulin. The activation of HSL upon β-adrenergic stimulation is primarly mediated by protein kinase A
(PKA)-dependent phosphorylation [34-40]. Additional phosphorylations, involved in the regulation of
HSL, are catalyzed by AMP-activated kinase (AMPK), extracellular signal-regulated kinase (ERK),
glycogen synthase kinase-4 as well as Ca2+/calmodulin-dependent kinase [41]. The major
phosphorylation sites controlling increased activity have been shown to be Ser659 and Ser660
(numbered for rat HSL), which are phosphorylated by either PKA or ERK [36, 42, 43]. In contrast,
phosphorylation of Ser565 by AMPK leads to antilipolytic effects by potentially antagonizing PKA-
dependent phosphorylation [34, 44]. All above mentioned phosphorylation sites are located in the
regulatory module (~150 amino acid (AA) loop) within the COOH-terminal region of the enzyme. Even
though phosphorylations of HSL are crucial, they cause only moderate changes in enzymatic activity.
The more important factor in HSL activation is the binding of the enzyme to LDs. In adipocytes, LDs
are shielded by perilipin-1, which surrounds LDs, forming a barrier between lipases and respective
lipid substrates [45]. In basal condition this LD barrier keeps lipolysis at a low rate [46]. Upon β-
adrenergic stimulation perlipin-1 is phosphorylated by PKA at six consensus serine residues (Ser81,
Ser222, Ser276, Ser433 Ser492, and Ser517; numbered for murine perilipin-1) [47-50]. This facilitates
binding of HSL to perilipin-1 at the NH2-terminal region, leading to a translocation of HSL to the LD
surface [51, 52] where it deploys full activity. The inactivation of HSL in adipocytes is triggered by
insulin as a consequence of nutrient uptake. In the cyclic AMP (cAMP)-dependent pathway, insulin
activates a variety of phosphodiesterases, which hydrolyze cAMP resulting in reduced cAMP levels
Introduction
9
and the loss of PKA activation [53, 54]. In the cAMP-independent pathway, insulin induces protein
phosphatase-1, which leads to HSL dephosphorylation and inactivation [55].
The long standing dogma that HSL acts as pacemaker of lipolysis, thereby hydrolyzing TAGs and
DAGs, was disproven when mice, carrying a global deletion of HSL (HSLko) showed no signs of obesity
on a high-fat diet (HFD) but demonstrated normal bodyweight and reduced fat mass as compared to
wildtype (wt) mice [56, 57]. Decreased adipose mass was partially explained by reduced FA
esterification counteracting decreased lipolytic activity [58]. Furthermore, HSL-deficient adipocytes
still showed a catecholamine dependent increase in FA release from WAT, which suggested an
additional lipase catalyzing lipid degradation [56, 59, 60]. The most intriguing finding in HSLko mice
was the drastic accumulation of DAG in several tissues [59] suggesting that HSL is responsible for
DAG hydrolysis.
Adipose triglyceride lipase (ATGL)
In 2004, a lipase which fulfilled all postulated requirements was identified independently by three
laboratories. The former denoted transport secretion protein-2.2 was renamed as ATGL/patatin-like
phospholipase domain containing A 2 (PNPLA2) [3], desnutrin [4], and calcium independent
phospholipase A2ζ [5].
ATGL is one of nine PNPLA family members found in humans (PNPLA1-9) [61]. The PNPLA protein
family is named after the patatin-domain, which was first identified in hydrolases of plants and
denominated after the most abundant protein of the potato tuber, patatin. Lipid hydrolases
containing this domain catalyze the non-selective hydrolysis of a variety of lipids, including PLs,
glycolipids, DAGs, and MAGs [62-64]. In addition to ATGL also other members of the PNPLA family
possess hydrolase (PNPLA3/adiponutrin; PNPLA4/gene sequence 2, GS2; PNPLA5/GS2-like),
phospholipase (PNPLA8 and 9), or lyso-phospholipase (PNPLA6 and 7) activity [5, 65, 66]. Within the
NH2-terminal half of human ATGL, the patatin domain (180 AA) is embedded in a 250 AA sized α/β/α
sandwich structure and includes a conserved serine lipase consensus sequence motif (GXSXG), which
contains the active site serine (Ser47) [66-68]. In contrast to the catalytic triad of classical lipases, the
enzymatic activity of ATGL relies on a catalytic dyad formed by Ser47 and Asp166, both identified to be
essential for hydrolytic activity [62, 69]. The COOH-terminal half of ATGL contains a lipid-binding
domain formed by a hydrophobic AA stretch (AA 315-360) [70, 71] as well as two putative
phosphorylation sites (Ser404, Ser428 in human ATGL or Ser406, Ser430 in murine ATGL) [72, 73].
Orthologs of ATGL exist in almost every eukaryotic species including invertebrates, fungi, and plants.
In contrast to the broad substrate spectrum of HSL, ATGL is highly specific for TAG and shows only
Introduction
10
weak or no activity against DAG, MAG, CE, or RE [3]. Furthermore, phospholipase A2 (PLA2) as well as
DAG transacylase activity was reported for ATGL [5, 74]. Yet, the physiological relevance of these
relatively minor activities is not established.
The key-role of ATGL in degradation of TAG is demonstrated by ATGL-deficient mice (ATGLko).
ATGLko mice exhibit 2-fold higher whole body fat mass as well as drastically enlarged adipose tissues
[75]. TAG accumulation is observable in all tissues reaching up to a 10-fold increase in TAG content
[75]. Furthermore, a dramatic accumulation of TAG in cardiomyocytes of ATGLko mice led to a
premature death of these animals due to severe cardiac dysfunction. In accordance to TAG
accumulation, isoproterenol-stimulated lipolytic activity of WAT explants is decreased around 70% as
compared to wt mice. Additionally, in vitro TAG hydrolase activity assay demonstrated the crucial
role of ATGL in the mobilization of TAG in WAT, BAT, cardiac muscle (CM), skeletal muscle (SM),
testis, and liver [75].
In mice, ATGL mRNA is expressed in all examined tissues. Highest expression is observed in WAT and
BAT. Lower expression levels are detectable in SM, CM, liver, and testis [3, 4, 66, 76]. In adipocytes,
ATGL expression is markedly upregulated during differentiation and highest when LD accumulation is
observable [3, 4]. Furthermore, ATGL shows increased expression upon fasting, treatment with
glucocorticoids [4], i.e. dexamethasone, or peroxisome proliferator activated receptor γ (PPARγ)
agonists, like thiazolidinedione [76-78]. Despite a downregulation of ATGL expression in murine
obesity models, like leptin- or leptin receptor-deficient mice (ob/ob; db/db), ATGL expression is
decreased upon feeding and by hormones associated with obesity like insulin, isoproterenol, and
TNF-α [79-82]. However, mRNA levels of ATGL do not always correlate with lipase activity, which
could be explained by substantial posttranslational regulation of ATGL. Posttranslational regulation
may include phosphorylation of Ser406 and Ser430 but reports regarding phosphorylation of ATGL are
controversial. Ser406 is phosphorylated by PKA [83] and/or AMPK [73], thereby increasing ATGL´s
hydrolase activity. In contrast, other publications showed an inhibitory effect of AMPK on TAG
hydrolysis [84] as well as a PKA-independent phosphorylation of ATGL [3].
Besides regulation via phosphorylation or nutritional condition, full stimulation of ATGL´s activity
requires the presence of an LD-associated protein named comparative gene identification-58 (CGI-
58; or α/β hydrolase fold domain containing protein 5, ABHD5) [68]. CGI-58 is highly expressed in
testis and shows lower expression in WAT, CM, SM, and liver [68]. Additionally, in vitro TAG-
hydrolase activity of cell lysates expressing murine ATGL as well as of WAT lysates is 20-fold and 2-
fold increased upon addition of exogenous CGI-58, respectively [68, 85]. Although CGI-58 belongs to
a lipase subfamily characterized by α/β hydrolase folds it does not possess intrinsic hydrolase activity
since a required putative active site serine within the canonical lipase motif (GXSXG) is replaced by an
Introduction
11
asparagine (Asp153) [86]. Noteworthy, two independent laboratories described CGI-58 as an acyl-CoA-
dependent lysophosphatidic acid acyltransferase (LPAAT) in vitro [87, 88] but metabolic implications
are so far unknown.
The molecular mechanism behind CGI-58 dependent stimulation of ATGL remains elusive. However,
a number of studies identified structural key features of both proteins. Whereas truncations of the
COOH-terminal part of ATGL increases the co-activation of ATGL by CGI-58 [70, 89], mutations within
the NH2-terminal region of ATGL result in defective CGI-58 co-activation [70]. In case of CGI-58,
truncation mutants lacking NH2-terminal part failed both to localize to the LD and to co-activate ATGL
[90]. The finding that a removal of three Trp-residues within the NH2-terminal part of CGI-58 results
in defective co-activation of ATGL but did not influence CGI-58/ATGL protein interaction suggests
that both LD binding and protein interaction is essential for activation.
A second protein has been recently identified to be strongly involved in the inhibition of ATGL
activity. The 103 AA sized G0/G1 switch gene 2 (G0S2) originally shown to be involved in cell cycle
transition selectively inhibits ATGL-dependent hydrolase activity [91]. G0S2 is found in many tissues
with highest expression levels in WAT and liver. Without competing CGI-58, G0S2 directly interacts
with ATGL, whereat the hydrophobic region of G0S2 as well as the patatin domain of ATGL are crucial
structural features [91, 92]. Similarly to CGI-58, the biochemical mechanism behind G0S2-depedent
inhibition of ATGL is unknown.
Both ATGL and HSL together are responsible for more than 90% of lipolytic (TAG-hydrolase) activity
in cultured adipocytes and WAT [85]. So, ATGL and HSL, together with monoglyceride lipase (MGL),
which was identified as potent MAG hydrolase [93, 94], represent the three major lipases responsible
for adipocyte lipolysis (Fig. 6).
FIGURE 6. Stereo/regioselectivity of enzymes involved in lipolysis. ATGL and ATGL/CGI-58 hydrolyze TAG with so far
unknown selectivity. In the second step HSL degrades DAG specifically at the sn-3 position. The resulting mixture of MAG
isoforms is further degraded to glycerol by MGL, which exhibits no selectivity. ATGL, adipose triglyceride hydrolase; CGI-58,
comparative gene identification-58; DAG, diacylglycerol; FA, fatty acid; G, glycerol; HSL, hormone-sensitive lipase; MAG,
monoacylglycerol; MGL, monoglyceride lipase; TAG, triacylglycerol.
Introduction
12
Phospholipase C
Phospholipases specifically hydrolyze PLs at different chemical position. The four major classes of
phospholipases are distinguished by the type of catalyzed reaction. Phospholipase A1 (PLA1) and A2
(PLA2) catalyze the hydrolytic cleavage of the acyl chains at respective sn-1 or sn-2 position. In
contrast, phospholipase C (PLC) and D (PLD) cleave phospho-glycerol and phospho-headgroup esters.
Hence, only PLCs contribute to the intracellular formation of DAG. So far, thirteen PLC isozymes were
identified and assigned to six subclasses, namely β (1-4), γ (1-2), δ (1,3,4), ε, ζ, and η (1,2) [95-98].
Virtually all PLC isozymes are highly expressed in different brain regions and only a few (β3, δ1, δ3,
δ4, ε) are distributed to other, peripheral tissues, like liver, SM, or CM [99]. Intracellularly, the
soluble PLC proteins are localized mainly in the cytoplasm. Upon cell activation PLCs translocate to
the plasma membrane and develop catalytic activity [99]. The PLC dependent hydrolysis of
phosphatidylinositol 4,5-bisphosphate (PIP2), a plasma membrane-associated PL, describes a key
event during regulation of a variety of cellular functions. By producing two intracellular messengers,
namely sn-1,2 DAG and inositol 1,4,5-triphosphate (IP3), this reaction mediates activation of protein
kinase C (PKC) as well as intracellular Ca2+ release, respectively [99]. Additionally, PIP2, which usually
exhibits an arachidonic acid (C20:4) at sn-2 position, represents the precursor for 2-
arachidonoylglycerol (2-AG) that is strongly involved in endocannabinoid signaling [100, 101]. Due to
the strict sn-3 position of the phosphate residue, PLC generates exclusively sn-1,2 DAG.
B) De novo synthesis of DAG
In addition to the catabolic formation of DAG, two other pathways contribute to anabolic generation
of DAG (see Fig. 4). In those pathways DAG arises as intermediates of the de novo biosynthesis of
TAG and PLs (Fig. 7). The glycerol-3-phosphate (G3P) pathway is one major pathway of TAG and PL
synthesis and takes place in most tissues, predominantly in liver and WAT. The G3P-pathway begins
with the consecutive acylation of G3P by acyl-CoA dependent glycerol-3-phosphate acyltransferase
(GPAT) and acyl-CoA acylglycerol-3-phosphate acyltransferase (AGPAT, LPAAT). The product of this
reaction is phosphatidic acid (PA), which is dephosphorylated to DAG by PA phosphatase (PAPase,
lipins) [102-104]. In the so called MAG-pathway, MAGs are esterified to DAG by monoacylglycerol-
acyltransferase (MGAT). This pathway plays a predominant role in enterocytes upon feeding and is
also involved in the storage of TAG in adipocytes [105, 106].
Introduction
13
FIGURE 7. De novo synthesis of DAG. DAG can be formed by either G3P or MAG pathway. AGPAT, acyl-CoA acylglycerol-3-
phosphate acyltransferase; DAG, diacylglycerol; G3P, glycerol-3-phosphate; GPAT, glycerol-3-phosphate acyltransferase;
LPA, lysophosphatidic acid; MAG, monoacylglycerol; MGAT, monoacylglycerol-acyltransferase; PA, phosphatidic acid;
PAPase, phosphaditic acid phosphatase.
The G3P pathway
It was recognized quite early that the liver exhibits enzymatic activities to generated PA from glycerol
and that DAG acts as precursor for both phosphatidylcholine (PC) and TAG. In the 1950´s, Kennedy
and coworkers described an enzymatic activity that dephosphorylates PA to form DAG in vitro [107].
This finding completed the enzymatic sequence of TAG and PL synthesis from glycerol. This pathway
is now known as Kennedy-pathway [108, 109].
In mammals, PA dephosphorylation is catalyzed by three recently identified members of the Lipin
family, namely Lipin1, 2, and 3. Among these, Lipin1 is the most extensively studied. It is highly
expressed in tissues with high rates of lipid flux, like CM, WAT, and SM [110, 111]. In WAT and SM,
Lipin1 was identified to account for the entire PAP activity, whereas the other two members
contribute essentially to total PA dephosphorylation of liver, brain and placenta [111, 112].
Furthermore, early studies revealed that lipins locate within the cytoplasm and translocate rapidly to
ER-membranes upon elevated levels of intracellular FAs [113]. Loss-of-function mutations within
Lipin1 cause dramatic metabolic impairments, like hypertriglyceridemia and severe hepatic steatosis,
as observed in fatty liver dystrophic mice [110]. The opposite effect is observed in transgenic mice
overexpressing Lipin1 in adipocytes. These mice exhibit increased amounts of TAG, which fits to
Lipin1 dependent generation of DAG as precursor for TAG [114]. Since G3P and PA are
phosphorylated at the sn-3 position of the glycerol backbone, their dephosphorylation generates
exclusively sn-1,2 DAG.
Introduction
14
The MAG-pathway
The esterification of MAG catalyzed by MGAT enzymes forms the first step in TAG synthesis following
dietary absorption by enterocytes. So far, three enzymes are known to possess MGAT activity,
MGAT1, MGAT2, and MGAT3. All three isoforms are located at the ER [115-119]. Besides similar
intracellular localization, MGAT isoforms differ in their tissue-specific expression pattern as well as in
their specific catalytic activity. In contrast to MGAT1, which is mainly expressed in stomach, AT and
kidney, MGAT2 and MGAT3 exhibit highest expression in small intestine [115-117, 119, 120]. MGAT3,
which is found exclusively in higher mammals, exhibits significantly higher specific DAG-
acyltransferase activity as compared to MGAT1 and MGAT2. This indicates that MGAT3 functions as
TAG synthase [121]. Furthermore, MGAT3 prefers sn-2 MAG as acyl-acceptor and activated palmitic
acid (C16:0) or C18:1 as acyl-donor [117]. Thus, the generated DAGs exhibit either sn-1,2 or sn-2,3
isoform. The prior mentioned lipid intermediates, sn-2 MAG, C16:0 and C18:1, are the major
hydrolytic products of the pancreatic lipase (PAL) dependent TAG hydrolysis during intestinal
digestion [122-124]. Hence, MGAT3 is supposed to be mainly involved in the re-esterification of
dietary absorbed fat within the small intestine.
MGAT2 was identified to possess little or no selectivity regarding chain-length or saturation-level of
FA-CoAs [115]. Furthermore, incubation of MGAT2 with rac-1/3 MAG results in the generation of sn-
1,3 and rac-1,2/2,3 DAG implicating that all position of the glycerol backbone can be esterified by
MGAT2 [115]. In contrast, MGAT1 displays a stricter selectivity for glycerol position as well as for
utilized FA-CoA species. MGAT1 shows highest activity when incubated with long-chain unsaturated
FAs, with the utmost with arachidonic acid (C20:4) [116]. Incubation with either sn-1 or sn-3 MAG
yields in the generation of sn-1,3 DAG suggesting that MGAT1 preferentially esterifies sn-1 or sn-3
position [116]. In line with these findings, incubation of sn-2 MAG results exclusively in the
generation of the rac-1,2/2,3 DAG [116].
In summary, intracellular reactions, which contribute to DAG formation, can generate all DAG
isoforms (Fig. 4). While the stereo/regioselectivity of most TAG lipases is presently unknown, PLC
isozymes generate exclusively sn-1,2 DAG. Additionally, sn-1,2 DAG is also formed during de novo
synthesis via the G3P-pathway. In contrast, DAGs generated by different MGAT enzymes can exhibit
other isomerism, like sn-1,3, depending on the catalyzing MGAT enzyme. The potential significance
of different DAG isoforms in regard to further utilization is addressed within the next sections.
Introduction
15
Utilization of DAG
In general, not only DAG generating enzymes, but also enzymes, which are involved in the utilization
of DAG might exhibit selectivity for specific DAG isoforms (Fig. 8).
FIGURE 8. Several enzyme classes are potentially involved in the utilization of different DAG isoforms. Different DAG
isoforms are assumable substrates for several enzyme classes, including transferases, kinases, and lipases.
Thus, isomerism of DAG species could influence (i) their degradation by lipases which leads to MAG
formation, (ii) their re-esterification to TAG by DAG-specific acyltransferases, (iii) their conversion to
PC by CDP-choline:1,2-diacylglycerol cholinephosphotransferases (CPTs), and (iv) their
phosphorylation to PA by diacylglycerol kinases (DGKs) (Fig. 9). So far, little is known about the
potential impact of DAG isomerism on the activity of the consuming enzymes.
FIGURE 9. Different enzyme classes utilize DAG. DAG is a potential substrat for lipase, kinase and acyltransferase reactions.
CPT, CDP-choline:1,2-diacylglycerol cholinephosphotransferases, DAG, diacylglycerol; DAGL, DAG lipase; DGAT, DAG-O-
acyltransferase; DGK, DAG kinase; MAG, monoacylglycerol; PA, phosphatidic acid; PL, phospholipid; TAG, triacylglycerol.
Introduction
16
DAG acyltransferases
The acylation of DAG is the final step of TAG synthesis. The esterification reaction consumes DAG and
FA-CoA and is catalyzed by diacylglycerol-O-acyltransferases (DGATs). So far, two mammalian DGAT
enzymes have been identified, DGAT1 and DGAT2.
DGAT1 was identified due to high sequence similarity with acyl-CoA:cholesterol-acyltransferase
(ACAT) enzymes. DGAT1 belongs to the large family of membrane-bound O-acyltransferases
(MBOAT) whose members catalyze the transfer of FAs onto thiol or hydroxyl groups of either lipid or
protein acceptors [125]. DGAT1 is highly expressed in small intestine, AT, SM, CM, skin, spleen, and
testis, where it localizes strictly to ER-membranes [126]. DGAT1 contains three transmembrane-
spanning domains and an active site within the COOH-terminal region facing the ER lumen [127]. The
NH2-terminal region, located in the cytoplasm, allows formation of tetramers and binds long-chain
FA-CoAs [128], but is not required for acyl transfer [129]. Whether the active site of DGAT1 localizes
lumenal (latent activity) is conrtoversial. Yet, several studies found mild, latent activity [130-132].
DGAT1 can catalyze a diversity of different acyltransferase reactions including MGAT, monoester wax
synthase, and retinol acyltransferase [133, 134].
DGAT2 shares little similarity with DGAT1. It belongs to a seven-member family including former
mentioned MGAT1, 2, and 3 [118, 135]. All members of this family contain the highly conserved
amino acid sequence HPHG, which in case of DGAT2 is supposed to be part of the active site [118,
125, 136]. Additionally, DGAT2 contains a FLXLXXXn consensus sequence, which displays a neutral
lipid binding domain, found in other neutral lipid metabolizing proteins, like plasma cholesteryl ester
transfer protein, HSL or Triacylglycerol hydrolase/Carboxylesterase 3 (TGH/Ces3) [137, 138]. Within
DGAT2 this domain is responsible for DAG binding, and mutations in this region markedly reduce in
vitro acyltransferase activity [136]. Expression of DGAT2 mRNA is highest in liver, AT, mammary
gland, testis, peripheral leukocytes, and CM [126]. In cultured cells DGAT2 localizes to the ER under
basal conditions. Upon supplementation of FAs and resulting induction of TAG-synthesis, DGAT2
partly localizes to mitochondria-associated membranes (ER domains tightly interacting with
mitochondria) and to LDs [139]. Unlike DGAT1, DGAT2 contains two ER-membrane spanning domains
and both the COOH- as well as the NH2-terminal domain face the cytoplasm [118], suggesting a
spatial difference of DGAT1 and DGAT2-dependent TAG synthesis. Furthermore, DGAT2 does not
possess activity towards substrates other than DAG.
Both DGAT enzymes have been shown in functional studies to be involved in intracellular TAG-
synthesis. Overexpression of either DGAT1 or DGAT2 in mammalian cells causes an increase of in
vitro DGAT activity, whereat specific activity of DGAT1 is significantly higher as compared to that of
Introduction
17
DGAT2 [140]. In contrast, cells expressing DGAT2 show higher TAG mass, bigger LDs, as well as
enhanced glycerol incorporation as compared to cells expressing DGAT1 [140], indicating differences
between in vitro activity and their importance in vivo. DGAT2 shows no preferences regarding
saturation level of FA-CoAs but prefers medium-chained FA-CoAs (C12:0) and short- and medium-
chained DAG species (e.g. C6:0, C8:0, C12:0) as substrate [118, 135]. In contrast, DGAT1 prefers
monounsaturated acyl-donors, like C18:1-CoA, as compared to saturated FA-CoAs, like C16:0-CoA
[118]. Noteworthy, studies on human DGAT1 of the small intestinal revealed equal preference for
both C16:0 and C18:1 [141]. An additional difference is the sensitivity of both enzymes against
magnesium, which was demonstrated by in vitro experiments. High concentrations (> 50mM) are
described to suppress DGAT2 activity whereas DGAT1 activity is much less affected [118].
The differences in subcellular localization as well as in enzymatic activity of DGATs suggest different
intracellular functions. This hypothesis is supported by the phenotypes of DGAT-deficient mice
(DGAT1ko and DGAT2ko). Interestingly, the enzymes cannot functionally compensate for each other
[140, 142]. DGAT1ko mice exhibit a moderate phenotype characterized by reduced TAG levels in
several tissues (e.g. WAT, SM, liver) and resistance against diet-induced obesity [142]. In contrast,
DGAT2ko mice die within the first days of life and suffer from a major skin dysfunction and severe
lipopenia [140]. The exact reasons for these drastic differences are unknown. Recently, distinct
functions of DGAT1 and DGAT2 in hepatic TAG synthesis were reported. DGAT2 is described to
mediate esterification of newly synthesized FAs, whereas DGAT1 catalyzes the synthesis via
utilization of exogenously supplied FAs [143, 144]. Whether this finding holds true for other tissues
needs to be tested.
The regio/stereospecificity of DGAT enzymes is currently unknown. Conclusive studies regarding this
topic have not been performed. Since neutral lipid metabolism leads to the formation of all DAG
isoforms it is conceivable that DAG isomerism influences the activity of DGAT enzymes. A diverging
stereo/regioselectivity of DGAT enzymes might partially explain the differences observed in DGAT1ko
and DGAT2ko mice.
DAG lipases
In addition to HSL, several other cellular lipases possess hydrolytic selectivity for DAG, generating
MAG and free FAs. In humans, two additional sn-1 specific DAG lipases were identified, named DAG
lipase α and β (DAGLα and β) [145]. Both enzymes share ~30% homology and comprise four
predicted transmembrane-spanning domains as well as a serine lipase motif [145]. DAGLα is highly
expressed in brain and pancreas and to a lower extend in AT. DAGLβ shows highest expression in
Introduction
18
bone marrow and the liver [145, 146]. Enzymatic characterization revealed that both are specific
DAG lipases and exhibit a 3 to 8-fold higher selectivity for the sn-1 over the sn-2 position of DAG
[145]. Additionally, DAGLβ prefers DAG species, containing linoleic acid (C18:2) > C18:1 > C20:4 >
stearic acid (C18:0) at sn-2 position, whereas DAGLα shows equal activity against all examined DAG
species [145]. Both lipases strictly localize to the plasma membrane where sn-1,2 DAG is generated
during PLC-dependent hydrolysis of PIP2. DAGL-mediated breakdown of sn-1,2 DAG results in 2-AG,
which is the most abundant endocannabinoid in tissues and acts as ligand for cannabinoid-receptors
(CB1, CB2) [100, 101]. The role of DAGLα and DAGLβ in the biosynthesis of the endocannabionid 2-
AG was established from the phenotype of mice, either deficient in DAGLα or DAGLβ (DAGLαko,
DAGLβko) [146]. In line with the tissue expression pattern, DAGLαko mice display an 80% reduction
in 2-AG levels in brain, whereas DAGLβko mice show 90% reductions in 2-AG levels in liver [146]. To
date it is unknown if plasma membrane-bound, sn-1 specific DAGL enzymes are additionally involved
in the degradation of DAGs, which derive from lipolysis of cytoplasmic TAG stores.
DAG kinases
DGKs catalyze the formation of PA by phosphorylating the free hydroxyl (-OH) group of DAG.
Together with PAPases/Lipins, DGKs are crucially involved in the maintenance of intracellular DAG
and PA levels. So far, ten DGKs isozymes have been identified in mammals [147, 148]. All mammalian
DGK isozymes share two common structural features, the cysteine-rich C1 domain, which is
responsible for DAG binding and potentially involved in protein-protein interaction and a catalytic
domain, responsible for enzymatic activity [149]. Almost every tissue expresses at least one member
of the DGK family. Moreover, numerous tissues express several different DGK isozymes, e.g. all ten
DGKs can be found in brain extracts [150]. DGKs localize to multiple cellular compartments, including
the nucleus, plasma membranes, the cytoskeleton, the golgi apparatus, and the ER [151]. Little is
known about the specific functions of different DGK enzymes with regard to their organelle
distribution. Whereas some isoforms translocate to the plasma membrane upon activating stimuli
(e.g. DGKδ1 upon exposure to phorbol esters, most likely due to attenuate DAG concentration raised
by PLCs), others stay located inside the nucleus potentially regulating nuclear DAG level (e.g. DGKα,
DGKζ) [149].
DGKs act sn-1,2 selective, since their C1 domain shares high homology to sn-1,2 DAG specific binding
motifs C1A and C1B of PKCs [152-154]. Up to now it is questionable if DGK isoforms exhibit activity to
also phosphorylate sn-1,3 or sn-2,3 DAG species. Some more information about DAG selectivity is
available for particular DGKs, like DGKε. DGKε was identified as smallest DGK isoform, strictly located
Introduction
19
at plasma- or ER-membranes [155]. Furthermore, DGKε exhibits specificity for DAG containing C20:4
at sn-2 position [156], which is the product of PIP2 hydrolysis by PLC [157].
DAG choline/ethanolamine phosphotransferases
All tissues and cell types can synthesize PC via the CDP-choline or the “Kennedy” pathway [108, 109].
In an analogue manner, phosphatidylethanolamine (PE) can be formed via the CDP-ethanolamine
pathway. The final step of both pathways, namely the direct conversion of DAG to either PC or PE is
catalyzed by CPT or CDP-ethanolamine:1,2-diacylglycerol ethanolaminephosphotransferase (EPT),
respectively.
In humans, two proteins exhibiting CPT activity have been identified. CPT, which acts as CDP-choline
specific enzyme and CEPT which can utilize CDP-choline as well as CDP-ethanolamine [158, 159].
Both CPT and CEPT proteins are integral membrane proteins and localize at the golgi apparatus and
the ER, respectively [160]. CPT expression is most abundant in colon, intestine, CM, and spleen,
whereas CEPT is expressed in all tissues examined [159]. Due to a cellular excess of CPT activity, both
enzymes are not supposed to be rate-limiting in PC synthesis [161]. Accordingly, overexpression of
either CPT or CEPT does not lead to elevated PC synthesis [162, 163]. Since no purified mammalian
CPT enzyme is available, information about substrate specificity originates from structure/function
analysis of CPT1 of S. cerevisiae. This protein prefers sn-1,2 DAG, more precisely dipalmitolein
(2xC16:1) > diC16:0 = C18:1/C16:0 > C16:0/C18:1 DAG as substrates in vitro [164]. Whether the
mammalian ortholouges exhibit similar stereospecificity needs to be shown.
EPT1 encodes for an enzyme, which in contrast to CPT utilizes CDP-ethanolamine as substrate,
thereby generating PE [165]. Little is known about expression or biochemical properties of EPT1.
Notably, incubation of hepatocytes with radiolabeled ethanolamine results in a highly specific
accumulation of radioactivity in C16:0/C22:6 PE in vivo indicating that EPT activity prefers
C16:0/C22:6 DAG as substrate [166].
Taken together, CPT, CEPT, and EPT are involved in the synthesis of PLs, thereby consuming
specifically sn-1,2 DAG.
Introduction
20
DAG and DAG-derived signals
Most of the reactions afore described either consume or generate FAs and DAGs. DAGs and FAs itself
as well as DAG- and FA-derived lipid species, like MAGs, FA-CoAs and ceramides are bioactive lipid
species, which act as second messengers within different signaling pathways (Fig. 10). Upon
dysregulation of the generation or utilization of DAGs and FAs, such lipid intermediates may
adversely affect cellular signaling.
FIGURE 10. DAG and DAG-derived lipid species are involved in various signaling pathways. Sn-1,2 DAG and ceramides can
activate novel/conventional and atypical PKC isoforms, respectively. FA and FA-CoA are involved in the activation of
different PPARs. The MAG species 2-AG binds to CBRs and activates endocannabinoid signaling. CBR, cannabinoid receptor;
DAG, diacylglycerol; FA, fatty acid; FA-CoA, FA-coenzyme A; MAG, monoacylglycerol; PKC, proteinkinase C; PPAR,
peroxisome proliferator-activated receptor.
A landmark study by Randle and co-workers [167] postulated the negative effects of FAs on insulin-
stimulated glucose oxidation in muscle. According to Randle´s theory, excessive uptake of FAs in the
muscle and concomitant increase of fat oxidation leads to a combined inhibition of glycolytic key
enzymes including pyruvate dehydrogenase, phosphofructokinase, and hexokinase. The ensuing
intracellular accumulation of glucose-6-phosphate and glucose inhibits further glucose uptake [167].
This interpretation was partially challenged when several studies revealed that increased FA levels in
the circulation were associated with a defect in glucose transport, provoked by an impaired insulin-
mediated translocation of glucose transporter 4 (GLUT4) rather than a defect in glycolysis [168, 169].
In addition, elevated levels of plasma FAs can result in an ectopic accumulation of lipids, like TAG,
DAG, and FAs. Furthermore, such lipid accumulations can lead to an altered insulin response of
insulin-sensitive tissues (e.g. muscle, liver).
Introduction
21
In muscle cells and adipocytes insulin activates the plasma membrane-bound insulin receptor, a
tyrosine kinase, which further phosphorylates the intracellular family of insulin receptor substrates
(IRS 1-4) on several tyrosine residues [170, 171]. Activated IRSs serve as docking station for proteins,
like phosphatidylinositide-3-kinase (PI3K) and downstream effectors, like protein kinase B (PKB/Akt)
[172, 173]. The subsequent generation of phosphatidylinositol-3,4,5-triphosphate (PIP3) by PI3K
facilitates the recruitment of PKB/Akt to the plasma membrane where it is phosphorylated by 3-
phosphoinositide-dependent protein kinase 1 (PDK1) [171]. This event enables further signal
transmission, which results in the release of GLUT4 to the plasma membrane [174]. Membrane-
associated GLUT4 facilitates glucose uptake. Additionally, activation of PKB/Akt in muscle and liver
cells leads to phosphorylation and inhibition of glycogen synthase kinase 3 [172, 175]. This inhibition
promotes glycogen synthesis and inhibits gluconeogenesis (Fig. 11). Defects within this signaling
cascade result in a loss of insulin-sensitivity and can lead to insulin resistance of affected
cells/tissues.
FIGURE 11. Intracellular pathway of insulin signaling. Insulin binds to insulin receptor that further activates IRS. This leads
to the activation of PI3K and furthermore to phosphorylation/activation of PKB/Akt by PDK1. Consequently GLUT4
translocate to the plasma membrane and enables glucose uptake in SM and adipocytes. Furthermore, activation of PKB/Akt
promotes glycogen synthesis in SM and liver. Additionally, insulin promotes lipogenesis and inhibits gluconeogenesis in
liver. GLUT4, glucose transporter 4; IRS, insulin receptor substrate; PDK1, 3-phosphoinositide dependent protein kinase 1;
PI3K, phosphoinositide-3-kinase; PIP2, phosphatidylinositol 4,5-bisphosphate; PIP3, phosphatidylinositol 3,4,5-triphosphate;
PKB/Akt, protein kinase B.
Introduction
22
DAG signal
The intracellular accumulation of DAG is assumed to be tightly connected to an altered insulin
response. More recently, ectopic DAG accumulation was found to be associated with disturbed
insulin signaling. These DAG effects are thought to derive from the action of the PKC-family members
which are known to play a crucial role in many signaling events, cellular differentiation, and cell
growth [176]. The PKC-family comprises three different subgroups, namely conventional (α, β1, β2
and γ; cPKC), novel (δ, ε, η and θ; nPKC), and atypical (ζ and λ/ι; aPKC) PKCs [176]. cPKS and nPKCs
display lipid-sensitive isoforms and are usually activated by PLC-dependent generation of sn-1,2 DAG,
either Ca2+-dependent (cPKC) or -independent (nPKC). In contrast, aPKC are mainly activated by
protein/protein interactions and are insensitivity towards DAG or Ca2+ [176]. These differences are
attributed to regulatory domains, designated as C1 and C2, which account for lipid binding and Ca2+-
sensing, respectively. The C1 domain of both conventional and novel PKCs binds DAG and phorbol
esters, whereas C1 domain of aPKCs binds PIP3 and ceramides. Only conventional isoforms contain a
functional C2 domain which binds anionic PLs in a Ca2+-dependent manner [176]. The activity of
cPKCs and nPKCs is highly influenced by intracellular levels of DAG. Noteworthy, earlier studies
showed that only the sn-1,2 DAG isoform has the ability to activate PKCs. The other isoforms, sn-1,3
and sn-2,3 are inactive [152-154].
Of all PKC subgroups mainly nPKC, more precisely PKCε and PKCθ, adversely affect insulin signaling
[177]. Earlier studies showed that PKCs are activated in diabetic rodent models [178, 179] and that
activation of PKCs by phorbol esters cause IR [180-182]. In rodents, an infusion of intralipid/heparin
leads to enhanced plasma FA concentrations causing impaired insulin signaling associated with
activation of PKCθ in SM [183]. The development of IR was thereby associated with an increase in
intramuscular DAG levels and independent of TAG or ceramide content [184]. In accordance, PKCθ-
deficient mice are protected from acute SM-IR after lipid infusion [185]. Additionally, PKCε is also
implicated in the development of hepatic IR. In rodents, 3 days of high-fat feeding cause hepatic
steatosis and hepatic IR without peripheral lipid accumulation or IR. The requirement of PKCε in this
process is evident by enhanced hepatic insulin response in PKCε-antisense oligonucleotide (ASO)
treated fat-fed rats [186]. Consistent with this, mice lacking PKCε exhibit slightly increased hepatic
lipid content but are resident against diet-induced IR following 1 week of high-fat feeding [187].
Up to now several mechanisms have been identified by which nPKCs impair insulin action. Recent
studies showed that PKCθ can phosphorylate IRS1 at Ser1101 which blocks insulin stimulated tyrosine
phosphorylation [188] and activation of PI3K [177]. In rat liver, PKCε and insulin receptor reside in
close proximity and inhibition of PKCε expression protects against HFD-induced reduction of insulin
receptor kinase activation [186] (Fig. 12).
Introduction
23
FIGURE 12. Inactivation of insulin signaling by DAG via novel PKC isoforms. sn-1,2 DAG activates novel PKC isoforms,
PKCε/θ, which leads to phosphorylation of IRS at serine residues. This event inhibits downstream effector signaling. GLUT4,
glucose transporter 4; IRS, insulin receptor substrate; PDK1, 3-phosphoinositide dependent protein kinase 1; PI3K,
phosphoinositide-3-kinase; PIP2, phosphatidylinositol 4,5-bisphosphate; PKB/Akt, protein kinase B; PKC, protein kinase C.
Phenotypes of a variety of genetic murine models lead to the hypothesis that enhanced DAG levels in
insulin responsive tissue ultimately result in an impaired insulin signaling. Mice lacking mitochondrial
GPAT accumulate FA-CoA, but not TAG or DAG when set on a high-fat diet [189]. Besides this
elevation of FA-CoA, these mice are protected from diet-induced hepatic IR. In contrast,
overexpression of mitochondrial GPAT does not change FA-CoA levels but leads to hepatic IR, which
is associated with increased levels of lysophosphatidic acid (LPA), DAG, and TAG [190]. Similarly, in
obese Zucker rats (non-functional leptin receptor) IR is associated with increased amounts of hepatic
and muscle ceramide and DAG contents [191]. Another model arguing for DAG as mediator of IR are
mice fed high-ketogenic diet. These mice develop severe hepatic steatosis and profound hepatic IR
which is associated with a 350% increase in hepatic DAG content [192]. This increase is followed by
activation of PKCε and decreased insulin-mediated tyrosine-phosphorylation of IRS2. In mice
overexpressing DGAT2 in liver, hepatic TAG as well as DAG and ceramide levels are markedly
increased [193, 194]. These mice were first reported to exhibit normal hepatic insulin sensitivity
[193] but were recently identified to exhibit enhanced PKCε activation accompanied by severe
hepatic IR [194]. Furthermore, the key interaction between DAG, PKCε activation and hepatic IR is
confirmed in numerous other rodent models [195]. Interestingly, a recent study discovered that
hepatic DAG content of cytoplasmic LDs is the best predictor of IR in obese, non-diabetic individuals
[196]. In the same study they observed distinct localization of PKCε at cytoplasmic LDs as well as
Introduction
24
enhanced activation of this PKC isoform in these patients [196]. So far, this study is unique in
connecting LD turnover with PKC-signaling. However, the question concerning the intracellular
localization as well as the primary metabolic source of DAGs, which are involved in PKC-signaling is
still under dispute.
A number of murine model argue against a causative role of DAG accumulation in the development
of IR. For example, HSLko animals accumulate large amounts of DAG in adipose and non-adipose
tissues [56, 59] but do not develop pronounced alterations in insulin signaling [57, 197-199].
Although, depending on genetic background, some HSLko strains show signs of impaired insulin
signaling [199, 200], others do not or even show increased insulin sensitivity [197, 198]. ASO-
dependent decrease of CGI-58 expression, the co-activator of ATGL, leads to a markedly increase of
hepatic TAG, DAG, and ceramide content in fat-fed mice [201]. However, CGI-58 knockdown in mice
is associated with improved insulin sensitivity and glucose tolerance [201] suggesting additional
mechanism and factors to be involved in insulin signaling. So far, little evidence points towards a
direct association between ATGL-dependent lipolysis and altered insulin signaling. ATGLko mice
exhibit markedly increased ectopic lipid levels, mainly TAG, but display improved glucose tolerance
[75]. A recent study observed that overexpression of ATGL in cultured myotubes leads to an
increased content of both DAG and ceramides, which is associated with impaired insulin signaling
[202]. Nevertheless, mice overexpressing ATGL in adipose tissue appear to be protected from IR
[203]. Thus, to date it is unclear whether ATGL- derived DAGs activate PKC and may thereby interfere
with insulin signaling.
In summary, above mentioned studies suggest a link between DAG, PKC-signaling, and the
development of IR. In this context, DAG isomerism as well as intracellular DAG compartmentation
might play a crucial role. As outlined above, a number of metabolic reactions degrade or consume
DAG. Yet, they are located at different cellular compartments and the resulting DAG isoforms are
mostly unknown. Earlier studies clearly demonstrated that DAG-binding of the C1 domain of PKCs is
highly specific for sn-1,2 DAG. Other isoforms, like sn-2,3 or sn-1,3 DAG, completely fail to activate
PKCs [152-154] and hence display no signaling properties. Due to the inability of DAG isoforms, other
than sn-1,2, to activate PKCs it is crucial to determine the DAG isoforms which are generated and
further utilized during different metabolic reactions. The predominant reactions, accounting for DAG
generation and utilization in adipocytes, are catalyzed by ATGL/HSL as well as DGAT1/DGAT2,
respectively. Hence, the identification of the stereo/regioselectivity of these lipases and
acyltransferases would markedly augment the understanding of connections between DAG
metabolism and signaling and my provide explanations to previously unclear data regarding this
topic.
Introduction
25
FA and ceramide signal
Besides mentioned FA-induced alterations of glucose uptake, FAs can influence a variety of other
cellular processes. Since TAG displays the major storage form of FAs, TAG metabolism has a large
influence on the intracellular amount and composition of FAs and also FA-CoA. Both FAs and FA-CoAs
can interfere with cellular signaling pathways via activation of peroxisome proliferator-activated
receptors (PPARs). PPARs are nuclear hormone receptors involved in the regulation of genes
associated with inflammation, energy homeostasis as well as lipid, and lipoprotein metabolism. The
PPAR family is composed of PPARα, PPARδ, and three isoforms of PPARγ [204]. Oxidative tissues, like
CM, SM, or liver express mainly PPARα. PPARα activates the expression of genes involved in FA
transport and oxidation as well as keto- and gluconeogenesis [205]. PPARδ, which is responsible for
the activation of genes involved in glucose and FA utilization displays ubiquitous expression [206].
PPARγ is mainly active in cells associated with lipid storage and act as inducer of genes involved in
lipogenesis [207]. Activation of all PPARs requires co-activation by PPARγ coactivator-1α or β (PGC1α
or PGC1β), which follows binding of lipid ligands and dimerization with retinoid X receptor [208-210].
Both PGCs are involved in the regulation of uptake, transport, and oxidation of energy substrates
[205, 206]. Hence, dysregulation of these genes leads to severe metabolic disorders. Currently the
specificity of PPAR activation, in respect to endogenous ligands, is not entirely clear. However, the
established hypothesis lists FA and FA-CoAs either as direct (ligand) or indirect (precursor for other
lipid ligands) PPAR activator. In this regard unsaturated but not saturated FAs exhibit high signaling
potential for PPARs [211]. In particular mono-unsaturated FAs (MUFAs), like C16:1 and C18:1, are
potent activators for PPARα, much less for PPARβ or PPARγ [211-213]. PUFAs, like C18:2 and linolenic
acid (C18:3) exhibit activation potential for all PPAR species [211, 212].
Recently, ATGLko and HSLko mice were investigated regarding cellular signaling as a consequence of
TAG hydrolysis. These studies revealed different effects of either ATGL or HSL on the expression
pattern of genes involved in oxidative metabolism [214]. Deficiency of ATGL or HSL leads to a
decreased expression of oxidative genes in BAT [214]. However, only ATGL deficiency attenuates
oxidative gene expression in other tissues, like CM and SM suggesting a role of ATGL-dependent
lipolysis in the transcriptional control of FA metabolism [214]. Compatible with this theory,
overexpression of ATGL in either adipocytes or hepatocytes leads to increased gene expression of
PPARα and δ and enhanced FA oxidation as well as to enhanced PPARα activity and further target
gene expression, respectively [203, 215]. Conversely, knockdown of either ATGL or its co-activator
CGI-58 in hepatocytes leads to a suppression of PPARα target gene expression in vivo [201, 216].
Additionally, a recent study uncovered the role of ATGL in PPAR activation in cardiomyocytes [217].
Therein, it was shown that ATGL deficiency leads to a drastic decrease of PGC1α and β and to a
Introduction
26
dysfunction of mitochondrial substrate oxidation and respiration within cardiomyocytes. The
resulting excessive lipid accumulation leads to cardiac insufficiency and a lethal cardiomyopathy.
These results indicate that ATGL-dependent TAG hydrolysis generates essential mediators involved in
the generation of PPAR ligands. Moreover, ATGLko mice treated with PPARα agonists display
completely restored mitochondrial function and are spared from premature death [217].
Besides PPAR activation, which can be triggered by virtually all FAs, other bioactive lipid species
require specific FAs. In this context sphingolipids, more precisely ceramides, have been shown to be
negatively involved in the regulation of insulin signaling [218-221]. Since the sphingolipid synthesis
requires C16:0 [222, 223], the synthetic pathway of sphingolipids depends on the availability of this
FA species [224, 225]. Similar as for DAG, also ceramide accumulation was observed in SM of insulin-
resistant animals [191, 226], lipid-infused humans [227], and patients suffering from T2DM [220].
Furthermore, several studies reported that an accumulation of ceramides, induced by excessive
C16:0, is accountable for the initiation of IR in lean tissues [228, 229]. Ceramides efficiently block the
translocation of PKB/Akt to plasma membranes [221], which is a key step in insulin signaling and
promotes GLUT4 translocation to the plasma membrane resulting in glucose uptake [230]. The
detrimental effect of ceramides on PKB/Akt translocation was observed in several cell types,
including muscle cells [226, 231, 232], and adipocytes [232].
FIGURE 13. Inactivation of insulin signaling by ceramides via atypical PKC isoforms. Ceramides activate PKCζ which in turn
phosphorylates PKB/Akt. This phosphorylation inhibits translocation of PKB/Akt and consequently prevents translocation of
GLUT4 to the plasma membrane. GLUT4, glucose transporter 4; IRS, insulin receptor substrate; PDK1, 3-phosphoinositide-
dependent protein kinase 1; PI3K, phosphoinositide-3-kinase; PIP2, phosphatidylinositol 4,5-bisphosphate; PIP3,
phosphatidylinositol 3,4,5-triphosphate; PKB/Akt, protein kinase B; PKC, protein kinase C.
Introduction
27
In some cells ceramides block PKB/Akt translocation by direct activation of phosphatases which are
responsible for the dephosphorylation of PKB/Akt [175]. However, recent studies demonstrated that
the atypical, ceramide-binding PKCζ [233] is involved in phosphorylation of PKB/Akt. This
phosphorylation prevents translocation of PKB/Akt [226, 231] (Fig. 13). Additionally, a repression of
ceramide synthesis, by inhibition of serine palmitoyltransferase results in diminished lipotoxicity,
enhanced glucose regulation, and improved insulin response in mice [175, 234]. Together, these data
indicate that excessive amounts of C16:0, which result in an accumulation of ceramides represents at
least one possible reason for the onset of IR.
In summary, FAs affect directly or indirectly a variety of signaling events. Since some of the observed
effects strictly depend on single FA species, the composition of FA species within the cell plays a
crucial role. Since TAG hydrolysis is highly involved in the homeostasis of FAs and ATGL is rate-
limiting in this process, the identification of FA species, which are specifically released by ATGL during
lipolysis, is of great interest. A clear picture of the FA species composition, which derives from ATGL-
mediated lipolysis, would consequently enlarge the understanding of the interplay between energy
metabolism and FA-associated cell signaling.
MAG signal
MAG derives from either TAG- or PL-breakdown as direct product of DAG hydrolysis. Specifically 2-AG
was found to be the most abundant, endogenous ligand of cannabinoid receptors (CBRs; CBR1, CBR2)
[100, 101]. 2-AG derives mainly from the hydrolysis of arachidonic acid-containing PLs through the
combined actions of PLC and DAGL [100, 101]. In the nervous system, 2-AG is released from
postsynaptic neurons and causes retrograde inhibition of the presynaptic neurotransmitter release
[235]. Following binding and activation of presynaptic CBR1, 2-AG is internalized and degraded by
MGL. Endogenous compounds activating CBRs are commonly summarized as endocannabinoids
(ECs). Together, EC-metabolizing enzymes, CBRs, and ECs form the endocannabinoid system (ECS)
[236], which is active in neurons and non-neuronal cells, including hepatocytes, adipocytes, and cells
of the immune system. The ECS regulates numerous neuronal processes, including emotional
behavior, cognition, and pain. Furthermore, it is also involved in the regulation of food intake, energy
balance and lipid turnover [235].
Physiological implications of the ECS have been demonstrated in mice and humans. Obese patients
exhibit decreased food intake, reduced lipogenesis, and increased energy expenditure after
treatement with the CBR1-antagonist rimonabant [237, 238]. The same effects are observable in
mice lacking CBR1. Conversely, treatment with CBR-agonists results in a hyperactive ECS, which
Introduction
28
causes a central orexogenic effect and a concomitant reduction of energy expenditure [239]. This
leads to an increased lipid deposition in peripheral tissues, including WAT and the liver. A hyperactive
ECS may be one reason of increased appetite (food intake) and enhanced lipid deposition in obese
patients [240-242].
Due to the effects of the ECS on the energy metabolism, it has been linked to the pathogenesis of
metabolic deseases, like obesity and T2DM. Whether ATGL/HSL-dependent LD lipolysis contributes to
the generation of arachidonic acid-containing MAG signals or HSL is involved in the degradation of 2-
AG at cellular membranes is not known.
Aim of the Thesis
Aim of the Thesis
30
ATGL is the rate-limiting enzyme in TAG degradation. Its activity has been shown to be crucial for
lipid and energy homeostasis but also for intracellular lipid signaling, in particular PPARα activation.
The first aim of this study was to elucidate the enzymatic characteristics of ATGL-dependent TAG
hydrolysis. The characterization includes ATGL´s substrate, FA-, and stereo/regioselectivity and
should clarify the preference of ATGL for FA species, the position of hydrolyzed FA ester within TAG
and, thus, the stereo/regiospecific conformation of produced DAG. Since ATGL belongs to a large
protein family, which in part also comprises of phospholipases, it appears feasible that ATGL might
also hydrolyze any of the FA ester bonds of LD-forming PLs.
The second aim of this study was to investigate possible stereo/regiochemical requirements of DAG-
utilizing enzymes, more precisely DGAT1, DGAT2, and HSL. Selectivities of these enzymes should
delineate which cellular reactions can directly utilize lipolysis-derived DAGs, the direct product of
ATGL-dependent TAG hydrolysis.
Results
Results modified after Eichmann et al. [243] are indicated (§).To describe the investigation of the stereo/regioselectivity of
ATGL consistently, results obtained during forgone diploma thesis [244] were recapitulated (indicated by §§
).
Results
32
I) ATGL selectivity
This section focuses on the substrate-, FA- and stereo/regioselectivity of ATGL-dependent TAG
hydrolysis.
A) Stereo/regioselectivity
TAG consists of three FAs which are esterified to the three carbon atoms of the glycerol backbone.
According to stereochemical rules the carbon atoms are stereospecifically numbered (sn) 1-3. The
selectivity of a lipase is denoted depending on the position of hydrolysis.
ATGL regioselectively hydrolyzes sn-2 FA esters of TAG generating sn-1,3 DAG
in vitro
To investigate if ATGL exhibits selectivity for the FA position on the glycerol backbone, TAG hydrolysis
experiments with subsequent analysis of generated DAG isoforms were performed. Therefore, Cos7-
cells expressing ATGL (Fig. 14A) were homogenized and cytosolic fractions were incubated with 14C-
glycerol-labeled triolein in the presence of HSL-specific inhibitor (76-0079) with or without purified
GST-tagged CGI-58. Since the regioisomeric forms of DAG, sn-1,2/2,3 and sn-1,3 DAG, can be
separated by thin layer chromatography (TLC), the generation of this species was investigated first.
Incubation of ATGL led to an almost exclusive generation of sn-1,3 DAG, over a time period of 60 min.
In contrast, ATGL co-activated by CGI-58 led, besides an increase in activity, to the generation of both
regioisomeric forms of DAG (Fig. 14B). Experiments were repeated with slight modifications;
prolonging incubation period as well as changing the radiolabeled TAG tracer to 3H-FA-labeled
triolein. In addition, experiments were performed for different time periods and DAG formation was
measured. Data obtained reconfirm that ATGL alone generates exclusively sn-1,3 DAG, whereas ATGL
co-activated by CGI-58 results in an overall increase in generated DAGs and in the generation of both
DAG regioisomers (Fig. 14C, D). Results indicate that ATGL cleaves TAG selectively at sn-2 position
and extends selectivity upon CGI-58 co-activation to sn-1/3 position.
Results
33
FIGURE 14. ATGL cleaves TAG at sn-2 position and expands selectivity to sn-1/3 upon co-activation by CGI-58. A,
Expression of his-tagged ATGL in Cos7-cells was assessed by immunoblotting. Cytosolic fractions of Cos7-cells expressing
ATGL were incubated with a HSL-specific inhibitor (76-0079) in the presence (B, D) or absence (B, C) of purified GST-tagged
CGI-58 with either 14
C-glycerol (B) or 3H-FA labeled (C, D) triolein emulsified with PC for either 40 (C, D), 60 (B) or 120 min
(C, D) at 37°C. Lipids were extracted, separated by TLC and radioactivity in DAG bands was determined by scintillation
counting. Data are normalized to LacZ and presented as means +/- S.D. and are representative for 2 independent
experiments. Statistical significance was determined applying Student´s unpaired t-test (***,p<0,001; **, p<0,01). n.s.…no
significance. (Fig. 14C,D)§,§§
.
ATGL co-activated by CGI-58 generates sn-1,3 and sn-2,3 DAG
To investigate if ATGL looses sn-2 selectivity upon CGI-58 co-activation, DAG species were analyzed
using chiral-phase HPLC. This method allows the discrimination between all DAG species namely sn-
1,3 DAG, sn-1,2 DAG and sn-2,3 DAG. Therefore, TAG-hydrolysis experiments were performed using
cytosolic fractions of Cos7-cells expressing ATGL in combination with purified GST-tagged CGI-58 in
the presence of HSL-specific inhibitor (76-0079). Samples were incubated with non-labeled triolein
for 60 min. Lipids were extracted and separated using TLC. Following isolation, DAGs were
0
100
200
300
400
500
600
ATGL ATGL/CGI-58
DA
G (
cpm
/µg
pro
tein
*h)
sn-1,3
sn-1,2/2,3
**
0
20
40
60
80
100
120
140
160
180
40 120
DA
G (
cpm
/µg
pro
tein
*h)
time [min]
ATGLsn-1,3
sn-1,2/2,3
n.s.
0
500
1000
1500
2000
2500
40 120
DA
G (
cpm
/µg
pro
tein
*h)
time [min]
ATGL/CGI-58sn-1,3
sn-1,2/2,3***
***
50 -
37 -
ATGL kDa
B
C D
A
Results
34
derivatized to their corresponding 3,5-dinitrophenylurethanes and separated by chiral-phase HPLC.
First, as “proof of principal”, a racemic diolein reference compound, which comprises all three DAG
species, was analyzed. The analysis proved the suitability of this method by separating all three
species into distinct peaks over a time range of 16 min (Fig. 15A). Next, as a positive control, “egg-
yolk”-lecithin was digested using phospholipase C (b. cereus). Breakdown of lecithin resulted in
exclusively sn-1,2 DAG (Fig. 15B). Subpeaks within the sn-1,2 DAG peak reflect the FA composition of
this biological sample and correspond to sn-1,2 DAGs comprising either C16:0, C18:1 or C18:2 in
different combinations. When triolein was hydrolyzed by ATGL co-activated by CGI-58 exclusively sn-
1,3 and sn-2,3 DAGs were generated (Fig. 15C). These data indicate a sn-2 selectivity of ATGL which is
enlarged to sn-1 position by CGI-58.
FIGURE 15. ATGL co-activated by CGI-58 generates sn-1,3 and sn-2,3 DAG. A-C, Chiral-phase HPLC resolution of different
DAG species. DAGs were analyzed as corresponding 3,5-dinitrophenylurethanes by chiral-phase HPLC. Analysis of a racemic
dioleoylglycerol reference mix (A), the reaction products of „egg yolk lecithin” hydrolyzed by purified PLC (b. cereus, the
three peaks at the retention time range 11.5-13min display the different FA composition of the resulting DAGs; I: 16:0-
18:1+18:1-18:1; II 16:0-18:2+18:1-18:2; III:18:2-18:2; B), and the reaction products of triolein hydrolyzed by ATGL contained
in cytosolic fraction of Cos7-cells, in the presence of purified GST-tagged CGI-58 and HSL-specific inhibitor (76-0079, C). Data
are representative for 2 independent experiments. x…unknown compound. (Fig. 15A,B,C)§,§§
.
A B
C
Results
35
Conversion of DAG substrates is not detectable
The next question was, whether sn-1,2 DAG is preferentially consumed in follow-up reactions
occuring in cellular cytosolic fractions, or if spontaneous occurring non-enzymatic transesterification
(racemization) could be a reason for the lack of sn-1,2 DAG. Hence a racemic mixture of diolein was
incubated with cytosolic fractions of Cos7-cells in the presence of HSL-specific inhibitor (76-0079) for
120 min. DAGs were isolated from extracted lipids and analyzed before and after incubation using
chiral-phase HPLC. No alterations in DAG species composition were observed over an incubation
period of 120 min (Fig. 16A). Furthermore, diolein isoforms were determined by TLC (Fig. 16B). Also
with this method, no evidence of transesterification was observed. This proves the stability of DAG
isoforms in this experimental setup.
FIGURE 16. DAG substrates show no transesterification or decomposition. A, A mixture of sn-1,3, sn-1,2 and sn-2,3 DAG or
either sn-1,3 (B) or rac-1,2/2,3 DAG (B) emulsified with PC was incubated with cytosolic fraction (A) or lysates (B) of Cos-7
cells in the presence of a HSL-specific inhibitor (76-0079) for 120 min at 37°C. Lipids were extracted before and after
incubation according to Folch et al, separated by TLC and DAGs were analyzed as corresponding 3,5-dinitrophenylurethanes
by chiral-phase HPLC (A) or visualized by iodine staining (B). Data are presented as means +/- S.D. and are representative for
2 independent experiments. (Fig. 16B)§.
HSLko mice show drastic accumulation of sn-1,3 DAG in WAT
To investigate, if the selectivity of ATGL is also present in an in vivo model, acylglycerol composition
of WAT from HSLko mice was examined. The deficiency of HSL is known to lead to a drastic
accumulation of DAGs, due to the fact that HSL represents the predominant intracellular DAG-lipase
in lipolysis. Lipid analysis revealed that total acylglycerol levels (including TAG, DAG, MAG) of HSLko
mice were unaltered compared to wt littermates (Fig. 17A). Further analysis revealed the expected
0
10
20
30
40
50
60
70
sn-1,3 sn-1,2 sn-2,3
DA
G is
om
ers
(%
of
tota
l)
timepoint: 0min
timepoint: 120min
A B
Results
36
substantial accumulation of DAG (~6-fold), which was accompanied by a moderate decrease in TAG
level (~10 %) (Fig. 17B).
FIGURE 17. Accumulation of sn-1,3 DAG in WAT of HSL-deficient mice. A, Lipids were extracted according to Folch et al.
and total acylglycerol levels in WAT of wt and HSLko mice were determined using Infinity-TAG kit. C, Neutral lipids of wt and
HSL-deficient WAT were separated by TLC using chloroform/acetone/acetic acid (90/8/1; v/v/v). B, D, Bands corresponding
to TAG and DAG were scraped off, extracted and acylglycerol content was determined using Infinity-TAG kit (B and D) and
DAG isomers were analyzed by chiral-phase HPLC (C, right half). Data are presented as means +/- S.D. Statistical significance
was determined applying Student´s unpaired t-test (***, p<0,001), n=4 (each genotype). (Fig. 17A,B,C,D)§.
Next, TLC analyses were performed and showed that sn-1,3 DAG was not detectable in lipid extracts
of wt mice. In contrast, sn-1,3 DAG constitutes a major DAG species in acylglycerols of HSLko mice
(Fig. 17C). The detailed species composition of isolated DAG from wt and HSLko WAT was analyzed
using chiral-phase HPLC. In wt WAT, acyglycerol content consisted of 1.7% of sn-1,2 DAG (80% of
total DAG). 0.06% of sn-1,3 (2% of total DAG) and 0.32% of sn-2,3 DAG (18% of total DAG) constitute
only a minor portion of total acylglycerol in wt mice (Fig. 17D left). Composition analysis confirmed
the drastic accumulation of sn-1,3 DAG in WAT of HSLko mice, which accounted for 7.7% (65% of
0
10
20
30
40
50
60
70
80
90
100
wildtype HSLko
acyl
glyc
ero
l (%
of
tota
l )
triacylglycerol
diacylglycerol97.9%
88.2%
2.1%
11.8%
0
50
100
150
200
250
wildtype HSLko
tota
l acy
lgly
cero
l (µ
mo
l/g
WA
T)
A B
C D
0
1
2
3
4
5
6
7
8
9
wildtype HSLko
DA
G (
% o
f to
tal a
cylg
lyce
rol)
sn-1,3 DAG
sn-1,2 DAG
sn-2,3 DAG
***
Results
37
total DAG) of total acylglycerols. Furthermore, it showed that sn-2,3 DAG was more abundant than
sn-1,2 DAG and accounted for 2.8% (24% of total DAG) of HSLko acylglycerols. The 1,3% of sn-1,2
DAG (11% of total DAG) were comparable to the sn-1,2 DAG content of wt mice (Fig. 17D right).
Together, data of this section clearly demonstrate that ATGL hydrolyzes TAG selectively at sn-2
position, yielding sn-1,3 DAG. Furthermore, co-activation of ATGL by CGI-58 leads to the concomitant
generation of sn-2,3 DAG but not sn-1,2 DAG (Fig. 18). The accumulation of sn-1,3 and sn-2,3 DAG in
HSLko mice corroborates the stereo/regioselectivity of ATGL in vivo.
FIGURE 18. Schematic depiction of the stereo/regioselectivity of ATGL. ATGL hydrolyzes TAG at sn-2 position, or at sn-1 or
sn-2 position when co-activated by CGI-58 yielding sn-1,3 DAG or sn-1,3 and sn-2,3 DAG, respectively. ATGL, adipose
triglyceride hydrolase; CGI-58, comparative gene identification-58; DAG, diacylglycerol; FA, fatty acid; TAG, triacylglycerol.
B) FA selectivity
TAGs occurring in biological systems are usually not uniformly esterifies with only one kind of FA but
instead contain different FAs with varying chain-length and saturation. Naturally abundant FAs
contain 8 to 22 carbon atoms and up to 6 double bonds. Annotation of FAs includes number of
carbon atoms followed by number of double bonds (saturation level) e.g. palmitic acid (C16:0, 16
carbon atoms:0 double bonds). Besides stereo/regioselectivity, lipases usually display substrate
selectivity against chain-length as well as saturation level of bound FA species.
This section focuses on the FA-selectivity of ATGL in vitro and in vivo.
Results
38
ATGL differentiates between chain-length of TAG-bound FAs in vitro
The next aim was to investigate, if ATGL exhibits selectivity for distinct FA species during TAG
hydrolysis. Therefore, murine ATGL and CGI-58 were expressed in Cos7-cells (Fig. 19A) and in vitro
TAG hydrolase assays were performed using various TAG species as substrate. In the first set of
experiments, the used TAG substrates were uniformly esterified with naturally abundant FA species,
namely palmitoleic acid (C16:1), C18:1, C18:2 and C18:3. The high melting points (>40°C) of TAGs
consisting of three saturated FAs (C12:0, C14:0, C16:0, C18:0) precluded the preparation of suitable
lipid emulsions. Furthermore, also liquid soluble TAG emulsions (unsaturated FAs) differ in their
solubility. To prevent any effects, caused by solubility, all substrates were diluted to the same final
concentration (0.25 mM). ATGL displayed highest activity against tripalmitolein (3xC16:1), followed
by triolein (3xC18:1), trilinolenin (3xC18:3) and trilinolein (3xC18:2) (Fig. 15B). ATGL co-activated by
CGI-58 showed an expected increase in hydrolase-activity. As a consequence differences in activities
against used substrates became more obvious (Fig. 19B). Results indicate that ATGL is capable of
hydrolyzing virtually all used TAG substrates with a marked preference for C16:1 esters.
FIGURE 19. ATGL hydrolyzes TAG with FAs of different chainlength in vitro. A, Recombinant proteins were expressed in
Cos7-cells and expression of his-tagged proteins was assessed by immunoblotting. B, Homogenates of Cos7-cells expressing
ATGL were incubated in the absence or presence of CGI-58 with different, homogeneously esterified TAG species emulsified
with PC as substrates for 1 h at 37°C. Released FAs were measured using NEFA-C kit. Data are normalized to LacZ and
presented as means +/- S.D. and are representative for 2 independent experiments. Statistical significance was determined
applying Student´s unpaired t-test (***, p<0,001; **, p<0,01). (Fig. 19A,B)§.
0
100
200
300
400
500
600
700
800
FA (
nm
ol/
mg
pro
tein
*h)
ATGL/LacZ
ATGL/CGI-58
***
***
*** ***
*****
150 -
100 -
75 -
50 -
LacZ
37 -
ATGL CGI-58 kDa kDa
A B
Results
39
ATGL cleaves saturated FAs of mixed-labeled TAG species in vitro
Next, the ability of ATGL to hydrolyze saturated FA esters within mixed TAG species was investigated.
First, ATGL-dependent hydrolysis of TAG containing C16:0 at the sn-1/3 or sn-2 and C18:1 at the
remaining positions was investigated. These TAGs reflect a substrate class mixed in chain-length as
well as saturation-level. Virtually all of these TAG species were hydrolyzed at the same rates by either
ATGL alone or ATGL co-activated by CGI-58 (Fig. 20A). A minor, yet significant decrease in the
hydrolytic activity of ATGL was found when C16:0 was esterified at sn-2 position, indicating that a
saturated FA at the preferred position of ATGL-dependent hydrolysis results in a slightly decreased
activity. Next, substrates composed of FAs exhibiting same chain-length (18 carbon atoms) but
different saturation-level were tested. ATGL alone as well as ATGL co-activated by CGI-58 showed
equal hydrolase activities against the different TAG substrates (Fig. 20B). The small decrease in
activity using TAG containing C18:2 at sn-2 position most likely reflects a weak preferential selectivity
of ATGL for C18:1 over C18:2 (Fig. 20B). All in all, these data suggest that the saturation-level of TAG-
bound FAs is not a crucial factor of ATGL-dependent TAG hydrolysis.
FIGURE 20. ATGL hydrolyzes TAGs with FAs differing in saturation-level in vitro. A, B, Homogenates of Cos7-cells
expressing ATGL were incubated in the absence or presence of CGI-58 with different, heterogeneously esterified TAG
species as substrate for 1 h at 37°C. Released FAs were measured using NEFA-C kit. Data are normalized to LacZ and
presented as means +/- S.D. and are representative for 2 independent experiments. Statistical significance was determined
applying Student´s unpaired t-test (***, p<0,001; **, p<0,01). (Fig. 20A)§.
0
50
100
150
200
250
300
350
400
FA (
nm
ol/
mg
pro
tein
*h)
ATGL/LacZ
ATGL/CGI-58***
18:1/18:1/18:1 18:1/18:1/16:016:0/18:1/18:1
18:1/16:0/18:1
triglyceride
0
50
100
150
200
250
300
350
400
FA (
nm
ol/
mg
pro
tein
*h)
ATGL/LacZ
ATGL/CGI-58**
18:1/18:1/18:1
triglyceride
18:1/18:0/18:1 18:1/18:2/18:1
A B
Results
40
FA-analysis by gaschromatography with flame-ionization detection was used
to assess the FA-selectivity of ATGL in vivo
To examine FA selectivity of ATGL in vivo, ATGLko mice and their wt littermates were used to
determine FA composition of WAT-TAG as well as FA composition of plasma lipids. This data should
indicate whether ATGL differentially hydrolyzes FA species in vivo. For this purpose, derivatization of
FAs to their corresponding methylesters (FAMEs) and analysis by gaschromatography with flame-
ionization detection (GC-FID) was performed. GC-FID exhibits the advantage that the
concentration/signal ratio is constant over a broad analyte concentration range and is equal for all
detectable FA species. To test the linearity of the method, different concentrations of FA species
were analyzed. Measurements of different concentrations of e. g. C18:1 and C18:3 showed accurate
linearity of the received signals, which made this method suitable for FA determination (Fig. 21A). A
second advantage of GC-FID is the excellent separation over a broad range of FA species in a short
time frame. The naturally most abundant FA species were clearly peak-separated within a time range
of around 17 min (Fig. 21B).
FIGURE 21. GC-FID analysis yields linear signal/concentration ratios as well as clear separation of different FAs analyzed
as FAMEs. A, Different concentrations of either oleic (upper panel) or linoleic acid (lower panel) were analyzed using GC-
FID. B, Chromatogram of a mixture of most abundant saturated and unsaturated FAs as FAMEs.
y = 26,49x + 18,111R² = 0,9979
0
500
1000
1500
2000
2500
3000
3500
4000
4500
5000
pe
ak a
rea
(AU
/10
0)
oleic acid (C18:1)
y = 25,458x + 7,4352R² = 0,9996
0
500
1000
1500
2000
2500
3000
3500
4000
4500
5000
0 50 100 150 200
pe
ak a
rea
(AU
/10
0)
pmol/µl
linolenic acid (C18:3)
A B
Results
41
Accumulating TAGs in ATGLko WAT contain all FAs with highest increase in
C16:1
To elucidate the FA-selectivity of ATGL in vivo, FA composition of WAT-TAG from wt and ATGLko mice
was determined. As previously described [75], ATGLko mice exhibited drastically increased TAG
content in WAT (~4-fold) (Fig. 22A). FA composition analysis of WAT-TAG expressed as “% of total”
showed an altered FA-pattern of ATGLko animals compared to wt, with a relative increase in C16:0
and C16:1 and a relative decrease in FAs containing 18 carbon atoms (Fig. 22B).
FIGURE 22. WAT-TAGs of ATGLko mice accumulate virtually all FA species. A, WAT lipids of non-fasted wt and ATGLko
mice were extracted according to Folch et al. and TAG content was measured using Infinity-TAG kit. B, C, Extracted WAT
lipids were separated by TLC and TAGs were isolated and transesterified. FAMEs were separated and analyzed using
GC/FID. FA composition demonstrated as percentage of total (B) or as amounts per tissue weight (C). D, Calculated changes
in FA species of WAT-TAGs in ATGLko versus wt mice. Data are presented as means +/- S.D. Statistical significance was
determined applying Student´s unpaired t-test (***, p<0,001; **, p<0,01; *, p<0,05). n=4 (each genotype). (Fig. 22C,D)§.
The molar depiction showed that the absolute amounts of all FA species massively accumulate in
ATGLko WAT-TAG (Fig. 22C). Calculation of the ratio of FAs comparing ATGLko and wt showed that all
0
3
6
9
12
15
WA
T TA
G-F
A (
fold
ch
ange
, A
TGLk
o t
o w
ildty
pe
)
C14:0 C16:0 C16:1 C18:0 C18:1 C18:2 C18:3 C20:1meanchange
***
***
*** ***
***
0
100
200
300
400
500
600
C14:0 C16:0 C16:1 C18:0 C18:1 C18:2 C18:3 C20:1
WA
T TA
G-F
A (
µm
ol/
g ti
ssu
e)
wildtype
ATGLko
***
**
***
***
***
***
*** ***
0
5
10
15
20
25
30
35
40
45
C14:0 C16:0 C16:1 C18:0 C18:1 C18:2 C18:3 C20:1
WA
T TA
G-F
A (
% o
f to
tal)
wildtype
ATGLko
**
***
***
**
**
**
**
B
C
D
A
0
50
100
150
200
250
300
350
400
450
500
WA
T-TA
G (
µm
ol/
g ti
ssu
e)
wildtype
ATGLko
*
Results
42
FA species are increased around 6 times, except C16:1 which accumulated up to 12-fold in ATGLko
WAT-TAG (Fig. 22D). This data fit to results from in vitro TAG hydrolase assays where ATGL shows
highest activity against C16:1. Hence, 16:1 accumulated most prominently in WAT-TAGs of ATGLko
mice.
Feeding/fasting-dependent changes in plasma-FA composition of wt mice
inversely correlate with FA-accumulation in WAT-TAGs of ATGLko mice
Next, plasma-FA composition of wt mice after 8 h fasting and in non-fasted state was determined.
Since, the metabolic switch, following starvation leads to an elevated, ATGL-mediated release of FAs
from WAT, determination of released FA species should reflect the preference of ATGL to hydrolyze
various FA species of WAT-TAGs.
FIGURE 23. Wt mice show a distinct pattern of FAs released into plasma. Plasma lipids of non-fasted and 8 h fasted wt
mice were extracted according to Folch et al. and separated by TLC. FAs were isolated and transesterified. FAMEs were
analyzed using GC/FID. A, Concentration of plasma FAs was calculated and expressed as sum of all measured FAs. B, Plasma
FA composition demonstrated as amounts per plasma volume. C, Calculated changes in plasma FA species of non-fasted
and 8 h fasted wt mice. Data are presented as means +/- S.D. Statistical significance was determined applying Student´s
unpaired t-test (***, p<0,001; **, p<0,01; *, p<0,05). n=4; nd…not detectable. (Fig. 23B,C)§.
0
0,1
0,2
0,3
0,4
0,5
0,6
0,7
pla
sma-
FA (
mm
ol/
l)
non-fasted
fasted
***
***
0
0,05
0,1
0,15
0,2
0,25
C14:0 C16:0 C16:1 C18:0 C18:1 C18:2 C18:3 C20:4 C22:6
pla
sma-
FA (
mm
ol/
l)
non-fasted
fasted
***
***
***
***
***
* nd nd
0
1
2
3
4
C14:0 C16:0 C16:1 C18:0 C18:1 C18:2 C18:3 C20:4 C22:6
pla
sma-
FA (
fold
ch
ange
, fas
ted
/no
n-f
aste
d)
meanchange
***
***
***
*****
nd nd
A B
C
Results
43
In agreement with published data [75], FA concentration in plasma of wt mice was doubled after 8 h
of food deprivation (Fig. 23A). Furthermore, composition analysis showed that the majority of
abundant FA species were increased around 2-fold (Fig. 23B). Calculation of the release changes for
each FA species upon fasting revealed that C16:1 as well as C18:1 and C18:2 displayed significantly
higher ratios, comparing fasted and non-fasted FA levels in plasma (Fig. 23C).
Fasting-induced FA release of WAT is blunted in ATGLko mice
To test if FA release observed in wt mice is caused by ATGL-dependent WAT-TAG hydrolysis the same
experiment was performed with ATGLko mice. FA release of WAT was completely diminished in
these mice. No increase in concentration of plasma FAs were observed upon 8 h fasting (Fig. 24A).
According to that, no changes in the pattern of plasma FAs were detected (Fig. 24B, C). These results
demonstrate that ATGL is crucial for WAT lipolysis and the release of FAs into plasma.
FIGURE 24. ATGLko mice display blunted release of FA into plasma. Plasma lipids of non-fasted and 8 h fasted ATGLko
mice were extracted according to Folch et al. and separated by TLC. FAs were isolated and transesterified. FAMEs were
separated and analyzed using GC/FID. A, Concentration of plasma FAs was calculated as sum of all measured FA
concentrations. B, Plasma FA composition demonstrated as amounts per plasma volume. C, Calculated changes in plasma
FA species of non-fasted and 8 h fasted ATGLko mice. Data are presented as means +/- S.D. n=4.
0
0,05
0,1
0,15
0,2
0,25
0,3
pla
sma-
FA (
mm
ol/
l)
non-fasted
fasted
***
0
0,02
0,04
0,06
0,08
0,1
0,12
C14:0 C16:0 C16:1 C18:0 C18:1 C18:2
pla
sma-
FA (
mm
ol/
l)
non-fasted
fasted
0
1
2
C14:0 C16:0 C16:1 C18:0 C18:1 C18:2
pla
sma-
FA (
fold
ch
ange
, fas
ted
/no
n-f
aste
d)
meanchange
A B
C
Results
44
To compare the fasting response of ATGLko and wt mice, both data sets were combined. Consistent
with the blunted FA release in ATGLko mice, the plasma FA levels were decreased around 3-fold as
compared to wt mice (Fig. 25A). Changes in FA composition of ATGLko plasma mirrored the ATGL-
dependent fasting-response of wt mice. Nearly all FA species were decreased in plasma of fasted
ATGLko mice and long chain FAs were undetectable (Fig. 25B). Calculation of the release ratio
between wt and ATGLko mice showed that especially C16:1, C18:1 and C18:2 were drastically
diminished in plasma of ATGLko mice (Fig. 25C). Taken together, ATGL-dependent hydrolase activity
in WAT leads to a release of predominantly unsaturated FA species, in particular C16:1, C18:1 and
C18:2 into circulation. Furthermore, these data provide clear evidence that ATGL acts as an essential
lipase during fasting-stimulated WAT lipolysis and, hence, determines FA release.
FIGURE 25. ATGL in WAT is essential for the fasting-induced FA release into plasma. Plasma lipids of 8 h fasted wt and
ATGLko mice were extracted according to Folch et al. and separated by TLC. FAs were isolated and transesterified. FAMEs
were analyzed using GC/FID. A, Concentration of plasma FAs was calculated as sum of all measured FAs. B, Plasma FA
composition demonstrated as amounts per plasma volume. C, Calculated changes in plasma FA species of fasted wt mice
compared to ATGLko littermates. Data are presented as means +/- S.D. Statistical significance was determined applying
Student´s unpaired t-test (***, p<0,001; *, p<0,05). n=4 (each genotype); nd…not detectable. (Fig. 25B,C)§.
0
0,05
0,1
0,15
0,2
0,25
C14:0 C16:0 C16:1 C18:0 C18:1 C18:2 C18:3 C20:4 C22:6
pla
sma-
FA, f
aste
d (
mm
ol/
l)
wildtype
ATGLko
***
***
***
*** ***
*
nd nd nd
0
0,1
0,2
0,3
0,4
0,5
0,6
0,7
pla
sma-
FA, f
aste
d (
mm
ol/
l)
wildtype
ATGLko
***
***
0
1
2
3
4
5
6
7
8
9
C14:0 C16:0 C16:1 C18:0 C18:1 C18:2 C18:3 C20:4 C22:6
pla
sma-
FA, f
aste
d (
fold
ch
ange
, wild
typ
e/A
TG
Lko
)
meanchange
***
***
***
***
nd nd nd
A B
C
Results
45
ATGL affects FA composition of plasma TAG
To assess possible effects of ATGL-dependent WAT-FA release on the FA composition of VLDL-TAG,
the FA composition of plasma TAG of 8 h fasted ATGLko and wt mice was determined using GC-FID.
In wt, plasma TAG levels were 4-times higher compared to ATGLko (Fig. 26A). Analysis of FA
composition revealed that most of the FA species were in about 3-fold higher (Fig. 26B, C) in plasma
TAG of wt mice as compared to ATGLko mice. Furthermore, plasma TAG-FA composition of wt mice
showed highest specific increase in C16:1 (6-fold) followed by C18:1 (5-fold). This indicates that the
loss of ATGL leads to most prominent decrease in unsaturated FAs, which presumably is a result of
decreased hydrolytic activity in WAT of these mice.
FIGURE 26. FA composition of plasma TAG is changed in ATGLko mice. Plasma lipids of 8 h fasted wt and ATGLko mice
were extracted according to Folch et al. and separated by TLC. TAGs were isolated and transesterified. FAMEs were
analyzed using GC/FID. A, Concentration of plasma TAGs were measured using Infinity-TAG kit. B, Plasma TAG-FA
composition shown as concentration per plasma volume. C, Numeric changes in plasma TAG-FA species of fasted wt mice
compared to ATGLko littermates. Data are presented as means +/- S.D. Statistical significance was determined applying
Student´s unpaired t-test (***, p<0,001). n=4 (each genotype).
0
1
2
3
4
5
6
7
pla
sma
TAG
-FA
, fas
ted
(m
mo
l/l)
wildtype
ATGLko
***
***
0
1
2
3
4
5
6
7
8
C 14:0 C 16:0 C 16:1 C 18:0 C 18:1 C 18:2 C 18:3 C 20:4 C 20:6
pla
sma
TAG
-FA
, fa
ste
d (
fold
ch
ange
, wild
typ
e/A
TGLk
o)
meanchange
***
***
0
1
2
3
4
5
6
C 14:0 C 16:0 C 16:1 C 18:0 C 18:1 C 18:2 C 18:3 C 20:4 C 20:6
pla
sma
TAG
-FA
, fas
ted
(m
mo
l/l)
wildtype
ATGLko
***
***
***
***
A B
C
Results
46
ATGL affects TAG content in the brain
To additionally investigate the effect of ATGL on brain lipid metabolism, neutral and polar lipid
fraction of whole brains, from 8 h fasted ATGLko and wt mice, were analyzed.
FIGURE 27. TAGs accumulate most abundant FA species in brain of ATGLko mice. Brain lipids of wt and ATGLko mice were
extracted according to Folch et al. and separated by TLC. Total lipid extract or TAGs isolated after TLC were transesterified.
FAMEs were analyzed using GC/FID. FA-composition of PLs (A), CEs (B), DAGs (C) and total brain lipids (D) shown as
percentage of total. E, Brain TAG content was calculated as sum of all measured FA species. F, Brain TAG-FA composition
shown as amount per tissue weight. Data are presented as means +/- S.D. Statistical significance was determined applying
Student´s unpaired t-test (***, p<0,001; *, p<0,05). n=5 (each genotype). (Fig. 27E,F modified after [245])
100
200
300
400
500
600
700
800
(nm
ol/
g ti
ssu
e)
wildtype
ATGLko***
***
***
*
0
20
40
60
80
100
C 14:0 C 16:0 C 16:1 C 18:0 C 18:1 C 18:2 C 18:3 C 20:1 C 20:4 C 22:6
bra
in T
AG
-FA
, fas
ted
*
0
5
10
15
20
25
30
35
C 14:0 C 16:0 C 16:1 C 18:0 C 18:1 C 18:2 C 18:3 C 20:1 C 20:4 C 22:4 C 22:6
bra
in li
pid
-FA
, fas
ted
(%
of
tota
l)
wildtype
ATGLko
0
100
200
300
400
500
600
700
800
bra
in T
AG
, fas
ted
(n
mo
l/lg
tis
sue
)
wildtype
ATGLko***
0
10
20
30
40
50
60
C 14:0 C 16:0 C 18:0 C 18:1 C 18:2 C 20:4
bra
in D
AG
-FA
(%
of
tota
l)
wildtype
ATGLko
0
10
20
30
40
50
60
70
C 16:0 C 18:0 C 18:1
bra
in C
E-FA
(%
of
tota
l)
wildtype
ATGLko
0
5
10
15
20
25
30
C 14:0 C 16:0 C 16:1 C 18:0 C 18:1 C 18:2 C 18:3 C 20:1 C 20:4 C 22:4 C 22:6
bra
in P
L-FA
(%
of
tota
l)
wildtype
ATGLko
A
C
E F
D
B
Results
47
Total brain lipids or single lipid species were isolated by TLC and FA compositions were determined as
FAMEs by GC-FID. FA composition of PLs, CEs, and DAGs were unaltered in ATGLko mice as compared
to wt mice (Fig. 27A, B, C). Similarly the FA composition of total brain lipids from ATGLko and wt mice
was identical (Fig. 27D). In contrast, large differences were found in total TAG content and TAG-FA
composition. TAG content in ATGLko brain exceeded that of wt about 14-fold (Fig. 27E). FA
composition analysis revealed that all TAG-FAs of brain were increased between 5 and 30-fold in
ATGLko mice as compared to wt mice. Besides that, many FA species, including C16:1 as well as long-
chained docosahexanoic acid (C22:6), which were not detectable in wt mice, were highly increased in
ATGLko mice (Fig. 27F). Although the role of ATGL and TAGs in brain is not known, the changes in
TAGs clearly speak for an active, ATGL-dependent TAG turnover in brain (results published in [245]).
In summary, in vitro and in vivo results of this section show that ATGL is able to hydrolyze all major
saturated und unsaturated FAs occuring in TAG. ATGL exhibits moderate preference for unsaturated
FA in particular C16:1. Even if C16:1 is not very abundant, increases in C16:1 levels were ubiquitous
when neutral lipids of ATGLko mice were investigated.
C) Substrate selectivity
Amphiphatic PLs form the lipid droplet surface monolayer, thereby enclosing lipophilic TAGs and
emulsifying them in the aqueous environment. Since ATGL belongs to a protein family which also
comprises phospholipases [65] it was tested whether ATGL also hydrolyzes LD-associated PLs.
ATGL hydrolyzes PLs independent of CGI-58
First, the hydrolytic activity of ATGL against PC, which is the major PL species on LD surface, was
investigated. Therefore, murine ATGL was expressed in Cos7-cells and hydrolase activity assays using
PC as substrate were performed. Purified PLA2 (naja mossambica m.) served as positive control. PC-
hydrolase assays were performed in the presence and absence of 1 mM Ca2+, since many, including
snake venom PLA2, hydrolyze PLs in a Ca2+-dependent manner. Little activity was observed for snake
venom PLA2 in the absence of Ca2+. Addition of Ca2+ led to a 32-fold increase in phospholipase
activity (Fig. 28A). To investigate the phospholipase activity of ATGL, PC-hydrolase assays in the
presence or absence of Ca2+ and purified GST-tagged CGI-58 were performed. ATGL exhibited
Results
48
hydrolytic activity against PC, which was independent of Ca2+ (Fig. 28B). Furthermore, the PC
hydrolase activity of ATGL was not co-activated by CGI-58 (Fig. 28B) which is in contrast to effects
observed for ATGL-dependent TAG hydrolase activity.
FIGURE 28. ATGL hydrolyzes PC independent of calcium and CGI-58. Purified phospholipase A2 (1 IU, naja mossambica m.)
(A) or homogenates of Cos7-cells expressing ATGL (B) were incubated in the absence or presence of purified GST-tagged
CGI-58 (B) or 1 mM Ca2+
(A,B) with PC as substrate for 1 h at 37°C. Released FAs were measured using NEFA-C kit. Data are
normalized to LacZ and presented as means +/- S.D. and are representative for 2 independent experiments.
Phospholipase activity of ATGL is co-activated by CGI-58 when PLs are mixed
with TAG
To investigate whether a LD-like lipid mixture is a better substrate for ATGL-dependent
phospholipase activity, experiments were performed using different neutral lipids emulsified with PC
as substrate. First, 14C-labeled PC micelles were used as a substrate and incubated with homogenates
of Cos7-cells expressing ATGL. As expected, ATGL hydrolyzed PC and was not co-activated by CGI-58
(Fig. 29A). Next, non-radiolabeled CE was emulsified with non-radiolabeled PC. This substrate should
reflect liposomes, which contain a hydrophobic core that cannot be hydrolyzed by ATGL. In this
setup, ATGL showed hydrolase activity against PC independent of CGI-58 (Fig. 29B). So far, the
discrepancy in absolute values of hydrolase activity using either radiolabeled or non-radiolabeled PC
substrates is unclear. Next, experiments were performed using 3H-labeled triolein emulsified with
14C-labeled PC. Both lipids are potential substrates of ATGL, and so, generated liposomes more or less
reflect the composition of a cytosolic LD. Unexpectedly, FAs released of PC showed that ATGL
exhibited the same activity against PC but increased activity 2-fold when co-activated by CGI-58 (Fig.
29C).
0
2
4
6
8
10
12
14
16
18
20
-Ca2+ +1mM Ca2+
FA (
nm
ol/
mg
pro
tein
*h)
ATGL ATGL/CGI-58
0
2
4
6
8
10
12
14
16
18
-Ca2+ +1mM Ca2+
FA (
mm
ol/
mg
pro
tein
*h)
phospholipase A2 (najamossambica m.)
A B
Results
49
FIGURE 29. The co-activation of the phospholipase activity of ATGL by CGI-58 depends on the neutral lipid species
emulsified within PC liposomes. Homogenates of Cos7-cells expressing ATGL were incubated in the absence or presence of
purified GST-tagged CGI-58 with 14
C-labeled PC (A) or CE emulsified with non-labeled PC (B) or 3H-labelled TAG emulsified
with 14
C-labeled PC (C,D) or non-labeled TAG emulsified with 14
C-labeled PC (E) as substrate for 1 h at 37°C. Released FAs
were determined by either NEFA-C kit (B) or scintillation counting (A, C, D). Data are normalized to LacZ and presented as
means +/- S.D. and are representative for 2 independent experiments. Statistical significance was determined applying
Student´s unpaired t-test (***, p<0,001; **, p<0,01).
0
1
2
3
4
5
6
7
ATGL ATGL/CGI-58
FA (
nm
ol/
mg
pro
tein
*h)
**
14C-phosphatidylcholine3H-triacylglycerol
14C-FA release
0
50
100
150
200
250
ATGL ATGL/CGI-58
FA (
nm
ol/
mg
pro
tein
*h)
***
14C-phosphatidylcholine3H-triacylglycerol
3H-FA release
0
1
2
3
4
5
6
ATGL ATGL/CGI-58
FA (
nm
ol/
mg
pro
tein
*h)
14C-phosphatidylcholine micelle
0
2
4
6
8
10
12
14
16
18
20
ATGL ATGL/CGI
FA (
nm
ol/
mg
pro
tein
*h)
phosphatidylcholinecholesterylester
0
2
4
6
8
10
12
ATGL ATGL/CGI-58
FA (
nm
ol/
mg
pro
tein
*h)
***
14C-phosphatidylcholinetriacylglycerol
14C-FA release
A B
C D
E
Results
50
Determination of 3H-labeled FA release revealed that the TAG hydrolase activity of ATGL was 9-fold
co-activated upon addition of CGI-58 (Fig. 29D). To exclude an error of measurement caused by an
interference of simultaneously used 3H- and 14C-labeling of FAs, experiments were performed using
non-labeled triolein emulsified with 14C-labeled PC. As observed in experiments using labeled TAG
and PC, ATGL hydrolyzed PC and increased activity 3-fold upon CGI-58 co-activation (Fig. 29E). These
data indicate that ATGL hydrolyzes PC and that the PC hydrolase activity of ATGL can be co-activated
by CGI-58 when PC is mixed with TAG. Furthermore, TAG hydrolase activity of ATGL and ATGL co-
activated by CGI-58 is 10-fold and 45-fold higher, respectively, as compared to PC hydrolase activity.
ATGL hydrolyzes most abundant PL species
To investigate if ATGL is able to hydrolyze PL species other than PC, PL hydrolase assays were
performed using homogenates of Cos7-cells expressing ATGL and PC, phosphatidylserine (PS), PE, PA
and phosphatidylglycerol (PG) as substrates. ATGL hydrolyzed all investigated PLs in micellar form
and showed comparable activities against PC, PS and PE as well as increased activity against PA and
PG (Fig. 30). These data suggest that ATGL exhibits phospholipase activity against a broad spectrum
of PLs, thereby slightly preferring PA and PG.
FIGURE 30. ATGL hydrolyzes highly abundant PLs, favoring PA and PG. Homogenates of Cos7-cells expressing ATGL were
incubated with different PL species as substrate for 1 h at 37°C. Released FAs were determined by NEFA-C kit. Data are
normalized to LacZ and presented as means +/- S.D. and are representative for 2 independent experiments. Statistical
significance was determined applying Student´s unpaired t-test (***, p<0,001).
0
5
10
15
20
25
30
PC PA PS PE PG
FA (
nm
ol/
mg
pro
tein
*h)
***
***
Results
51
Thus, ATGL can act as phospholipase, with preference for PG and PA as substrates. Thereby, the
position selectivity of ATGL as well as the impact of the phospholipase activity of ATGL on cellular
lipid homeostasis needs to be investigated.
II) Selectivity of DAG hydrolysis and re-esterification
This section investigates the isomer-selectivity of possible downstream reactions of DAGs generated
by ATGL during lipolysis. On the one hand, the re-esterification of DAG by either DGAT1 or DGAT2,
and on the other hand, the degradation of DAG by HSL (Fig. 31).
FIGURE 31. Potential hydrolysis and re-esterification reactions of DAG generated on cytoplasmic LDs. DAG can act as
precursor for TAG-synthesis catalyzed by either DGAT1 or DGAT2. Furthermore, DAG can be a substrate for degradation by
HSL yielding MAG.
A) Stereo/regioselectivity of DGAT enzymes
The regeneration of TAG by esterification of DAG with FA-CoA is one possible reaction of lipolysis-
derived DAGs. This reaction is catalyzed by either DGAT1 or DGAT2. If one of these enzymes exhibit
certain selectivity for DAG isomers is unknown and topic of this section.
Results
52
DGAT1 and DGAT2 differentiate between DAG isoforms and are highly active
on nano-sized liposomes
To investigate stereo/regioselectivity of DGAT enzymes in vitro, murine DGAT1 and DGAT2 were
expressed in Cos7-cells (Fig. 33A). Acyltransferase assays were performed using different DAG
isoforms as well as different DAG/PC compositions. The published method of DGAT activity assays
describes a DAG/PC ratio of ~1/4 [246] as substrate, which results in liposomes with a distinct size.
Given that cellular LDs differ in size, it was of interest if liposomal size affects DGAT activity.
Therefore, a constant substrate concentration of DAG (200 µM) was used while PC concentration
was decreased from 800 to 200 and 50 µM, giving DAG/PC ratios of 0.25, 1 and 4, respectively. To
examine the size of generated liposomes the substrate solutions were incubated with lipophilic
BodiPy® and liposomes were visualized using fluorescence microscopy. The pictures showed that an
increase in DAG/PC ratio, led to an increase in liposomal size. DAG/PC ratios of 0.25, 1 and 4 resulted
in a mean liposome size of 50-15 nm, 500-1000 nm and 2-3 µm, respectively (Fig. 32A, B, C).
FIGURE 32. Micelles size of DAG/PC substrate increases with increasing DAG/PC ratio. 200 µM of DAG substrate was
emulsified with either 800 µM (DAG/PC – 0.25, A), 200 µM (DAG/PC – 1, B) or 50 µM (DAG/PC – 4, C) of PC. Micelle size was
visualized by staining with BodiPy® and imaged by fluorescence microscopy.
Furthermore, either sn-1,2, rac-1,2/2,3 or sn-1,3 DAG was emulsified with PC in the mentioned ratios
and used as substrate. Both DGAT enzymes showed a reduction in activity for all provided DAG
isoforms when DAG/PC ratio was increased from 0.25 to 4 (Fig. 33B). Unexpectedly, DGAT1 and
DGAT2 exhibited different selectivity for DAG isoforms. Using a DAG/PC ratio of 0.25, at which both
DGATs were most active, DGAT1 showed highest activity against sn-1,2 and rac-1,2/2,3 DAG whereas
DGAT2 showed highest activity in esterifying sn-1,3 DAG (Fig. 33C). For further experiments a
DAG/PC ratio of 0.25 was used. Since expression levels of both DGATs differ, the selectivity quotient
for both enzymes was calculated. Results showed a 80/20 ratio of sn-1,2/sn-1,3 DAG for DGAT1 and,
in contrast, a 30/70 ratio of sn-1,2/sn-1,3 DAG for DGAT2 (Fig. 33D). Thus, DGAT1 as well as DGAT2
A B C
Results
53
prefer nano-liposomes as substrate and DGAT1 prefers sn-1,2 and sn-2,3 DAG as acceptor, while
DGAT2 favours sn-1,3 DAG.
FIGURE 33. DGAT1 and DGAT2 display different activities against DAG isoforms and micelle size. A, Expression of his-
tagged LacZ and flag-tagged murine DGAT1 and DGAT2 in Cos7-cells was assessed by immunoblotting. B, Homogenates of
Cos7-cells expressing DGAT1 or DGAT2 were incubated with either sn-1,2, rac-1,2/2,3 or sn-1,3 DAG emulsified with PC in
different molar ratios and 14
C-labeled C18:1-CoA as substrate for 10 min at 37°C. Reaction was stopped by addition of
CHCl3/MeOH (2/1, v/v). Lipids were extracted, separated by TLC and radioactivity in TAG bands was determined by
scintillation counting. C, Comparison of DGAT1 and DGAT2 activities against different DAG isoforms emulsified with PC in a
molar ratio of 800/200 (PC/DAG, µM/µM). D, Regioselectivity of recombinant DGAT enzymes against sn-1,2 or sn-1,3
diolein, expressed as percentage of total acyltransferase activity. Data are normalized to LacZ and presented as means +/-
S.D. and are representative for 2 independent experiments. Statistical significance was determined applying Student´s
unpaired t-test (***,p<0,001). (Fig. 33A,C,D)§
Selective inhibiton of DGAT enzymes is required to assign endogenous DGAT
activity to either DGAT1 or DGAT2
To additionaly investigate the stereo/regioselectivity of endogenous DGAT1 and DGAT2 in WAT, a
specific inhibition of either DGAT1 or DGAT2 is required. Therefore, a variety of DGAT inhibitors was
0
10
20
30
40
50
60
70
80
90
DGAT 1 DGAT2
rati
o (
% o
f to
tal a
cylt
ran
sfe
rase
act
ivit
y)
sn-1,2 diolein
sn-1,3 diolein
***
***
0
20
40
60
80
100
120
140
sn-1,2 rac-1,2/2,3 sn-1,3
14C
-TA
G f
orm
ed
(n
mo
l/m
g p
rote
in*h
)
DGAT1
DGAT2
***
***
diolein
0
20
40
60
80
100
120
140
0.25 1 4 0.25 1 4
14C
-TA
G f
orm
ed
(n
mo
l/m
g p
rote
in*h
)
sn-1,2 diolein
rac-1,2/2,3 diolein
sn-1,3 diolein
ratio diolein:phospatidylcholine
DGAT2DGAT1
D
A
C
B
Results
54
tested. DGAT activity assays were performed in vitro using reported inhibitors and a 50/50 mixture of
sn-1,2/sn-1,3 DAGs as substrate. Niacin, proposed as DGAT2-selective inhibitor [247] showed a 90%
inhibition of DGAT2 whereas DGAT1 activity was unaffected (Fig. 34A).
FIGURE 34. The effects of DGAT-inhibitors on the enzymatic activity of DGAT1 and DGAT2 in cell homogenates. A, B, C,
Homogenates of Cos7-cells expressing DGAT1 or DGAT2 were incubated in the absence or presence of different inhibitory
compounds with either sn-1,2 or sn-1,3 DAG or a 1/1 mixture of sn-1,2/sn-1,3 DAG substrate emulsified with PC in a molar
ratio of 800/200 (PC/DAG, µM/µM) and 14
C-labeled C18:1-CoA for 10 min at 37°C. Reaction was stopped by addition of
CHCl3/MeOH (2/1, v/v). Lipids were extracted, separated by TLC and radioactivity in TAG bands was determined by
scintillation counting. Data are presented as means +/- S.D. and are representative for 2 independent experiments.
Statistical significance was determined applying Student´s unpaired t-test (***,p<0,001).
0
20
40
60
80
100
120
140
14C
-TA
G f
orm
ed
(n
mo
l/m
g p
rote
in*h
)
--
Niacin (5mM)DGAT1 inh. (5µM)
LacZ DGAT1 DGAT2
***
***
--
+-
-+
++
--
+-
-+
++
0
20
40
60
80
100
120
140
160
14C
-TA
G f
orm
ed
(n
mo
l/m
g p
rote
in*h
)
--DGAT1 inh. (5µM)
LacZ DGAT1 DGAT2
***
***
***
MgCl2 (100mM) --
+-
-+
++
--
+-
-+
++
0
5
10
15
20
25
30
35
LacZ DGAT1 DGAT2
14C
-TA
G fo
rme
d (
nm
ol/
mg
pro
tein
*h) 10mM MgCl2
100mM MgCl2
sn-1,3 diolein
***
***
0
20
40
60
80
100
120
140
DGAT2DGAT1LacZ
14C
-TA
G fo
rme
d (
nm
ol/
mg
pro
tein
*h)
sn-1,2 diolein
***
C
A
B
Results
55
Conversely, with the same efficiency, DGAT1-specific inhibitor (2-((1s,4s)-4-(4-(4-amino-7,7-dimethyl-
7H-pyrimido[4,5-b][1,4]oxazin-6-yl)phenyl)cyclohexyl) acetic acid, [139]) inhibited DGAT1 but not
DGAT2 (Fig. 34A). Next, the effect of MgCl2 on DGAT activity was investigated. Low concentrations
are required for DGAT activity while high concentrations (>100 mM) should selectively inhibit DGAT2
[118]. As expected, a concentration of 100 mM MgCl2 led to a complete inhibition of DGAT2 activity.
Unexpectedly, the same concentration also significantly inhibited 60% of DGAT1-dependent activity.
The residual activity of DGAT1 was eliminated by using DGAT1-specific inhibitor (Fig. 34B). Since
published data only use sn-1,2 DAG as substrate, it was conceivable that enzyme inhibition depends
on substrate composition. To test this, either sn-1,3 or sn-1,2 DAG was used as substrate in
acyltransferase experiments. With sn-1,3 DAG as substrate, a higher activity of DGAT2 was observed
as compared to DGAT1 (Fig. 34C). Furthermore, 100 mM MgCl2 led to a complete inhibition of DGAT2
and a ~70% inhibition of DGAT1. In contrast, using sn-1,2 DAG as substrate, DGAT1 showed a 4-fold
higher activity compared to DGAT2. Using sn-1,2 DAG as substrate, 100mM MgCl2 had no effect on
DGAT1 activity while again completely inhibiting DGAT2. This led to the conclusion that due to the
substrate-dependent inhibition of DGAT1, MgCl2 is inappropriate to discriminate activities of
endogenous DGAT enzymes.
TAG hydrolase activity in WAT homogenates affects in vitro acyltransferase
measurements
To investigate the stereo/regioselectivity of endogenous DGAT enzymes in WAT of wt mice, the
previously tested inhibitors were applied to acyltransferase experiments using WAT homogenates.
Acyltransferase assays were performed in the presence of niacin as well as the DGAT1-specific
inhibitor [118]. Notably, results differed between niacin treatment of WAT and Cos7-cell
homogenates. In WAT homogenates DGAT1-specific inhibitor led to a 90% reduction of TAG
formation. In contrast, niacin enhanced DGAT activity, resulting in a 40% increase of TAG (Fig. 35A).
This may be due to an effect of niacin on enzymes involved in lipolysis, which may degrade TAGs
formed during acyltransferase assays. If niacin modulates lipase activity of ATGL or HSL in
acyltransferase experiments is unknown. To investigate this hypothesis, hydrolase inhibitors like HSL-
specific inhibitor (76-0079), bromoenol lactone (BEL) or orlistat were used. To test whether these
hydrolase inhibitors additionaly interfere with DGAT enzymes, homogenates of Cos7-cells expressing
DGAT1 and DGAT2 as well as described inhibitors were used in combination. Neither the two
unspecific hydrolase inhibitors BEL and orlistat, nor the HSL-specific inhibitor resulted in significant
differences of DGAT activity in homogenates of Cos7-cells expressing DGAT1 or DGAT2 (Fig. 35B).
Results
56
Next, hydrolase inhibitors as well as the DGAT1-specific inhibitor were used alone and in combination
in acyltransferase experiments using WAT homogenates.
FIGURE 35. The effects of different lipase and/or DGAT-inhibitors on the enzymatic activity of DGAT1 and DGAT2 in WAT
and cell homogenates. Homogenates of WAT from wt mice (A, C) or homogenates of Cos7-cells expressing DGAT1 or
DGAT2 (B) were incubated in the absence or presence of different inhibitory compounds with a 1/1 mixture of sn-1,2/sn-1,3
DAG substrate emulsified with PC in a molar ratio of 800/200 (PC/DAG, µM/µM) and 14
C-labeled C18:1-CoA for 10 min at
37°C. Reaction was stopped by addition of CHCl3/MeOH (2/1, v/v). Lipids were extracted according to Folch et al, separated
by TLC and radioactivity in TAG bands was determined by scintillation counting. Data are presented as means +/- S.D. and
are representative for 2 independent experiments. Statistical significance was determined applying Student´s unpaired t-
test (***,p<0,001). n=3 (pooled).
0
10
20
30
40
50
60
70
80
90
14C
-TA
G f
orm
ed
(n
mo
l/m
g p
rote
in*h
)
BEL (5µM)
Orlistat (20µM)
76-0079 (12,5µM)DGAT1 inh. (5µM)
-
WAT homogenate
---
+---
-+--
--+-
-++-
+-+-
+-++
-+++
***
***
0
10
20
30
40
50
60
70
80
90
14C
-TA
G f
orm
ed
(n
mo
l/m
g p
rote
in*h
)
DGAT1 inh. (5µM)
Niacin (5mM) --
***
***
WAT homogenate
+-
-+
0
20
40
60
80
100
120
140
160
180
14C
-TA
G f
orm
ed
(n
mo
l/m
g p
rote
in*h
)
Orlistat (20µM)
76-0079 (12,5µM)
LacZ DGAT1 DGAT2
BEL (5µM) ---
---
+--
-+-
--+
---
+--
-+-
--+
A
C
B
Results
57
Unexpectedly, inhibition of lipolytic enzymes in WAT clearly affected acyltransferase experiments.
BEL, orlistat, and HSL-specific 76-0079 resulted in a 1.7-fold increase of TAG formation. In the
presences of lipase inhibitors, DGAT1-specific inhibition led to a 90% reduction of TAG formation (Fig.
35C). These results lead to the conclusion that inhibition of TAG hydrolases is crucial for an accurate
determination of DGAT-dependent acyltransferase activity of WAT samples.
Endogenous DGAT1 and DGAT2 exhibit preference for sn-1,2 and sn-1,3 DAG,
respectively
In the next set of experiments the stereo/regioselectivity of endogenously expressed DGAT enzymes
of WAT was investigated. WAT homogenates of non-fasted C57BL/6J mice were separated into
microsomal (including plasma membranes), cytplasmic and LD fraction using ultracentrifugation.
Microsomal fractions showed highest DGAT1 and DGAT2 expression (Fig. 36A). In contrast, DGAT1
and DGAT2 were undetectable in the cytosolic fraction and in the LD fraction. Each cellular fraction
was subjected to acyltransferase assays using combinations of orlistat/76-0079 and DGAT1-specific
inhibitor and either sn-1,2 or sn-1,3 DAG as substrate.
FIGURE 36. DGAT1 and DGAT2 of WAT display different specific activities against DAG isoforms. A, Expression of
endogenous DGAT1 and DGAT2 in microsomal fraction of WAT was assessed by immunoblotting. B, Microsomal fraction of
WAT from wt mice were incubated in the absence or presence of different inhibitory compounds with either sn-1,2 or sn-
1,3 DAG substrate emulsified with PC in a molar ratio of 800/200 (PC/DAG, µM/µM) and 14
C-labeled C18:1-CoA for 10 min
at 37°C. Reaction was stopped by addition of CHCl3/MeOH (2/1, v/v). Lipids were extracted according to Folch et al,
separated by TLC and radioactivity in TAG bands was determined by scintillation counting. C, Calculated regioselectivity of
DGAT enzymes in microsomal fraction of WAT against sn-1,2 or sn-1,3 DAG expressed as percentage of total acyltransferase
activity. Data are presented as means +/- S.D. and are representative for 2 independent experiments. Statistical significance
was determined applying Student´s unpaired t-test (***,p<0,001). n=3. (Fig. 36A,B,C)§
0
10
20
30
40
50
60
70
DGAT1 DGAT2
rati
o (
% o
f to
tal a
cylt
ran
sfe
rase
act
ivit
y)
sn-1,2 diolein
sn-1,3 diolein
***
***microsomal fraction
0
20
40
60
80
100
120
140
160
180
14C
-TA
G f
orm
ed
(n
mo
l/m
g p
rote
in*h
) ***
***
76-0079 (12,5µM)DGAT1 inh. (5µM)
Orlistat (5mM)++
+sn-1,2 diolein sn-1,3 diolein
***
***
microsomal fraction
+-
+--
-++
++-
+--
-
C B A
Results
58
Microsomal fraction exhibited highest endogenous DGAT activity against both DAG isoforms.
Addition of lipase inhibitors resulted in a 1.6-fold increased formation of TAG when sn-1,2 DAG was
used as substrate. Upon inhibition of DGAT1, specific endogenous acyltransferase activity for sn-1,2
DAG was decreased by around 70% (Fig. 36B). DGAT activity of microsomal fraction was 20% lower
using sn-1,3 DAG as substrate and 1.6-fold increased upon lipase inhibiton. Addition of the DGAT1-
specific inhibitor led to a 50% reduction (Fig. 36B). Assuming that residual DGAT activity can be
attributed to DGAT2 and taken into account that expression levels of DGAT enzymes were different,
the selectivity quotient of microsomal fraction was calculated. It showed a 65/35 ratio for sn-1,2/sn-
1,3 DAG for DGAT1 and a 40/60 ratio for sn-1,2/sn-1,3 DAG for DGAT2 (Fig. 36C).
FIGURE 37. DGAT1 and DGAT2 of WAT display preference for different DAG isoforms. A, Cytoplasmic and C, LD fraction of
WAT from C57BL/6J mice were incubated in the absence or presence of different inhibitory compounds with either sn-1,2
or sn-1,3 DAG substrate emulsified with PC in a molar ratio of 800/200 (PC/DAG, µM/µM) and 14
C-labeled C18:1-CoA for 10
min at 37°C. Reaction was stopped by addition of CHCl3/MeOH (2/1, v/v). Lipids were extracted according to Folch et al,
separated by TLC and radioactivity in TAG bands was determined by scintillation counting. Calculated regioselectivity of
DGAT enzymes in B, cytoplasmic, and D, LD fraction of WAT against sn-1,2 or sn-1,3 DAG expressed as percentage of total
acyltransferase activity. Data are presented as means +/- S.D. and are representative for 2 independent experiments.
Statistical significance was determined applying Student´s unpaired t-test (***,p<0,001). n=3.
0
10
20
30
40
50
60
70
80
90
100
DGAT1 DGAT2
rati
o (
% o
f to
tal a
cylt
ran
sfe
rase
act
ivit
y)
sn-1,2 diolein
sn-1,3 diolein
***
***
cytoplasmic fraction
0
1
2
3
4
5
6
7
14C
-TA
G f
orm
ed
(n
mo
l/h
*mg)
***
***
-76-0079 (12,5µM)
DGAT1 inh. (5µM) -
Orlistat (5mM) -+-
+++
+--
-+-
+++
+
sn-1,2 diolein sn-1,3 diolein
cytoplasmic fraction
0
5
10
15
20
251
4C
-TA
G f
orm
ed
(n
mo
l/h
*mg)
***
***
-76-0079 (12,5µM)
DGAT1 inh. (5µM) -
Orlistat (5mM) -+-
+++
+--
-+-
+++
+
sn-1,2 diolein sn-1,3 diolein
******
lipid droplet fraction
0
10
20
30
40
50
60
70
80
DGAT1 DGAT2
rati
o (
% o
f to
tal a
cylt
ran
sfe
rase
act
ivit
y)
sn-1,2 diolein
sn-1,3 diolein
***
***lipid droplet fraction
A
B
C
D
Results
59
Next, DGAT activity of the cytoplasmic fraction and the LD fraction was assessed. Despite much lower
DGAT activities, comparable effects were observed using sn-1,2 DAG as substrate in combination
with lipase and DGAT1-specific inhibitors. Addition of lipase inhibitors resulted in a 3-fold increase of
TAG formation. Combined inhibition of lipases and DGAT1 decreased DGAT activity by 90%. In
contrast, no inhibitory effects were detectable using sn-1,3 DAG as substrate (Fig. 37A). Calculation
of the selectivity quotient for sn-1,2/sn-1,3 DAG led to a 90/10 ratio for DGAT1 and to a 20/80 ratio
for DGAT2 (Fig. 37B). Addition of lipase inhibitors to DGAT assays using LD fraction had no effects,
whereas addition of DGAT1-specific inhibitor led to >90% reduction of TAG formation using either sn-
1,2 or sn-1,3 DAG as substrate (Fig. 37C). The calculated selectivity quotients of DGAT activity within
LD fraction were similar to that obtained for microsomal and cytosolic fraction. DGAT1 exerts 70%
preference for sn-1,2 DAG whereas DGAT2 prefers sn-1,3 DAG (70%, Fig. 37D).
Taken together, results of this section demonstrate that DGAT1 as well as DGAT2 exhibit
regioselectivity for DAG species. DGAT1 prefers sn-1,2 DAG as substrate, which is mainly generated
during de novo lipid synthesis, located at the ER. In contrast, DGAT2 predominantly esterifies sn-1,3
DAG, which is generated by ATGL-dependent hydrolysis at the LD (Fig. 38).
FIGURE 38. Cellular acyltransferase-reactions utilizing DAG. sn-1,3 DAG, which is generated by ATGL in the course of
lipolysis displays the preferred substrate for DGAT2-dependent re-esterification. In contrast, DGAT1 predominantly
esterifies sn-1,2 DAG, which is mainly produced during de novo lipid synthesis at the ER.
Results
60
B) Stereo/regioselectivity of HSL-dependent DAG hydrolysis
In the process of lipolysis the breakdown of DAG on cytoplasmic LDs is catalyzed by HSL.
This section investigates the selectivity of HSL to degrade specific DAG isoforms. Since HSLko mice
accumulate large amounts of DAG, which can influence insulin signaling, insulin tolerance was
additionally assessed.
HSL preferentially hydrolyzes sn-1,3 DAG
First, the stereo/regioselectivity of the second consecutive reaction in the breakdown of TAG, namely
the degradation of DAG by HSL, was investigated. Hydrolase experiments were performed using
Cos7-cell homogenates containing murine HSL (Fig. 39A) and different DAG isoforms as well as TAG
as substrates. As previously described [23], HSL showed around 10-fold higher activity against
racemic DAG as compared to TAG. HSL exhibited highest hydrolase activity against sn-1,3 DAG when
sn-1,2, rac-1,2/2,3, and sn-1,3 DAG were separately used as substrate. DAG hydrolase activity of HSL
with sn-1,3 DAG as substrate was 1.5-fold increased as compared to hydrolase activity using either
sn-1,2 or rac-1,2/2,3 DAG. (Fig. 39B). This result indicates that sn-1,3 DAG displays a preferred
substrate for HSL.
FIGURE 39. HSL preferentially hydrolyzes sn-1,3 DAG. A, Expression of his-tagged LacZ and HSL in Cos7-cells was assessed
by immunoblotting. B, Homogenates of Cos7-cells expressing HSL were incubated with either TAG or isomeric different DAG
species as substrate, emulsified with PC for 1 h at 37°C. Generated FAs were measured using NEFA-C kit. Data are
normalized to LacZ and presented as means +/- S.D. and are representative for 2 independent experiments. Statistical
significance was determined applying Student´s unpaired t-test (***,p<0,001). (Fig. 39A,B)§
0
50
100
150
200
250
300
350
triolein rac sn-1,2 rac-1,2/2,3 sn-1,3
FA (
nm
ol/
mg
pro
tein
*h)
***
diolein
***
B A
Results
61
HSLko mice show signs of increased insulin sensitivity fed a normal chow diet
To address the question if DAG accumulation in HSLko mice causes IR, studies were performed using
male HSLko mice and wt littermates fed a normal chow diet (CD). NMR analysis revealed that body
composition of CD fed HSLko mice did not differ from the body composition of wt mice. Both,
showed comparable body weights (HSLko: 22.8±1.9 g; wt: 24.5±0.7 g) as well as 75% lean mass
versus 20% of fat mass (Fig. 40A). To test the effect of insulin, an intraperitoneal insulin tolerance
test (IPITT) was performed. Mice were fasted for 2 h following an intraperitoneal bolus of bovine
insulin (0.6 IU/kg body weight). Subsequently, blood glucose levels were measured at different time
points. Results of this IPITT showed that HSLko mice displayed increased insulin response as
compared to wt mice. This is evident by a significant decrease of blood glucose after 50 min of
injection as compared to wt mice (Fig. 40B). Calculating the area under the curve for both mice
showed that HSLko mice displayed significantly increased insulin tolerance (Fig. 35C). Accordingly,
the accumulation of DAG in tissues of HSLko mice does not promote the development of IR, but
instead increases insulin sensitivity.
FIGURE 40. HSLko mice show increased insulin sensitivity fed a normal CD. A, Body composition of male HSLko and
wildtype mice fed a normal CD was determined using NMR. B, Mice were fasted from 8:00 to 10:00 am and received an
intraperitoneal bolus of bovine insulin (0.6 IU/kg body weight). Blood glucose was determined at indicated time points
using Accu-Check glucometer. C, Areas under the curves were calculated using trapezoid method. Data are presented as
means +/- S.D. Statistical significance was determined applying Student´s unpaired t-test (*,p<0,05); n=4 (each genotype).
0
10
20
30
40
50
60
70
80
90
wildtype HSLko
we
igh
t (%
of
tota
l)
lean mass
fat mass
A B
0
20
40
60
80
100
120
140
160
180
200
0 50 100 150 200
glu
cose
(m
g/d
l)
time (min)
wildtype
HSLko
0
5
10
15
20
25
30
wildtype HSLko
are
a u
nd
er
the
cu
rve
(A
U/1
00
0)
*
C
Results
62
HSLko mice show no signs of impaired insulin signaling fed a HFD
To further investigate insulin signaling, male HSLko and wt mice were fed a HFD. Subsequently, body
composition was determined using NMR. Wt mice showed an extensive, 2.5-fold increase of fat mass
whereas HSLko mice showed a moderate 1,4-fold increase in fat mass as compared to that of mice
fed normal CD (Fig. 41A). To study insulin signaling, mice were fasted for 8 h following a refeeding
period of 1 h. After that, mice were sacrificed and liver as well as SM (m. gastrocnemius) samples
were collected. Tissues were homogenized and separated into cytoplasmic and microsomal fraction
(including plasma membranes) using ultracentrifugation. Samples were then subjected to
immunoblot analysis of proteins potentially involved in insulin signaling. GAPDH, which displays a
cytoplasmic protein, was not detectable in microsomal fractions but showed comparable expression
in cytoplasmic samples, which confirmed purity of fractionation.
FIGURE 41. HSLko mice show no signs of impaired insulin signaling fed a HFD. A, Body composition male HSLko and
wildtype mice fed a HFD was determined using NMR. B, Mice were fasted from 12:00 to 8:00 am and reefed for 1 h.
Subsequently mice were sacrificed and liver and gastrocnemius muscle were collected. Tissue homogenates were separated
into cytoplasmic and microsomal fraction (including plasma membranes) by ultracentrifugation. 40 µg of cellular fractions
were used to determine expression of indicated proteins by immunoblotting. Data are presented as means +/- S.D.; n=4
(each genotype).
0
10
20
30
40
50
60
70
80
wildtype HSLko
we
igh
t (%
of
tota
l)
lean mass
fat mass
B A
Results
63
Furthermore, HSL expression was missing in tissues of HSLko mice, proving correct genotypes (Fig.
41B). Tissue specific PKC isoforms, namely PKCθ in skeletal muscle and PKCε in liver showed
increased expression in cytoplasmic fraction of wt compared to HSLko mice and were much less
expressed in microsomal fractions. Both PKC isoforms are known to be positively correlated with
defective insulin signaling. PKCα which is induced by insulin showed slightly increased expression in
microsomal fractions of HSLko tissues. PKCα showed higher expression in microsomal fraction
(translocation) of SM as compared to the expression in cytoplasmic fraction. This translocation was
not observable in liver, were expression in cytoplasmic fraction was higher as compared to
microsomal fraction (Fig. 41B).
Together, the results of IPITT and analysis of insulin signaling in wt and HSLko mice showed that
HSLko mice are more insulin responsive as compared to wt mice and do not show defects in insulin
signaling even on a HFD.
Discussion
Discussion
65
During the last decades the incidence of obesity and concomitant T2DM and IR in the western world
increased alarmingly. Elevated levels of circulating free FAs are known to be involved in the
development of IR in peripheral tissues [167]. These circulating free FAs are deposited in non-adipose
tissues, a phenomenon called ectopic fat deposition [248]. The lipid overload of these tissues also
promotes the accumulation of intracellular signaling molecules, which causes cell dysfunction and
damage (generally termed as lipotoxicity) [249, 250]. Specifically the enrichment of DAGs and
ceramides is suggested to result in an inhibition of insulin signaling [230], caused by an activation of
c/nPKC and aPKC, respectively. This activation blocks the phosphorylation and/or translocation of
important proteins involved in insulin-signaling and inhibits cellular insulin response [230].
In cells, DAG can be generated by three different pathways. (i) The breakdown of PLs catalyzed by
PLC, (ii) the glycerolipid de novo synthesis (by the consecutive reaction of GPATs, AGPATs and
PAPases/lipins) or (iii) the hydrolysis of TAG, catalyzed by TAG hydrolases, like ATGL or HSL. All of the
mentioned pathways are located in different compartments of the cell. Degradation of PLs catalyzed
by phospholipases occurs mainly at the plasma membrane. In contrast, most of the enzymes involved
in DAG synthesis are located at ER membranes. Finally, TAG hydrolysis occurs mainly at the surface of
LDs. DAG generation in different cellular compartments suggests that their metabolic fate as well as
their signaling potential may differ. The signaling potential of DAGs also depends on their
stereospecific conformation. Several studies delineated that only sn-1,2 but not the two other DAG
isoforms, sn-2,3 and sn-1,3, are able to activate novel and conventional PKC isoforms [152-154].
Moreover, several studies have shown that increased levels of DAG in peripheral tissues but not of
adipose tissue are associated with IR in both rodents and humans [177]. Together, these findings
argue for distinct DAG pools in non-adipose and adipose tissues, which may differ in their
stereochemistry as well as in their localization. PKC-activating sn-1,2 DAG is the intermediate of both
PL/TAG de novo synthesis and the hydrolysis of PLs by PLC-dependent activity. In contrast, the
stereochemistry of DAG, which derives from the lipolytical breakdown of TAG, catalyzed mainly by
ATGL, is unknown and may explain why adipose tissues do not develop IR.
In this study the stereo/regioselectivity of ATGL was investigated to unravel DAG isoforms, which are
formed in the initial step of TAG hydrolysis. In addition, the FA-preference of ATGL as well as the
activity of ATGL against PL substrates was studied. Results showed that ATGL hydrolyzes all, in
rodents highly abundant, species of long-chain, saturated, and unsaturated FAs in vitro. Highest
activity was observed for C16:1 and C18:1 esters. The same selectivity of ATGL was concluded from
Discussion
66
the analysis of neutral lipids, which accumulate in adipose tissue of ATGLko mice in vivo.
Interestingly, ATGL also hydrolyzes LD-associated PLs. Yet, in comparison to TAG hydrolysis the rate
for PL hydrolysis is much lower. However, the regioselectivity of ATGL for PL-bound FAs is currently
unknown. This study also demonstrated that ATGL by itself (without co-activation by CGI-58)
hydrolyzes TAG highly selective at sn-2 position, thereby generating sn-1,3 DAG. Interestingly, the sn-
2 regioselectivity of ATGL expands to sn-1 position upon co-activation by CGI-58, generating
additionally sn-2,3 DAG. Notably, ATGL did not hydrolyze sn-3 esters, hence did not generate sn-1,2
DAG in detectable amounts. Additionally, HSL and DGAT2, which catalyze important, subsequent
reactions of DAG, also prefer the generated sn-1,3 DAG isomer as substrate.
Results of this study, in combination with published data, reveal the entire selectivity of the enzymes
involved in TAG catabolism of adipose tissue LDs, based on stereo/regiochemical considerations (Fig.
42).
FIGURE 42. Stereo/regioselectivity of enzymes involved in the lipolytic breakdown of TAG. TAG is hydrolyzed at sn-2
position by ATGL, or at sn-1 or sn-2 position by ATGL co-activated by CGI-58 yielding sn-1,3 DAG or sn-1,3 and sn-2,3 DAG,
respectively. All of the generated DAG species exhibit a FA ester at sn-3 position, which is preferentially hydrolyzed by HSL
yielding either sn-1 or sn-2 MAG. MGL does not exhibit preference for MAG isomers and hydrolyzes both sn-1 and sn-2
MAG. ATGL, adipose triglyceride hydrolase; CGI-58, comparative gene identification-58; DAG, diacylglycerol; FA, fatty acid;
G, glycerol; HSL, hormone-sensitive lipase; MAG, monoacylglycerol; MGL, monoglyceride lipase; TAG, triacylglycerol.
Discussion
67
In basal state, ATGL generates sn-1,3 DAG. ATGL additionally generates sn-2,3 DAG upon co-
activation by CGI-58. Both generated DAG species are preferred substrates for sn-3 selective HSL
[27], which generates sn-1 and sn-2 MAG. MGL finally catabolizes both provided MAG isoforms [93].
Thus, the stereo/regioselectivity of the entire enzymatic cascade catalyzing TAG breakdown at
cytoplasmic LDs suggests, that the involved enzymes have co-evolutionary developed in consecutive
reactions of lipolysis to optimize hydrolytic efficiency (Fig. 42).
The stereo/regioselectivity of ATGL in the absence of the co-activator CGI-58 is unique in that ATGL is
so far the only mammalian lipase described to hydrolyze TAG selectively at sn-2 position. Earlier
studies, which focused on the stereochemical characterization of a variety of animal and microbial
lipases clearly showed that the large majority of lipases exhibit sn-1 or sn-3 selectivity. Only very few
microbial lipases are able to catalyze the hydrolysis of the secondary ester bond at sn-2 position of
TAG. Out of those, only Candida Antarctica A lipase was found to exhibit a clear positional selectivity
for the sn-2 position [14]. Furthermore, all stereochemically characterized TAG lipases involved in
digestion and lipid absorption, like lingual lipase (LL), PAL, and gastric lipase (GL) exhibit preference
for sn-1 or sn-3 position of TAG. PAL, GL, and LL hydrolyze TAG specifically at the sn-3 position but
also hydrolyze DAGs. Thus, they break down TAG into FAs and MAG [13, 14, 251-253]. An example
for a positionally unspecific lipase is bile-salt stimulated lipase (BSSL), which hydrolyzes all positions
of TAG, generating glycerol and FAs [254]. In contrast to intestinal lipases, lipoprotein lipase (LPL),
which depletes circulating lipoproteins from TAGs, exhibits sn-1 specificity for TAG and also
hydrolyzes DAG at the sn-2 position, generating FAs and sn-3 MAG [13, 14, 255].
Intracellularly, several other lipases are supposed to hydrolyze TAG. In this context, a number of
studies focused on triacylglycerol hydrolase/carboxylesterase 3 (TGH1/Ces3), which is mainly
expressed in liver, and to a lower extent in WAT, kidney, and heart [256, 257]. TGH/Ces3 localizes to
the ER, especially to areas surrounding cytosolic LDs and mitochondria [258], and catalyzes the
hydrolysis of long-, medium-, and short-chained TAG substrates with so far unknown
stereoselectivity [257, 259]. Besides TGH1/Ces3, several members of PNPLA protein family, like
adiponutrin (PNPLA3), GS2 (PNPLA4), and GS2-like (PNPLA5) as well as triacylglycerol hydrolase
2/carboxylesterase ML1 (TGH2), are described to possess TAG-hydrolase activity, even though
activities are not comparable with that of ATGL. For most of them, like TGH2, GS2-like, or adiponutrin
the stereo/regioselectivity, if any, is unknown. Only human GS2 was shown to hydrolyze TAG at rac-
1/3 and sn-2 position [74]. So far, more detailed studies on the physiological role of GS2 are lacking,
since the mouse genome lacks GS2 and the rat GS2 lacks activity [66, 74]. Besides ATGL, HSL is the
only other intracellular TAG hydrolase involved in TAG-turnover. HSL exhibits also DAG, MAG, CE, and
Discussion
68
RE hydrolase activity. Thus, if HSL hydrolyzes TAG, it most likely also degrades the product (DAG) to
MAG and FAs. HSL exhibits sn-1/3 preference for TAG and hydrolyzes DAG predominantly at sn-3
position, thereby generating sn-2 MAG [27, 260]. Consistent with previous reports, in vitro studies of
this work revealed that HSL prefers sn-1,3 DAG and not sn-1,2 or rac-1,2/2,3 DAG as substrate. In
contrast, ATGL exhibits clear sn-2 preference for TAG hydrolysis and does not hydrolyze DAG. Upon
co-activation by CGI-58, ATGL becomes much more active and expands its selectivity (to sn-1). The
mechanism by which CGI-58 co-activation of ATGL broadens the regioselectivity of the enzyme
remains to be elucidated. The regioselectivity of ATGL may have important physiological relevance. It
can be assumed that during basal hydrolysis of TAG, when ATGL is not co-activated by CGI-58, only
MUFAs and PUFAs are released from the sn-2 position of TAG since unsaturated FAs are very
abundant at the sn-2 position of TAG [261]. In contrast, activation of lipolysis by hormones, like ß-
adrenergic receptor agonists, leads to stimulation of lipolysis and co-activation of ATGL by CGI-58.
During stimulated lipolysis ATGL broadens its stereo/regioselective spectrum from sn-2 to sn-1
position, thus generating both sn-1,3 and sn-2,3 DAGs. In addition, stimulated lipolysis also leads to
increased FA mobilization, which are predominately released into circulation and used from
peripheral tissues for energy production.
Another aim of this study was to investigate the FA preference of ATGL. In vitro measurements
showed that ATGL exhibits highest activities against tripalmitolein, followed by triolein, trilinolenin,
and trilinolein. As expected, upon co-activation by CGI-58 ATGL hydrolase activity increased but FA-
selectivity remained unchanged. Furthermore, no major differences in ATGL and ATGL/CGI-58-
dependent activity were found when one of the C18:1 esters of trioleate was substituted by a
saturated FA, like C16:0 or C18:0. These results show that ATGL is able to hydrolyze various TAG-
substrates, even if they contain unsaturated FAs. In agreement, FA-composition analysis of plasma-
FAs and of WAT-TAG from ATGLko and wt mice indicated that ATGL hydrolyzes all major saturated
and unsaturated FA species. Highest differences in FA composition of WAT-TAG between wt and
ATGLko mice were found for unsaturated FAs, in particular for C16:1 and C18:1. The observed FA
preference for length and degree of saturation of ATGL might be a direct result of its
stereo/regioselectivity for TAG hydrolysis, because early studies of Brockerhoff et al. showed that in
WAT-TAG more than 50% of total C16:1 are located at the sn-2 position and more than 75% of all FAs
at sn-2 position of TAG are unsaturated FA species [261]. Thus, it is conceivable that ATGL-mediated
hydrolysis of TAG releases preferentially unsaturated FAs because they are very abundant at sn-2
position of TAG and not because ATGL preferentially cleaves unsaturated FA at the sn-2 position.
Discussion
69
In addition to the FA-composition of WAT-TAG, also the FA-composition of circulating lipid species
was investigated. Upon fasting, WAT lipolysis releases FAs into the circulation. A large proportion of
FAs is further utilized for VLDL-TAG synthesis within hepatocytes. Thus, the FA-selectivity of ATGL in
adipose tissue lipolysis may indirectly influence the FA-composition of circulating VLDL-TAG. If the FA
release of WAT is blunted, such as in ATGLko mice, then also the plasma levels of VLDL-TAG are
drastically reduced. Furthermore, FA-composition analysis revealed that, besides an overall decrease
of plasma FAs in ATGLko mice, especially unsaturated FAs like C16:1 and C18:1 were largely
diminished in plasma FAs and VLDL-TAG. This goes along with the FA-preference of ATGL observed
during in vitro experiments since ATGL showed highest activity for C16:1 and C18:1 esters. The direct
involvement of hepatic ATGL in VLDL production is unclear, since liver-specific ATGLko mice show
drastic accumulation of TAG in cytosolic LDs but no alterations in circulating VLDL-TAGs [262].
The FA preference of ATGL is further relevant in the perspective that FAs are known ligands involved
in intracellular signaling, e.g. via activation of PPARs [211, 212]. Interestingly, unsaturated, but not
saturated FAs have strong signaling potential, because they exhibit high activation potential for
PPARs [211]. MUFAs, like C16:1 and C18:1, which are preferentially hydrolyzed by ATGL, are potent
activators for PPARα. In contrast, activation potential of MUFAs is much less for PPARβ or PPARγ
[211-213]. Studies regarding PPAR-activation of PUFAs, like C18:2 and C18:3, showed that PUFAs can
activate all PPAR species to similar extends [211, 212]. PUFAs in plasma and WAT TAG were not
determined in this study since they are not very abundant and were most of the times below
detection limit of the method. However, since unsaturated FAs are preferentially release by ATGL-
mediated TAG hydrolysis it seems likely that these FA species may in part also act as ligands and
facilitate PPAR signaling. Thus, if ATGL-mediated TAG breakdown is defective (e.g. ATGLko), the
impaired MUFA release may be causative for reduced PPARα activation as observed in ATGLko
animal model [217].
Additionally, the hydrolysis of specific FA species could induce the synthesis of bioactive molecules,
e.g. ceramides. Ceramide synthesis highly depends on C16:0 supply and ceramide levels are accepted
to be tightly connected to alterations in aPKC-dependent signaling. It is known that ceramides act as
ligand and activator of e.g. PKCζ which leads to an inhibition of PKB/Akt and consequently disturbed
insulin response and glucose uptake [230]. The fact that C16:0 levels are largely unaltered in ATGLko
animals and that ATGL exhibits no specific preference for C16:0 in vitro suggests that ATGL is not
directly involved in the provision of C16:0. Thus, ATGL may not directly affect intracellular ceramide
levels.
Discussion
70
Intracellular LDs are formed by a core of hydrophobic TAGs surrounded by a monolayer of
amphipathic PLs. This serves as mediator between TAG and the aqueous cellular environment. In
agreement with previous studies, which reported PLA2 activity for ATGL [5], this study demonstrates
that ATGL is able to hydrolyze PLs. Testing a variety of common PLs, it was evident that ATGL
hydrolyzes all investigated PL species, even though activities against PLs are much smaller than
against TAG. Interestingly, the rate of hydrolysis of micellar PLs by ATGL could not be co-activated by
CGI-58. The finding that ATGL can hydrolyze PLs independent of the co-activator CGI-58 raises the
questions about the metabolic implications of this hydrolytic activity. From results of this study, it
can be deduced that ATGL can hydrolyze PLs if presented in micellar form (with no hydrophobic core)
as well as in liposomal form, surrounding other neutral lipids. The latter liposomal substrate may
more reflect cellular LDs. Interestingly, addition of CGI-58 to ATGL led to an increase of ATGL´s
phospholipase activity, when TAG but not CE liposomes were used as substrate. The mechanism
underlying this substrate-dependent effect is so far unclear.
The phospholipase activity of ATGL as well as the positional selectivity against TAG is consistent with
enzymatic activities of other PNPLA protein family members. Next to ATGL, this protein family also
includes characterized sn-1 and sn-2 specific phospholipases (PNPLA8/9) and sn-1 specific lyso-
phospholipases (PNPLA6/7) [263-266]. The observed similarities in stereo/regioselectivity suggest a
common ancestry of these enzymes. To date it is unknown whether PNPLA6-9 contribute directly to
lipid turnover on cytoplasmic LDs. The fact that phospholipase activity of ATGL is much weaker than
TAG-hydrolase activity could match the abundance of the two substrates at LDs. Since PLs form the
surface of LDs, the molar amount of PLs is much smaller than that of TAG of the core. An explanation
why ATGL also exhibits phospholipase activity could be that the enzyme gains better access to the
hydrophobic TAGs by hydrolyzing PLs of the LD monolayer. Another explanation might be a spatial
consideration that during LD degradation also PLs must by hydrolyzed since LDs loose TAG and
become smaller. This may also implicate an explanation for the CGI-58-dependent enhancement of
ATGL´s phospholipase activity when TAG liposomes were used. Since TAG-hydrolysis is the main
activity of ATGL a supply of both TAG and PLs yields primary in TAG-degradation. Consequently the
size of used liposomes decreases, thus, surface area increases and provides potentially more PL
substrate, thereby enhances ATGL´s activity against PLs. The phospholipase activity of ATGL also
raises the possibility that in the process of lipolysis, lipid intermediates, such as arachidonic acid (a
very abundant FA at the sn-2 of PLs) and lyso-PLs, which could act as signaling molecules in cells, are
generated. In any case, further studies are necessary to confirm and clarify the phospholipase activity
of ATGL and its cellular function.
Discussion
71
The stereochemical analysis of DAG, which are formed by ATGL hydrolysis of TAG showed that ATGL-
mediated catabolism of TAG generates sn-1,3 and sn-2,3 DAGs. Interestingly, the stereochemistry of
DAG isoforms, which derive from ATGL catabolism is also corroborated by the accumulation of sn-1,3
DAG (65% of total DAG) in WAT of HSLko mice. In addition to sn-1,3 DAG, HSLko mice also
accumulate sn-2,3 DAG, constituting approximately 25% of the residual DAG. Interestingly, these
DAG isoforms are apparently not the “right” stereochemical species, to act as ligand for PKC [152-
154]. This conclusion is supported by the phenotype of HSLko mice. These mice do not develop a
clear defect in insulin response, despite massive accumulation of DAGs in many tissues, including SM
[56, 59]. In this regard the genetic background of the HSLko strains may play a role. Some strains
show indications for impaired insulin signaling [199, 200], whereas other, including the HSLko strain
used in this study, showed the opposite, increased hepatic insulin sensitivity [197, 198]. So far, this
discrepancy is entirely unsolved. Results obtained in this study show no indications of impaired
insulin sensitivity/signaling. In contrast, insulin tolerance tests indicated rather increased insulin
sensitivity in HSLko mice as compared to wt littermates. Similarly, Voshol et al. [197] and Park et al.
[198] monitored increased hepatic insulin sensitivity during hyperinsulinemic-euglycemic clamp
studies. In contrast, both clamp studies of Mulder et al. [199] and glucose tolerance tests of Roduit et
al. [200] suggest a decrease in insulin sensitivity. Additionally, results of this study show that the
expression levels of SM and liver specific protein kinase C isoforms, PKCθ and PKCε, respectively,
were tentatively decreased in HFD-fed HSLko mice as compared to wt animals. Both PKC isoforms
target IRS proteins, one of the initial events in insulin signaling. PKCθ can directly phosphorylate IRS1
on Ser1101 and Ser307 and PKCε increases IRS1 phosphorylation at Ser636/639 [230]. Such serine
phosphorylations of IRS1 are described to prevent insulin-induced IRS1 tyrosine phosphorylation,
which leads to inhibition of PI3K activity and PKB/Akt phosphorylation and hence blocks insulin-
induced cellular glucose uptake [230]. Since both PKC isoforms are described to negatively regulate
insulin signaling [177] these data indicate that in the present study, wt mice are more prone to HFD-
induced IR than HSLko mice. All in all, data of this study give no indication that DAGs, which
accumulate due to HSL deficiency, lead to an impairment of insulin signaling. Additionally, the fact
that TAG-derived DAGs are sn-1,3 and sn-2,3 isoforms and are generated at LDs and, thus, may never
end up at the plasma membrane where PKC resides also argues against a role of lipolysis-derived
DAGs in PKC signaling.
Intracellular DAGs can potentially undergo several anabolic reactions. HSL can catabolize them in the
process of lipolysis. DGKs can convert them to PA by phosphorylation of the free OH-group. CPT can
transform them to PC and the acyltransferases DGAT1 and DGAT2 can acylate them thereby forming
Discussion
72
TAG. CPT and DGKs represent membrane-bound enzymes. Hence, these reactions are unlikely to
occur on cytoplasmic LDs. Additionally, neither sn-1,3 DAG nor sn-2,3 DAG are substrates for these
enzymes (only sn-1,2 DAG is the intermediate of PL metabolism and glycerophospholipid de novo
synthesis). Both, the wrong stereochemistry of TAG-derived DAGs and the “mislocalization” on LDs
makes it unlikely that these DAGs are endogenous substrates for PL synthesis without prior
transesterification to sn-1,2 isomer and DAG transport systems. To exclude a misinterpretation of
obtained data, caused by DAG remodeling in vitro, transesterification of DAG was additionally
investigated. Results show that no conversion of rac-1,2/2,3 to sn-1,3 and vice versa was detectable.
Consistently, no mammalian DAG-transport proteins and no in vivo occurring transesterification of
DAG have been described yet. This suggests that the DAG pool, formed by ATGL, is not a precursor
for PL synthesis.
Another aim of this study was to determine the stereo/regioselectivity of the re-esterification of
DAG, which is catalyzed by DGAT1 and DGAT2. It turned out that DGAT2, but not DGAT1, exhibits
clear preference for sn-1,3 DAG. This finding was independent of the enzyme sources and confirmed
in acyltransferase assays using either cell homogenates containing DGAT1 or DGAT2 or homogenates
of murine WAT. DGAT1 converts rac-1,2/2,3 DAG much more efficiently than sn-1,3 DAG, whereas
DGAT2 exhibits the opposite substrate selectivity. These results were rather unexpected since the
role of DGATs on LDs has not been investigated so far. Until now, both DGAT enzymes are thought to
catalyze the same, final reaction of de novo TAG biosynthesis by esterifying sn-1,2 DAG with long-
chain FA-CoAs at the ER. The observed differences in substrate selectivity may be due to structural
differences of these two enzymes. Both belong to unrelated protein families. DGAT1 is a member of
the ACAT/DGAT1 gene family which is a subfamily of the MBOAT superfamily, whereas DGAT2
belongs to the DGAT2 gene family [126]. These two enzymes show no homology and it is known that
they cannot functionally compensate for each other [140, 267]. To date, the reaction mechanism
behind the observed metabolic differences, apart from structural differences, is elusive. Both DGATs
localize to the ER but only DGAT2 has been also reported to be associated within cytosolic LDs [139,
268]. Our finding that DGAT2 prefers sn-1,3 DAG as acyl-acceptor, generated by ATGL on LDs, marks
a new difference with regard to biochemical properties of both DGAT enzymes. Furthermore, the
finding that DGAT2 localizes on LDs together with the preference for sn-1,3 DAG suggests that ATGL
and DGAT2 may act coordinately on LDs to facilitate TAG hydrolysis and re-esterification, thereby
remodeling the degree of saturation of sn-2 FA esters. If this holds true, two distinct DAG pools for
DGAT enzymes would exist (Fig. 43).
Discussion
73
Both DGAT enzymes would esterify DAG pool 1 consisting of sn-1,2 DAGs, generated during de novo
glycerolipid synthesis at the ER. On the other hand, a sn-1,3 DAGs (pool 2), generated by ATGL on
cytoplasmic LDs would serve as substrate for DGAT2, exclusively. Such a local separation as well as
stereochemical distinction between different DAG pools could explain the differences observed in
mice carrying a global DGAT1 or DGAT2 deficiency. In line, a recent study overexpressing either
DGAT1 or DGAT2 in McA-RH7777 cells described surprisingly different phenotypes [140]. Despite
higher in vitro acyltransferase-activity when overexpressing DGAT1, cells expressing DGAT2 show
significant higher TAG mass, as well as glycerol incorporation into cellular TAG moiety. Furthermore,
LDs of cells overexpressing DGAT1 were considerably smaller compared to LDs of DGAT2 expressing
cells [140]. Another recent publication suggested different substrate sources of DGATs in HepG2
hepatoma cells. Studies using stable isotope-labeled substrates show that DGAT1 mainly
incorporates exogenously FAs to glycerol, whereas DGAT2 primary uses endogenously synthesized
FAs for TAG generation [144]. In combination with the observations of this study, it can be concluded
that DGAT1 and DGAT2 exhibit different functions in cellular TAG synthesis as evident by their
different localization, substrate source, and stereo/regioselectivity.
The observation that high doses of MgCl2 (100 mM) can result in an inhibition of either DGAT2 alone
or DGAT1 and DGAT2 in in vitro acyltransferase experiments was rather unexpected. High doses of
MgCl2 (> 100 mM) are described to inhibit specifically DGAT2, which has been demonstrated in mice
lacking DGAT1 as well as in cells overexpressing DGAT2 [118]. Results of this study show that the
presence of 100 mM MgCl2 leads to an exclusive inhibition of DGAT2 when sn-1,2 DAG is used as
substrate. In contrast to the expected inhibition of DGAT2, inhibition of DGAT1 (~50%) is additionally
observable when sn-1,3 DAG is used as substrate. Thus, MgCl2-dependent inhibition of DGAT activity
depends on the involved enzyme and the supplied substrate isoform. A comparison with the former
mentioned study of Cases et al. [118] is difficult because they did not use cells overexpressing DGAT1
do confirm their assumed selectivity of inhibition. In addition, they exclusively used sn-1,2 DAG as
substrate and thus may have overlooked the finding that MgCl2 also inhibits DGAT1 when sn-1,3 is
used as acyl-acceptor. In summary, data indicate that MgCl2 can be used as specific inhibitor for
DGAT2 in experiments containing sn-1,2 DAG. In experiments using sn-1,3 DAG the additional
inhibition of DGAT1 can falsify results. So far, the the mechanism by which MgCl2 differentially
inhibits DGAT enzymes is unknown.
In summary, results of this study demonstrate that ATGL hydrolyzes TAG selectively at the sn-2
position, thereby generating sn-1,3 DAG in vitro and in vivo. Generated sn-1,3 DAG constitutes the
preferred substrate for further hydrolysis by HSL or re-esterification catalyzed by DGAT2.
Discussion
74
Furthermore, co-activation of ATGL by CGI-58 leads to enhanced hydrolytic activity and to the
expansion of stereo/regioselectivity to sn-1 but not sn-3 position, therby additionally generating sn-
2,3 DAG but not sn-1,2 DAG. Since ATGL-dependent TAG hydrolysis does not generate sn-1,2 DAG,
the isoform that is known to activate PKCs, it is unlikely that lipolysis-derived DAGs regulate PKC-
activity. The preference of ATGL for long-chain unsaturated FAs, which are known PPAR ligands,
suggests that ATGL activity may contribute to PPAR activation in that way. Interestingly, ATGL is also
able to hydrolyze a variety of PLs, though with low activity. Moreover, results obtained from HSLko
mice showed that these mice cumulate sn-1,3 and sn-2,3 DAGs that are selectively generated by
ATGL-dependent lipolysis. Furthermore studies on HSLko mice suggest that accumulation of TAG-
derived DAG is not crucially involved in disturbed insulin signaling.
FIGURE 43. “3-pool” model of intracellular DAG distribution. Pool I consists of sn-1,2 DAG formed at the ER during de novo
lipogenesis. DGAT1, DGAT2 as well as CPT can metabolize newly generated sn-1,2 DAG. Pool II consists of sn-1,3 ± sn-2,3
DAGs, which are generated on cytoplasmic LDs by ATGL ± CGI-58-dependent TAG hydrolysis. Both DAG species are targets
for subsequent hydrolysis catalyzed by HSL or re-esterification catalyzed by DGAT2. sn-1,2 DAGs of pool III function as
activators of various PKCs and are generated by PLC at the plasma membrane. Furthermore, these DAGs are substrates for
DGKs and DAGLs. ATGL, adipose triglyceride lipase; CPT, choline:1,2-diacylglycerol cholinephosphotransferase; CGI-58,
comparative gene identification-58; DAG, diacylglycerol; DGK, diacylglycerol kinase; DAGL, diacylglycerol lipase; DGAT1/2,
diacylglycerol-O-acyltransferase 1/2; HSL, hormone-sensitive lipase; MAG, monoacylglycerol; PA, phosphatidic acid; PKC,
protein kinase C; PL, phospholipid; PLC, phospholipase C; TAG, triacylglycerol.
Observations of the present study in combination with previously published data support a “3-pool”
model of intracellular DAG compartmentation. Pool 1 is located at the ER and consists of sn-1,2 DAGs
that are generated during de novo lipogenesis by either MGATs or PAPases/lipins. This pool is
accessible and metabolized by DGAT enzymes and CPT (Fig. 43). TAG-derived DAGs form pool 2 at
Discussion
75
cytoplasmic LDs. This pool comprises sn-1,3 DAGs ± sn-2,3 DAGs and is generated by ATGL ± CGI-58.
DAGs of this pool are catabolized by HSL-mediated hydrolysis or re-esterified to TAG by DGAT2 (Fig
43). Pool 3 contains sn-1,2 DAGs, which are generated by PLC-dependent hydrolysis of PLs located at
the plasma membrane. DAGs of this pool can either serve as precursor for de novo synthesis of PL via
phosphorylation catalyzed by DGKs, or be degraded by membrane-bound DAGLs. So far, only sn-1,2
DAGs of pool 3 are widely accepted to be involved in cellular signaling events by activating novel and
conventional isoforms of PKC (Fig 43). The local separation of these pools within the cell as well as
the clear differences in stereo/regiochemistry of included DAGs may provide explanations to
previously confusing date regarding DAG-induced signaling and lipid metabolism.
Materials & Experimental Procedures
Materials & Experimental Procedures
77
I) Materials
Chemicals and buffers
Materials, chemicals and radiochemicals were obtained from either Sigma-Aldrich (St. Louis, MO) or
GE Healthcare (Waukesha, WI). Lipids were supplied by either Larodan Fine Chemicals (Malmō,
Sweden; TAGs), Sigma-Aldrich (DAGs) or Avanti Polar Lipids (Alabaster, AL; PLs).
Solution A: 0.25 M sucrose, 1 mM EDTA, 1 mM dithiothreitol, 20 µg/ml leupeptine, 2 µg/ml antipain
and 1 µg/ml pepstatin, pH of 7.0
Phosphate buffered saline (PBS): 137 mM NaCl, 2.7 mM KCl, 4.7 mM Na2HPO4, 1.4 mM KH2PO4, pH
7.4
Animals
Mice were housed on a regular light/dark cycle (12 h/12 h) and had ad libitum access to water and
normal CD (fat: 4.5%, protein: 22.1%, starch: 35.8%, sugar: 5.0%; w/w fat from Ssniff, Soest,
Germany) or HFD (fat: 34.0%, protein: 24.1%, starch: 1.1%, sugar: 23.8%; w/w fat from Ssniff, Soest,
Germany). 8-10 weeks-old male mice on CD or 20-22 weeks old male mice kept the last 12 weeks on
a HFD were used for studies. Mice with a global deletion of HSL (HSLko) or ATGL (ATGLko) were
generated by targeted homologous recombination and backcrossed to C57BL/6J at least 5 times, as
described previously [59, 75]. Blood and WAT of non-fasted and 8 h (12:00 – 08:00 am) fasted mice
on CD were used for lipid analyses. Tissue samples of liver and m. gastrocnemius (SM) of HFD-fed
mice fasted for 8 h (12:00 – 08:00 am), following 1 h of refeeding, were used for immunoblot
analysis. Body mass composition was assessed using TD-NMR minispec Live Mice Analyzer system
(LF90II, Bruker Optik GmbH, Ettlingen, Germany). Blood was collected via retro-orbital puncture from
isoflurane-anesthetized (Baxter, Deerfield, IL) mice. WAT, LIV, and SM samples were surgically
removed from cervically-dislocated mice. Tissue samples were washed in PBS, containing 1 mM
EDTA, 100 IU/ml heparin and disrupted using an Ultra Turrax® (IKA, Staufen, Germany), either in ice-
cold solution A for enzymatic activity measurements and immunoblotting or in methanol for further
lipid extraction and analysis. For analysis of brain lipids, mice were treated as described previously
[245].
Materials & Experimental Procedures
78
II) Experimental Procedures
cDNA cloning of recombinant proteins
pcDNA4/HisMaxC vector (Invitrogen, Carlsbad, CA) constructs containing the entire open reading
frame of murine ATGL, murine CGI-58, and murine HSL were generated as described previously [70].
Constructs of flag-tagged DGAT1 and DGAT2 were kindly provided by Robert V. Farese Jr. (Gladstone
Institute of Cardiovascular Disease, and Departments of Medicine and Biochemistry & Biophysics,
University of California, SF) and generated as described [140]. As control, pcDNA4/HisMaxC
containing β-galactosidase was used (Invitrogen). For purification of CGI-58, respective coding
sequence was subcloned into pYex4T-1 vector (CLONTECH Laboratories Inc., Mountain View, CA) as
previously described [68].
Purification of GST-tagged CGI-58
GST-tagged CGI-58 was expressed in S. cerevisiae BY4742 strain. Affinity purification of the GST-
fusion protein was performed as described [68]. pYex4T-1 vector was transformed into the S.
cerevisiae BY4742 strain. Copper promoter-driven expression of GST-CGI was induced by raising S.
cerevisiae in YNB-urea media containing 0.5 mM CuSO4. Cells were harvested and protoplasts were
generated using zymolyase. Protoplasts were disrupted by sonication using a Virsonic 475 (Virtis,
Gardiner, NJ) in the presence of 0.2% NP-40. GST-tagged CGI-58 within the supernatant was purified
using Glutathione-Sepharose beads (GE Healthcare) and further dialyzed overnight with 150 mM KCl,
10 mM potassium phosphate buffer (pH 7.0), and 0.01% NP-40.
Expression of recombinant proteins
SV-40 transformed monkey embryonic kidney cells (Cos7; ATCC, CRL-1651) were cultivated in
Dulbecco´s modified eagle medium (DMEM; GIBCO, Invitrogen) containing 10% fetal calf serum (FCS)
supplemented with penicillin (100 IU/ml, GIBCO) and streptomycin (100 µg/ml, GIBCO) at standard
conditions (95% humidified atmosphere, 37°C, 7.5% CO2). Transfection of cloned constructs was
performed using 1 µg DNA complexed with Metafectene (Biontex, Munich, Germany) in FCS-free
medium. After 4 h media were changed to DMEM containing 10% FCS. Finally, after 48 h cells were
washed twice in PBS and collected using a cell scraper.
Materials & Experimental Procedures
79
Preparation of tissue and cell homogenates
Cells were disrupted in solution A by sonication (3x6 sec on ice, 20% output; Virsonic 475). Both, cell
and tissue homogenates were centrifuged to remove nuclei and unbroken cells (1,000 x g, 4°C, 30
min). For microsomal-free cytoplasmic fraction, homogenates were further centrifuged (100,000 x g,
4°C, 60 min). The microsomal pellet was resuspended in ice-cold solution A to obtain microsomal
fraction.
Protein determination
Protein concentrations of homogenates and prepared cellular fractions were determined using Bio-
Rad protein assay reagent (BioRad laboratories, Hercules, CA) and bovine serum albumin (BSA) as
standard.
Immunoblotting
Cos7-cell homogenates (20 µg protein) or WAT microsomal fraction (30 µg protein) or microsomal or
cytosolic fraction of LIV and SM samples (40 µg protein) were separated according to their molecular
weight by sodium-dodecylsulfate-polyacrylamide gel electrophoresis (SDS-PAGE) using a 10%
polyacrylamide gel and tris/glycine (1.6 M glycine, 0.8% SDS, 200 mM Tris-HCl, pH 8.3) as
electrophoresis buffer. Then, proteins were transferred onto polyvenylidenfluorid (PVDF) membrane
(Carl Roth GmbH, Karlsruhe, Germany) using CAPS buffer (1 mM 3-cyclohexylamino-1-propansulfonic
acid). Unspecific binding sites were blocked using 10% non-fat dry milk (Carl Roth) dissolved in TST
buffer (0.15 M NaCl, 0.1% tween-20 v/v, 50 mM Tris-HCl, pH 7.4) for 2 h. His- or Flag-tagged proteins
(His-tag: ATGL, CGI-58, β-galaktosidase (LacZ) and HSL; Flag-tag: DGAT1 and DGAT2) were detected
by hybridization with either primary anti-His (GE-Healthcare) or anti-Flag antibodies (1:5000, 2% milk
in TST buffer), following anti-mouse horseradish-peroxidase (HRP)-linked secondary antibody (GE
Healthcare) (1:10000, 2% milk in TST buffer). Endogenous expression of DGAT1 and DGAT2 in murine
WAT was detected using polyclonal DGAT1 or DGAT2 primary antibody (ProSci, Poway, CA) (1:1000,
2% milk in TST buffer) and anti-rabbit HRP-linked secondary antibody (Vector laboratories Inc.,
Burlingame, CA) (1:10000, 2% milk in TST buffer). Endogenous expression of PKCα, PKCε, PKCθ,
GapDH, and HSL in LIV or SM samples was detected using respective primary antibodies (Cell
Signaling, Danvers, MA, USA) (1:1000, 2% milk in TST buffer) and anti-rabbit HRP-linked secondary
Materials & Experimental Procedures
80
antibody. Chemiluminescence was induced by ECLplus Kit (GE Healthcare) and detected by exposure
to X-ray films (Hyperfilm™ ECL, GE Healthcare).
Total lipid extraction and separation
Lipids of homogenized tissue samples (WAT, brain [245], or blood) were extracted using
chloroform/methanol/water (2/1/0,6; v/v/v; 1 % acetic acid). For GC-FID measurements, extracts
were supplemented with 20 µg margaric acid (C17:0) and 20 µg trimargarinate per ml as internal
standard. Extraction was performed under steady shaking for 1 h at room temperature (RT). After
centrifugation (1,000 x g, 4°C, 15 min), the organic phase was collected, dried under nitrogen, and
dissolved in 500 μl chloroform. Aliquots (30 μl) were separated by TLC using silica-gel coated plastic
TLC plates (solvent for TAG and FFA separation: hexane/diethyl ether/acetic acid (70/29/1; v/v/v) and
solvent for DAG separation: chloroform/acetone/acetic acid (95/4/1; v/v/v)). Lipids on TLC plates
were stained using iodine vapor. Bands corresponding to selected lipid species were scraped off and
either dissolved in 500 μl toluene, containing 500 μg butylated hydroxytoluene (1 mM in toluene) as
antioxidant and used for determination of FA composition (GC-FID) or lipids were re-extracted with
chloroform and used for HPLC lipid analysis (see below).
Fatty acid determination by gaschromatopgraphy with flame-ionization
detection
FA species were analyzed by GC-FID according to Sattler et al. [269] with some modifications: For
transesterification, 2 ml of borone trifluorid (BF3) were added to lipids, dissolved in 500 µl toluene,
and incubated for 1 h at 110°C in an incubator. Reactions were stopped by addition of 1 ml ice-cold
H2O. Resulting fatty acid methylesters (FAMEs) were extracted twice by addition of 2ml of
hexane/chloroform (4:1, v/v) and shaking for 10 min at RT. After centrifugation (1,000 x g, RT, 10
min) the upper-phase was collected. The combined phases were evaporated under nitrogen and
FAMEs were dissolved in 100 µl of hexane. The GC conditions were set to: split injection (split flow:
15 ml/min, split ratio: 1/5, injection volume: 2 µl), using an injector temperature of 230 °C, and a
wall-coated open tubular fused silica column (25 m, 0.32 mm inner diameter, FFA phase coated, film
thickness = 0.3, Agilent technologies, Santa Clara, CA). The carrier gas consisted of helium. As a
temperature gradient two consecutive ramps from 150 to 260 °C (ramp 1: 5°C/min to 250°C hold for
2 min, ramp 2: 10°C/min to 260 hold for 5 min) were used. FID (Trace-GC 2000series, ThermoQuest
corp., Atlanta, GA) conditions were as follows: base temperature 150°C; gas flows: 200 ml/min air, 30
Materials & Experimental Procedures
81
ml/min hydrogen, and 20ml/min helium. Data acquisition and analysis was done with Xcalibur 2.0
software (Thermo Fisher Scientific, Waltham, MA). For quantitative analysis the corresponding peaks
of FAMEs were integrated and peak areas were calculated using C17:0 as internal standard. FAME
concentrations were calculated as percentage of total FAMEs in a given sample and/or as amounts
per wet tissue weight (nmol/g) or plasma volume (nmol/l).
Enzymatic generation of DAG
The triolein substrate (0.3 mM final concentration in the used buffer) containing no tracer or either
1.0 x 106 cpm 3H-FA [9,10] labeled triolein or 0.5 x 106 cpm 14C-glycerol labeled triolein as tracer (per
sample) and PC (45 µM final concentration in the used buffer) were emulsified on ice in 100 mM
potassium phosphate buffer (pH 7.0) by sonication using a sonicator (3x60 sec on ice, 20% output;
Virsonic 475). Then, substrates were adjusted to 2% bovine serum albumin (BSA; FA free) and 100 µl
of substrate were incubated with cytoplasmic fractions of Cos7-cell lysates overexpressing ATGL (50
µg of protein in 100 µl solution A) without or with 200 ng of purified GST-tagged CGI-58 (in 10 mM
potassium phosphate buffer (pH 7.0), 0.01% NP-40) and HSL-specific inhibitor (12.5 µM of 76-0079 in
DMSO; Novo Nordisk, Kopenhagen, Denmark) in a water bath at 37°C for different periods (40 min,
60 min, or 120 min). The reaction was terminated by extracting the lipids according to Folch et al.
[270] using 1 ml chloroform/methanol (2/1; v/v). Subsequently, lipids were separated by TLC using
chloroform/acetone/acetic acid (95/4/1, v/v/v) as solvent. Bands corresponding to DAG were
scraped off and either re-extracted using chloroform and used for further analysis or radioactivity
was determined by liquid scintillation counting (Tri-Carb 2300 TR; Packard, Meridan, CT).
Determination of hydrolase activity
Different TAG (different FA composition) or diolein (sn-1,2; sn-1,3 or rac-1,2/2,3) substrates were
prepared by emulsification (sonication) with di-C18:1 PC (45 µM final concentration in the used
buffer) in 100 mM potassium phosphate buffer (pH 7.0) on ice using a sonicator (3x60 sec on ice, 20%
output; Virsonic 475). Final concentration was set to 0.25 mM for TAG substrates and to 0.3 mM for
DAG substrates using 100 mM potassium phosphate buffer (pH 7.0). Then, substrates were adjusted
to 2% bovine serum albumin (BSA; FA free). Cos7-cell homogenates (50 µg protein) containing either
HSL or ATGL were incubated without or with cell homogenates containing CGI-58 (50 µg protein) to
give a total volume of 100 µl in solution A to which 100 µl of indicated substrate was added and
incubated for 60 min in a water bath at 37°C. Then 200 µl of 0.1% Triton X-100 were added and
Materials & Experimental Procedures
82
samples were agitated for 10 min at RT. Released FAs were determined enzymatically using NEFA-C
kit (Principle: Activation of FA by acyl-CoA synthetase; Oxidation of FA-CoA by acyl-CoA oxidase;
Peroxidase dependent colour reaction; Wako chemicals, Neuss, Germany) after 10 min of shaking at
RT.
Determination of DAG rearrangement/transesterification
Racemic diolein (mixture of sn-1,2/sn-2,3/sn-1,3 or rac-1,2/2,3) or either sn-1,3 or sn-1,2 diolein
substrate (0.3 mM final concentration in the used buffer) was prepared by emulsification (sonication)
with PC (45 µM final concentration in the used buffer) in 100 mM potassium phosphate buffer (pH
7.0) on ice using a sonicator (3x60 sec on ice, 20% output; Virsonic 475). Then, substrates were
adjusted to 2% bovine serum albumin (BSA; FA free). 100 µl of substrate were incubated either with
100 µg cytosolic protein or 100 µg lysate protein of Cos7-cells in the presence of a HSL-specific
inhibitor (12.5 µM of 76-0079 in DMSO) (total volume: 100 µl in solution A) in a water bath at 37 °C
for 120 min. Before and after incubation, lipids were extracted according to Folch et al [270] by
adding 1 ml chloroform/methanol (2/1; v/v) and separated by TLC (silica gel-coated plastic TLC
plates) using chloroform/acetone/acetic acid (95/4/1, v/v/v) as solvent. Bands corresponding to DAG
were either stained using iodine vapor or scraped of, re-extracted with 10 ml chloroform, dried
under nitrogen and used for chiral analysis.
Determination of DGAT activity
WAT cellular fraction (microsomal, cytosolic, lipid droplet) or Cos7-cell homogenates containing
either DGAT1 or DGAT2 (50 µg protein) were incubated with HSL-specific inhibitor (12,5 µM of 76-
0079 in DMSO), tetrahydrolipstatin/orlistat (20 µM in DMSO, xenical®), niacin (5 mM in water), MgCl2
(100 mM in water), bromoenol lactone (BEL, 5 µM in DMSO, [271]), or DGAT1-specific inhibitor (5 µM
of (2-((1s,4s)-4-(4-(4-amino-7,7-dimethyl-7H-pyrimido[4,5-b][1,4]oxazin-6-yl)phenyl)cyclohexyl)
acetic acid [139] in DMSO) and different diolein substrates (final volume: 100 µl). Substrates were
composed of different diolein isoforms (0.2 mM of sn-1,2, sn-1,3 or rac-1,2/2,3 final concentration in
the used buffer) in Tris-buffer (50 mM, 20 mM MgCl2, pH 7.4) and either 0.8 mM (DAG/PC ratio:
0.25), 0.2 mM (DAG/PC ratio: 1) or 0.05 mM (DAG/PC ratio: 4) PC (final concentration in the used
buffer) and were emulsified on ice by sonication (3x60 sec on ice, 20% output; Virsonic 475).
Subsequently, C18:1-CoA (30 µM final concentration within substrate) and 14C-labeled C18:1-CoA (55
µCi/µmol; 20 µM final concentration within substrate) were added. Substrate was mixed 1:1 with
Materials & Experimental Procedures
83
samples to give a final volume of 200µl and incubated for 10 min at RT. Reactions were stopped and
lipids extracted by addition of chloroform/methanol (2:1, v/v) and separated on TLC (silica gel-coated
plastic TLC plates) using hexane/diethylether/acetic acid (70/29/1, v/v/v) as solvent. Bands
corresponding to TAG were scraped off and radioactivity was determined by liquid scintillation
counting (Tri-Carb 2300 TR).
Acylglycerol determination
Acylglycerol content (TAG, DAG) of lipid extracts from homogenized tissue preparations (WAT, brain
[245]), plasma as well as different non-radiolabeled TAG substrates were determined enzymatically
using INFINITY Triglyceride kit (Principle: Lipase-dependent hydrolysis of acylglycerol to glycerol;
Phosphorylation of glycerol by glycerol kinase; Oxidation of G3P by glycerolphosphate oxidase;
Colour reaction catalyzed by peroxidase using hydrogen peroxide; Thermo Scientific Fisher,
Middletown, VA). Plasma lipids or lipids extracted from TLC bands were measured directly, whereas
TAG substrates where diluted 1:1 with Triton X-100 to give a final concentration of 0.05 % Triton X-
100. Samples were measured photometrical after 10 min of shaking at RT.
Enzymatic hydrolysis of lecithin
Five mg of lecithin (egg yolk, average molecular weight ~800 g/mol) were emulsified in 10 ml of
potassium-phosphate buffer (100 mM, pH 7.4) by sonication (3x60 sec on ice, 20% output; Virsonic
475). Then, emulsion was adjusted to 2% bovine serum albumin (BSA; FA free). Lipid emulsion (500
µl, ~0.7 mM) was incubated with 100 µl of recombinant PLC (B. cereus, 100 IU/ml) for 2 h at 37°C.
Lipids were extracted according to Folch et al. [270] and separated by TLC using
chloroform/acetone/acetic acid (95/4/1, v/v/v) as solvent. Bands corresponding to DAG were
scraped off, re-extracted using chloroform, and subjected to chiral-phase HPLC analysis.
Determination of DAG isomers by chiral-phase HPLC
DAGs obtained either from rearrangement/transesterification experiments, of WAT tissue extracts,
or from TAG hydrolase assays were dried under nitrogen and derivatized according to Itabashi et al.
[272] with slight modifications. In detail, dried DAGs were dissolved in 400 µl dry toluol, 40 µl
pyridine and 2 mg 3,5-dinitrophenylisocyanate were added. The reaction mixture was shaken for 1 h
Materials & Experimental Procedures
84
at RT, dried under nitrogen, resolved in chloroform, and reaction products were separated by TLC
using UV254 silica plates and chloroform/methanol (95/5, v/v) as solvent. Bands corresponding to 3,5-
dinitrophenyl-urethanes were visualized under UV-light, scraped off, and extracted twice in
chloroform. The solvent was evaporated, samples were dissolved in 100 µl
hexane/dichloroethane/ethanol (80/20/2, v/v/v), and subjected to HPLC analysis. The HPLC system
consisted of Waters alliance e2695 separation module (Millford, MA) and an UV/VIS 2489 detector.
For separation a YMC-Pack A-K03 column (YMC Inc, Kyoto, Japan) containing (R)-(+)-1-(1-
naphtyl)ethylamine as stationary phase was used. Samples were analyzed under isocratic conditions
using hexane/dichloroethane/ethanol (80/20/2, v/v/v; flow rate: 1 ml/min) as mobile phase. DAG
derivatives were detected at 226 nm using an UV/VIS 2489 detector (Waters). Peaks corresponding
to DAG species were integrated and expressed as percentage of total DAG using Empower pro
software (Waters).
Intraperetoneal insulin tolerance test (IPITT)
Mice were fasted for 2 h and anestethized using isoflurane following an intraperetoneal bolus of 0.6
IU/kg bodyweight bovine insulin. Blood glucose levels were determined at indicated time points
using an Accu-Check glucometer (Roche Diagnostics, Basel, Switzerland). For every measurement
fresh blood was sampled by tail cutting. Area under the curve was calculated from absolute glucose
values using trapezoid method.
Determination of phospholipase activity
250 µM of 14C-labeled or non-radiolabeled PC was used as substrate for phospholipase activity
analysis. For determination of PL species selectivity, 500 µM non-labeled PC, PS, PE, PG, or PA was
used as substrates. For determination of phospholipase activity against mixed micelles either 50 µM
of 14C-labelled PC and 300 µM of 3H-labelled triolein or 50 µM non-labelled PC and 300 µM
cholesterylpalmitate were used as substrates. All substrates were prepared by emulsification in 100
mM potassium phosphate buffer (pH 7.0) in the presence or absence of 2 mM Ca2+ on ice using a
sonicator (3x60 sec on ice, 20% output; Virsonic 475). Then, substrates were adjusted to 2% bovine
serum albumin (BSA; FA free). All used PLs contained two C18:1 residues. Either purified PLA2 (1 IU;
Naja mossambica m.) or 50 µg protein of Cos7-cell homogenates containing ATGL without or with
purified GST-tagged CGI-58 (total volume: 100 µl in solution A) were incubated with 100 µl of
Materials & Experimental Procedures
85
respective substrate in a water bath at 37 °C for 60 min. Resulting FAs were determined either by
NEFA-C kit (Wako chemicals) or by scintillation counting.
Visualization of DAG substrate size by laser scanning microscopy
One ml of DAG substrates emulsified with PC in the DAG/PC ratios, 0.25 (50 µM/200 µM), 1 (200
µM/200 µM), and 4 (800 µM/200 µM) were incubated with the neutral lipid-specific dye BodiPy®
558/568 C12 (BodiPy, 15 µg/ml, Invitrogen) for 20min at RT. Subsequently, imaging of fluorescently
labeled structures was performed on a Leica SP2 confocal microscope (Leica microsystems, Wetzlar,
Germany) using a ×100, NA 1.40 oil immersion objective. BodiPy fluorescence was exited at 514 nm
and detected at 570 nm.
Statistics
Data are shown as means ± S.D. Statistical significance between two groups was determined by
unpaired Student´s two-tailed t-test. Following levels of statistical significance were used: *, p<0.05;
**, p<0.01; ***, p<0.001.
Publications
Publications
87
First author
Studies on the substrate and stereo/regioselectivity of adipose triglyceride lipase, hormone-
sensitive lipase, and diacylglycerol-O-acyltransferases
Thomas O. Eichmann, Manju Kumari, Joel T. Haas, Robert V. Farese Jr., Robert Zimmermann, Achim
Lass, and Rudolf Zechner
J. Biol. Chem. 2012.
Adipose triglyceride lipase (ATGL) is rate-limiting for the initial step of triacylglycerol (TAG) hydrolysis,
generating diacylglycerol (DAG) and fatty acids (FAs). DAG exists in three stereochemical isoforms.
Here we show that ATGL exhibits a strong preference for the hydrolysis of long-chain FA esters at the
sn-2 position of the glycerol backbone. The selectivity of ATGL broadens to the sn-1 position upon
stimulation of the enzyme by its co-activator CGI-58. sn-1,3 DAG is the preferred substrate for the
consecutive hydrolysis by hormone-sensitive lipase (HSL). Interestingly, diacylglycerol-O-
acyltransferase 2 (DGAT2), present at the endoplasmic reticulum and on lipid droplets (LDs),
preferentially esterifies sn-1,3 DAG. This suggests that ATGL and DGAT2 act coordinately in the
hydrolysis/re-esterification cycle of TAGs on LDs. Since ATGL preferentially generates sn-1,3 and sn-
2,3 it suggests that TAG-derived DAG cannot directly enter glycerophospholipid synthesis or activate
protein kinase C without prior isomerization.
Publications
88
Co-author
FAT SIGNALS - Lipases and Lipolysis in Lipid Metabolism and Signaling.
Zechner R, Zimmermann R, Eichmann TO, Kohlwein SD, Haemmerle G, Lass A, Madeo F.
Cell Metab. 2012 Mar 7;15(3):279-91.
Adipose triglyceride lipase affects triacylglycerol metabolism at brain barriers.
Etschmaier K, Becker T, Eichmann TO, Schweinzer C, Scholler M, Tam-Amersdorfer C, Poeckl M,
Schuligoi R, Kober A, Chirackal Manavalan AP, Rechberger GN, Streith IE, Zechner R, Zimmermann R,
Panzenboeck U.
J Neurochem. 2011 Dec;119(5):1016-28. doi: 10.1111/j.1471-4159.2011.07498.x. Epub 2011 Oct 20.
ATGL-mediated fat catabolism regulates cardiac mitochondrial function via PPAR-α and PGC-1.
Haemmerle G, Moustafa T, Woelkart G, Büttner S, Schmidt A, van de Weijer T, Hesselink M, Jaeger D,
Kienesberger PC, Zierler K, Schreiber R, Eichmann TO, Kolb D, Kotzbeck P, Schweiger M, Kumari M,
Eder S, Schoiswohl G, Wongsiriroj N, Pollak NM, Radner FP, Preiss-Landl K, Kolbe T, Rülicke T, Pieske
B, Trauner M, Lass A, Zimmermann R, Hoefler G, Cinti S, Kershaw EE, Schrauwen P, Madeo F, Mayer
B, Zechner R.
Nat Med. 2011 Aug 21;17(9):1076-85. doi: 10.1038/nm.2439.
Publications
89
Growth retardation, impaired triacylglycerol catabolism, hepatic steatosis, and lethal skin barrier
defect in mice lacking comparative gene identification-58 (CGI-58).
Radner FP, Streith IE, Schoiswohl G, Schweiger M, Kumari M, Eichmann TO, Rechberger G, Koefeler
HC, Eder S, Schauer S, Theussl HC, Preiss-Landl K, Lass A, Zimmermann R, Hoefler G, Zechner R,
Haemmerle G.
J Biol Chem. 2010 Mar 5;285(10):7300-11. Epub 2009 Dec 18.
Adipose triglyceride lipase plays a key role in the supply of the working muscle with fatty acids.
Schoiswohl G, Schweiger M, Schreiber R, Gorkiewicz G, Preiss-Landl K, Taschler U, Zierler KA, Radner
FP, Eichmann TO, Kienesberger PC, Eder S, Lass A, Haemmerle G, Alsted TJ, Kiens B, Hoefler G,
Zechner R, Zimmermann R.
J Lipid Res. 2010 Mar;51(3):490-9. Epub 2009 Nov 25.
Neutral lipid storage disease: genetic disorders caused by mutations in adipose triglyceride
lipase/PNPLA2 or CGI-58/ABHD5.
Schweiger M, Lass A, Zimmermann R, Eichmann TO, Zechner R.
Am J Physiol Endocrinol Metab. 2009 Aug;297(2):E289-96. Epub 2009 Apr 28.
Appendix
Appendix
91
Abbreviations & Acronyms
2-AG 2-arachidonoyl glycerol
AA amino acid
AGPAT 1-acylglycerol-3-phosphate-O-acyltransferase
AMPK AMP-activated kinase
ASO antisense oligonucleotide
AT adipose tissue
ATGL adipose triglyceride lipase
BAT brown adipose tissue
BSA bovine serum albumin
BSSL bile-salt stimulated lipase
cAMP cyclic AMP
CD chow diet
CE cholesteryl ester
Ces carboxylesterase
CGI-58 comparative gene identification-58
CIP Cahn-Ingold-Prelog
CM cardiac muslce
CoA coenzyme A
CPT CDP-choline:1,2-diacylglycerol cholinephosphotransferases
DAG diacylglycerol
DAGL diacylglycerol lipase
DGAT diacylglycerol-o-acyltransferase
Appendix
92
DGK diacylglycerol kinase
EPT CDP-ethanolamine:1,2-diacylglycerol ethanolaminephosphotransferase
ER endoplasmic reticulum
ERK extracellular signal-regulated kinase
FA fatty acid
FAME fatty acid methylester
G0S2 G0/G1 switch gene 2
G3P glycerin 3-phosphate
GAPDH glycerinaldehyde 3-phosphate dehydrogenase
GC-FID gas chromatography with flame-ionization detector
GL gastric lipase
GLUT4 glucose transporter 4
GPAT glycerol-3-phosphate acyltransferase
GS2 gene sequence 2
HFD high fat diet
HSL hormone-sensitive lipase
IP3 inositol 1,4,5-triphosphate
IPITT intraperetoneal insulin tolerance test
IR insulin resistance
IRS insulin receptor substrate
LacZ β-galaktosidase
LD lipid droplet
LL lingual lipase
LPA lyso-phosphatidic acid
Appendix
93
LPAAT lyso-phosphatidic acid acyltransferase
LPL lipoprotein lipase
MAG monoacylglycerol
MBOAT membrane-bound O acyltransferases
MGAT monoacylglycerol acyltransferase
MGL monoglyceride lipase
mTOR mammalian target of rapamycin
MUFA mono-unsaturated fatty acid
PA phosphatidic acid
PAL pancreatic lipase
PAPase phosphatidic acid phosphatase
PBS phosphate-buffered saline
PC phosphatidyl choline
PDK 3-phosphoinositide-dependent protein kinase 1
PE phosphatidyl ethanolamine
PGC peroxisome proliferator-activated receptor gamma co-activator
PI3K phosphatidylinositide-3-kinase
PIP2 phosphatidylinositol 4,5-bisphosphate
PKA proteinkinase A
PKB/Akt protein kinase B
PKC, c/n/a protein kinase C, conventional/novel/atypical
PL glycerophospholipid
PLA phospholipase
PNPLA patatin-like phospholipase domain containing A
Appendix
94
PPAR peroxisome proliferator-activated receptor
PS phosphatidyl serine
PUFA poly-unsaturated fatty acid
RE retinylester
Rictor rapamycin-insensitive companion of mammalian target of rapamycin
SM skeletal muscle
sn stereospecific numbering
T2DM Type 2 diabetes mellitus
TAG triacylglycerol
TGH triacylglycerol hydrolase
TLC thin layer chromatography
WAT white adipose tissue
wt wild type
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