Lipid droplet growth and adipocyte development: mechanistically distinct processes connected by phospholipids

    Lipid Droplet Growth and Adipocyte Development: Mechanistically Distinct Processes Connected by Phospholipids Yanfei Qi, Lei Sun, Hongyuan Yang PII: DOI: Reference:

S1388-1981(17)30120-8 doi:10.1016/j.bbalip.2017.06.016 BBAMCB 58172

To appear in:

BBA - Molecular and Cell Biology of Lipids

Received date: Revised date: Accepted date:

6 May 2017 20 June 2017 23 June 2017

Please cite this article as: Yanfei Qi, Lei Sun, Hongyuan Yang, Lipid Droplet Growth and Adipocyte Development: Mechanistically Distinct Processes Connected by Phospholipids, BBA - Molecular and Cell Biology of Lipids (2017), doi:10.1016/j.bbalip.2017.06.016

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ACCEPTED MANUSCRIPT

Lipid Droplet Growth and Adipocyte Development: Mechanistically Distinct Processes

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Connected by Phospholipids

Yanfei Qi1, Lei Sun2 and Hongyuan Yang1, * School of Biotechnology and Biomolecular Sciences, The University of New South Wales, Sydney, NSW, 2052, Australia DUKE-NUS Graduate Medical School Singapore, 8 College Rd, Singapore, 169857

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Corresponding author: School of Biotechnology and Biomolecular Sciences, The University of New South Wales, Sydney NSW 2052 Australia Tel: 61-2-93858133 Fax: 61-2-93851483 E-mail address: [email protected]

Keywords: lipid droplets, seipin, lipin, CDP-DAG synthase, AGPAT2, BSCL2, Lipodystrophy.

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ACCEPTED MANUSCRIPT Abstract The differentiation of preadipocytes into mature adipocytes is accompanied by the growth and

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formation of a giant, unilocular lipid droplet (LD). Mechanistically however, LD growth and

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adipogenesis are two different processes. Recent studies have uncovered a number of proteins that are able to regulate both LD dynamics and adipogenesis, such as SEIPIN, LIPIN and CDP-Diacylglycerol Synthases. It appears that phospholipids play a critical role in both LD growth

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and adipocyte development. This review summarizes recent advances, and aims to provide a better

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understanding of LD growth as well as adipogenesis, two critical aspects in mammalian fat storage.

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ACCEPTED MANUSCRIPT Overview of lipid droplet biology Lipid storage has been recognized as a central issue in metabolic disorders, including insulin

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resistance, type 2 diabetes, non-alcoholic fatty liver disease, and cardiovascular diseases. In a

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challenging environment, living organisms need to store energy in times of plenty to supply energy when food becomes scarce [1, 2]. At the cellular level, energy is stored in form of neutral lipids within lipid droplets (LDs) [3, 4]. Neutral lipids in LDs can serve as the source of energy substrates,

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membrane components, hormone precursors and signaling messengers [3, 5]. In response to free

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fatty acid (FFA) overload, lipid storage in LDs also provides a means of protection from lipotoxicity

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[6, 7].

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LDs are evolutionarily conserved organelles present in nearly all organisms, from bacteria to

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mammals [8, 9], and in almost all eukaryotic cell types [10]. The content of neutral lipid core varies in different cell types. White adipocytes exclusively store triacyglycerols (TAGs), with little

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cholesterol esters (CEs); whereas CEs are predominant in macrophage LDs [11]. Yeast LDs contain a mixed pool of TAGs and CEs in almost equal amounts [12]. The size and number of LDs also vary in different cell types. For instance, white adipocytes are characterized by a giant, unilocular LD whereas brown adipocytes normally contain numerous small LDs.

LDs are believed to originate from the endoplasmic reticulum (ER), where most enzymes responsible for lipid synthesis reside. The prevailing model of LD biogenesis starts with LD nucleation, in which LDs emerge from lipid accumulation in spatially restricted subdomains within two leaflets of the ER membrane, followed by budding off into the cytoplasm[13, 14].

This model is based on two key 3

ACCEPTED MANUSCRIPT assumptions: 1. The ER membrane has limited capacity to allow storage lipids such as TAG to undergo free lateral diffusion; 2. Lipid/TAG globules would nucleate at discrete sites of the ER,

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triggering the budding and formation of nascent LDs and initial LDs (iLDs)[15]. Although attractive

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and widely accepted, this model still requires additional work to provide molecular details. For instance, the sites of LD nucleation remain to be clearly defined. Moreover, little is known about the protein machinery that drives LD budding, despite recent work implicating fat storage-inducing

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transmembrane protein 2 (FITM2) in LD biogenesis [16, 17]. The involvement of

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curvature-producing phospholipids (PLs) in LD budding also requires further analyses [5, 18].

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Once formed, the nascent initial LDs (with a diameter typically less than 200nm in cultured

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mammalian cells) will grow into mature iLDs (typically 300-500nM in diameter). This growth likely

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requires lipid/TAG transfer from ER to nascent iLDs through ER-LD contact sites. From birth, LDs are believed to retain connection with the ER, as manifested by their dynamic cargo exchange [15,

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19]. Therefore, some ER resident proteins can regulate LD growth. SEIPIN is an integral ER membrane protein. Cells deficient in mammalian SEIPIN or its yeast ortholog Fld1p/Sei1p exhibit altered LD morphology, with either clustering of multiple small LDs or formation of few strikingly enlarged, or supersized LDs [20-22]. Recent studies have demonstrated that SEIPIN/Fld1p is stably localized to ER-LD contact sites, facilitating the incorporation of protein and lipid cargo into growing LDs [23-25]. SEIPIN may also regulate the growth of iLDs by helping the formation and maintenance of the ER-LD contact sites [23, 25, 26].

Prolonged incubation of cells with fatty acids can trigger further iLD growth through an alternative 4

ACCEPTED MANUSCRIPT pathway: a subset of iLDs can acquire key enzymes in producing core neutral lipids and surface phospholipids, enabling their further growth into expanding LDs (eLDs, typically with a diameter

O-acyltransferase

2

(DGAT2)

for

TAG

synthesis,

as

well

as

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CoA:diacylglycerol

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over 1µM)[15]. These enzymes include glycerol-3-phosphate acyltransferase (GPAT3/4) and acyl

CTP:phosphocholine cytidylyltransferase 1 (CCT1), which binds to phosphatidylcholine (PC)-poor surfaces of LDs to synthesize PC. Through the actions of Arf1/COP-1, iLDs may undergo

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back-fusion with the ER and form membrane bridges, enabling the migration of GPAT4/DGAT2

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from ER to LD surface. However, the molecular mechanism of iLD back-fusion with the ER remains to be elucidated. Another means for LDs to grow is through a lipid transfer process mediated by cell

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death inducing DFF45-like effector (CIDE) family of proteins including CIDE A, B and C. CIDEC,

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also known as FSP27, is highly expressed in white adipocytes, and is required for the formation of

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the characteristic unilocular LDs in white adipocytes. Elegant work from Li and colleagues has demonstrated that CIDEC concentrates at LD-LD contact sites (MCS) and generates a “pore” that

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enables unidirectional transfer of neutral lipids from small to large LDs [27]. In the absence of CIDEC, white adipocytes accumulate multiple small LDs. Due to increased surface area for lipase access, CIDEC-deficient adipocytes exhibit a higher rate of lipolysis, which ultimately impairs the lipid storage capacity of adipose tissue in vivo [28]. CIDEA and CIDEB are found in brown adipocytes and hepatocytes, respectively, and are believed to mediate LD growth in those cells through a mechanism that is similar to CIDEC [28-30].

Depending on species examined, LDs have been shown to contain 40 to 300 different proteins, some of which are specific to LDs, while others are shared with the ER and possibly other organelles [3, 8]. 5

ACCEPTED MANUSCRIPT LD proteins regulate synthesis, degradation, transport, and sensing of lipids across PL monolayer, and mediate interconnection with other cellular processes [31]. Members of PLIN family are most

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extensively studied LD surface proteins. PLIN1 and PLIN2 are found as major protein constituents

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on larger LDs, whereas PLIN3 and PLIN4 are localized to smaller LDs [32]. This may be due to interactions of PLIN proteins with distinct PLs and proteins on LD of different sizes [33]. Li and her colleague have identified PLIN1 as active regulator of LD size by interacting with CIDEC and

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promoting unilocular LD formation in adipocytes [34]. During adipocyte lipolysis, PLIN1 on LD

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surface is phosphorylated by protein kinase A (PKA), causing dissociation of CGI-58 that in turn recruits adipose triglyceride lipase (ATGL) to LD surface. On the other hand, hormone sensitive

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lipase (HSL) is recruited by PLIN1 to LDs, where HSL is phosphorylated by PKA to ensure maximal

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lipolytic activity. ATGL hydrolyzes TAG into diacylglycerol (DAG), which is subsequently

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and FFAs [35-37].

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hydrolyzed into monoacylglycerol (MAG) by HSL, and MAG is eventually broken down to glycerol

Glycerolipid synthesis and lipid droplet growth Glycerolipids such as TAG, are synthesized from FFAs and glycerol-3-phosphate by enzymes in the ER. The first committed step of de novo TAG biosynthesis is the acylation of glycerol-3-phosphate by glycerol-3-phosphate acyltransferases (GPATs) to generate lysophosphatidic acid (LPA). An additional

acyl

chain

is

subsequently

transferred

to

LPA

by

members

of

the

1-acylglycerol-3-phosphate acyltransferases (AGPATs) family to produce phosphatidic acid (PA) (Figure 1). These two reactions take place in the ER. PA is sitting at a branching point in the synthesis of membrane phospholipids (PLs) and TAGs. In the following steps, PA can be converted 6

ACCEPTED MANUSCRIPT to diacylglycerol (DAG) through action of phosphatidate phosphatases (PAPs), referred as Lipin proteins in mammals or Pah1p in yeast. DAG is the key intermediate for synthesis of TAG as well as

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PLs, such as phosphatidylcholine (PC), phosphatidylethanolamine (PE) and phosphatidylserine (PS)

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through the Kennedy pathway in mammals. PA can also be converted to CDP-DAG through CDP-DAG synthases (CDSs), and CDP-DAG serves as precursor for the synthesis of

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phosphatidylinositol (PI), phosphatidylglycerol (PG) and cardiolipin (CL) [10, 31, 38].

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The neutral lipid core of LDs is coated by a monolayer of PLs. In mammalian LDs, PC and PE are the major PL species, followed by PI and PA; while in yeast LDs, PI is the second abundant PL

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species after PC [39, 40]. In addition neutral lipid synthesis, PL composition on LD surface is

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another key determinant of LD morphology. In general, limited PL synthesis leads to formation of

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supersized LDs, as it is thought that larger LDs minimize surface to volume ratio due to shortage in surface PLs, in particular PC [41, 42]. Genome-wide screening analyses have revealed that

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accumulation of PA or increased ratio of PE to PC on LD surface favors the formation of supersized LDs [41, 43]. The physical properties of lipids also influence LD dynamics: lipids with negative curvature, such as free sterols and PA, can increase surface tension and promote LD coalescence; in contrast, lipids with positive curvature, such as lyso-PA, may favor the budding and “pinching-off” of LDs [4, 18]. In this respect, four key enzymes regulating PA metabolism, i.e. GPAT3/4, AGPAT2, PAP/lipin and CDS1/2, have all been linked to LD growth/morphology (Figure 1)[21, 38, 43]. Reducing PAP or CDS activity led to the formation of supersized LDs in yeast and mammalian cells [38, 43]. Depleting AGPAT2, which is known to increase cellular PA, also increased LD clustering and the formation of supersized LDs (our unpublished observations)[44, 45]. Likewise, 7

ACCEPTED MANUSCRIPT overexpressing ER GPATs in yeast or mammalian cells increased PA and formed supersized LDs [21]. Notably, CIDEA was demonstrated to promote LD growth through PA [46]. Therefore,

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intermediates in glycerolipid synthesis such as LPA, PA and DAG may be crucial regulators of LD

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biogenesis and growth.

Overview of adipogenesis and lipodystrophy

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In mammals, adipose tissue is the primary depot of lipid storage [47]. The adipocytes in white

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adipose tissue (WAT) have unique capacity to store large amount of lipids, and the stored lipids can be released rapidly for the use by other tissues. In addition to fat storage, WAT has long been

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recognized as a bona fide endocrine organ that secrets adipokines which regulate systemic energy

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homeostasis [48]. In contrast, brown adipose tissue (BAT) primarily functions for heat generation

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and energy consumption [49]. Obesity is characterized by accumulation of fully-differentiated white adipocytes loaded with LDs. Classic LDs in white adipocytes is unilocular, taking up to 90% of

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cytoplasmic space and 60~85% of weight of cells. In contrast, brown adipocytes possess multilocular LDs, consistent with their thermogenic characteristics [3, 10, 47]. In response to over nutrition, WAT accommodates excessive lipids via both increase in size (hypertrophy) and number (hyperplasia) of adipocytes [50]. When the storage capacity of WAT is overwhelmed or compromised, excessive lipids are diverted to non-adipose tissues, such as liver, skeletal muscle and heart, where they cause lipotoxicity, leading to metabolic syndrome [7, 37, 51]. A widely held view is that ectopic lipids, rather than the “safe” fat stored in adipose tissue, drive the progression of metabolic disorders [52]. Indeed, in obese animal models, genetically manipulated expansion of adipose tissue mitigates the progression of metabolic defects [53]. 8

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Both white and brown adipocytes arise from embryonic mesoderm. White adipocytes stem from both

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MYF5+ and MYF5- lineages, whereas brown adipocytes derive from MYF5+ lineage only [54].

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Adipocyte differentiation can be divided into two phases: in the determination phase, mesenchymal stem cells responding to yet-to-be characterized cues first develop into preadipocytes, with no distinguished morphological alterations from their progenitor cells [55]; in terminal phase, i.e. from

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preadipocytes to fully differentiated mature adipocytes, adipogenic factors, such as insulin, cortisol

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and others initiate similar adipogenic transcriptional cascade in both white and brown adipocytes. Firstly, two members of the CCAAT/enhancer binding protein family, C/EBP-β and C/EBP-δ are

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induced, which in turn directly activate the “master” regulators of terminal adipogenesis, peroxisome

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proliferator activated receptor γ (PPARγ) and C/EBP-α. Once PPARγ and C/EBP-α are expressed,

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they cooperate to stimulate the expression of a large number of adipocyte-specific gene products such as fatty acid binding protein 4/ adipocyte protein 2 (FABP4/aP2), adiponectin/Adipoq, glucose

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transporter type 4 (GLUT4), and lipoprotein lipase (LPL) [7, 10, 47]. However, the exact mechanisms whereby adipogenesis is induced in vivo remain elusive. Although PPARγ and C/EBP-α co-regulate most genes during in vitro adipogenesis [56], these two master regulators share very few transcriptional targets in vivo [57]. C/EBPα is not required for the maturation of subcutaneous WAT (sWAT) during embryogenesis or epididymal WAT (eWAT) during early postnatal development. In contrast, they are PPARγ-dependent [57, 58]. PPARγ is a ligand-activated nuclear receptor. It has a large, promiscuous ligand-binding pocket and can be activated by a diverse spectrum of lipid species, including select leukotrienes, prostaglandins, oxidized or nitrated FAs, and oxidized PLs [55, 59, 60]. 15-prostaglandin-E2 can act as a ligand of PPARγ to enhance adipogenesis of 3T3-L1 cells [61]. In 9

ACCEPTED MANUSCRIPT contrast, another endogenous ligand, cyclic PA has been shown to inhibit PPARγ activity and block adipogenic

differentiation

of

3T3-L1

cells

[62].

Besides

naturally

occurring

ligands,

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thiazolidinediones (TZDs) are PPARγ agonists that stimulate adipocyte differentiation in vitro [63],

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but these drugs tend to divert murine and human white adipocyte precursors towards beige cell development [64, 65]. To date, many pro-adipogenic and anti-adipogenic regulators have been identified to associate with PPARγ-mediated transcriptional cascade. For instance, phosphorylation

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of PPARγ at serine 112 by MAP kinase represses PPARγ activity, leading to inhibition of

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adipogenesis [66].

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Lipodystrophy is an extreme lipid storage disease, which was first documented in medical literature

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about 130 years ago [67]. Opposite to obesity that manifests adipose tissue expansion, lipodystrophy

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is featured by selective loss of adipose tissue [67]. Partial and generalized lipodystrophy predispose to developing an array of complications, including insulin resistance, type 2 diabetes and acanthosis

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nigricans; hypertriglyceridemia, hepatic steatosis, and proteinuric kidney disease [51, 68]. The limited lipid storage capacity of adipose tissue underlies severe ectopic lipid deposition and associated metabolic disorders. The severity of metabolic syndrome is proportional to the extent of loss of adipose tissue [51]. Lipodystrophy can be classified as genetic and acquired syndrome in term of the causation. Two major types of genetic lipodystrophy observed in patients are: generalized loss of adipose tissue at birth or soon thereafter which is an autosomal recessive condition called congenital generalized lipodystrophy (CGL), also known as Berardinelli-Seip congenital lipodystrophy (BSCL), and partial loss of adipose tissue during late childhood and puberty which can be either autosomal recessive or dominant trait called familial partial lipodystrophy (FPL) [51]. By 10

ACCEPTED MANUSCRIPT far, there are 300~500 CGL cases and ~1,000 FPL cases documented, but it is assumed that undiagnosed cases are three times that of reported cases [69]. Studying the etiology of lipodystrophy

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will not only help develop novel therapies in the battle against this condition itself, but will also shed

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critical insight into the broader area of lipid storage diseases, e.g. obesity, and the underlying mechanisms. Both CGL and FPL are monogenic diseases, and so far nearly 20 disease loci have been identified [69]. The most common CGLs are caused by the mutations in CGL1/BSCL1/AGPAT2,

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CGL2/BSCL2/SEIPIN, CGL3/CAV1 and CGL4/PTRF; however, CGL1 and 2 are likely caused by

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defective adipogenesis whereas CGL3 and 4 are believed to arise from premature death of adipocytes [10]. FPLs arise from mutations in PPARG, LMNA, PLIN1, CIDEC/FSP27, LIPE/HSL, ZMPSTE24

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and AKT2 [1, 51], but the exact mechanism for localized fat loss varies from case to case [1]. In mice,

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loss of lipin-1 function also causes severe generalized lipodystrophy [70, 71]. This review will focus

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on recent advances in the studies of CGL, particularly BSCL2/SEIPIN because of its dual roles in regulating both LD biology and adipogenesis. The possible relationship between SEIPIN, GPAT,

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AGPAT2, lipin-1 and CDS1/2 will also be discussed.

Biochemical understanding of SEIPIN The most severe form of lipodystrophy is the type 2 CGL (CGL2; OMIM # 269700). It is an autosomal recessive disorder caused by loss-of-function mutations in the BSCL2 gene that is located at 11q13 on chromosome and encodes a protein called SEIPIN. Distinct from other types of CGL that preserve mechanical adipose tissue in the palms, soles, orbits, scalp, and periarticular regions, CGL2 patients lose mechanical fat along with metabolically active adipose tissues at most subcutaneous, intermuscular, bone marrow, intraabdominal, and intrathoracic regions [51, 72]. In 11

ACCEPTED MANUSCRIPT addition to the near complete lack of fat since birth, CGL2 patients also manifest mild mental

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retardation and cardiomyopathy [73].

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Magre et al first cloned human BSCL2 gene that encodes 398 amino acids (a.a.) protein with at least two hydrophobic stretches, and thus speculated that SEIPIN is a transmembrane protein [72]. Northern blot analyses showed that human BSCL2 is transcribed into at least three different mRNAs

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in lengths of 1.6, 1.8 or 2.2kb. The 1.8kb mRNA is exclusively expressed in all parts of brain,

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whereas the other two variants express ubiquitously [74, 75]. In 2004, a longer form (462a.a.) of human SEIPIN with additional 64aa at N-terminus was proposed based on the statistical predictions

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of the Kozak's consensus sequence. [76]. The 1.6kb mRNA can only be translated to 398a.a. protein,

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while 1.8 and 2.2bp mRNAs can be translated to both long and short protein isoforms [74, 75]. The

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462a.a. protein was suggested as predominant form of SEIPIN, rather than, as originally proposed, the 398a.a. isoform [75]. However, due to the lack of a sensitive and specific antibody against

determined.

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SEIPIN, the exact tissue distribution pattern as well as relative quantity remains to be firmly

The topology of SEIPIN was first investigated based on its yeast ortholog, few lipid droplets protein 1 (Fld1p)/Sei1p in the extensive analyses of 37 yeast membrane proteins with unknown functions [77]. Fld1p was predicted to possess two transmembrane domains, with both termini exposed to the cytosol [77]. In a later study, the long form of human SEIPIN has been shown to have both N and C termini facing the cytoplasm, with a long luminal loop between the two transmembrane helices, using engineered glycosylation sites [75]. Such a topology has been further confirmed by others [78]. 12

ACCEPTED MANUSCRIPT The first 280a.a. of human SEIPIN is 88% identical to the homologous regions in rodents and primates SEIPIN isoforms [76]. Although human SEIPIN and yeast Fld1p share striking

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conservations in secondary structure and membrane topology, they show weak sequence similarity.

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Fld1p only possess 12 to 16 a.a at N- and 11 to 19 a.a. at C- termini which are exposed to the cytosol, distinct from the long cytoplasmic C-terminal and isoform-specific N-terminal extensions in human SEIPIN (Figure 2) [20]. To date, all known missense mutation sites in CGL2 patients reside in the

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ER lumenal loop of SEIPIN such as A212P [72]; while two mutations of SEIPIN glycosylation sites,

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N88S and S90L (also lumenal) are associated with neuronal seipinopathy [74]. It should be noted that the membrane topology of SEIPIN/Fld1p/Sei1p remains to be firmly determined by structural

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studies.

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SEIPIN’s architecture was first reported in 2010, which confirmed the topological predictions in the previous studies, and further demonstrated that approximately nine units of yeast Fld1p assembly a

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stable homooligomer in the form of a toroid [79]. G225P (orthologous pathogenic mutation of A212P in human SEIPIN) is unstable and can only form a smaller homooligomer. To better understand the structure of human SEIPIN, a variety of naturally occurring mutants, including four premature stop mutations (E113X, R138X, R275X, Q391X) and three single amino acid substitution mutations (T78A, L91P, A212P), have been comprehensively investigated. All these mutants can be expressed in both HEK293 cells and C3H10T1/2 preadipocytes, except R275X that is not expressed in C3H10T1/2 [80]. In term of subcellular localization, wild-type (WT) and most mutants localize to the ER, whereas L91P and A212P are partially mislocalized to the nuclear envelope. In addition, L91P and A212P form smaller and less homogenous oligomers, as compared with WT and T78A. In 13

ACCEPTED MANUSCRIPT contrast to yeast Fld1p that forms a nonamer, WT and T78A human SEIPIN assemble into a dodecamer as observed by atomic force microscopy [80]. However, the complete atomic model of

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SEIPIN architecture, the organization and orientation of SEIPIN upon homooligomer assembly or

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physical interaction with other proteins, as well as the structure-function relationship remain to be elucidated. It should also be noted that A212P is extremely unstable when expressed in certain cell

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lines [21].

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SEIPIN in LD growth

LDs are highly dynamic organelles, whose number and size dramatically vary between cell types and

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undergo constant change under different physiological states [10, 81]. Yeast cells usually form LDs

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of less than 250 nm in diameter; while white adipocytes contain giant unilocular LDs up to 100 μm

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[12]. Hepatocytes normally contain a few small LDs; however, the size of hepatocellular LDs can increase tremendously in hepatic steatosis [82]. Even within a single cell, the sizes of LDs can vary

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greatly. For example, in Drosophila S2 cells, TAG synthesis enzymes DGAT2 and GPAT4 are found to associate with one subset of LDs, which keep expanding; whereas another subset of LDs devoid of these enzymes remain small [15].

Two independent genome wide screens both identified Fld1p/Sei1p as a critical regulator of LD morphology in Saccharomyces cerevisiae [20, 22]. In the absence of Fld1p, cells contain either many small, aggregated LDs or a few supersized LDs, depending on growth conditions. In fld1Δ yeast cells, overexpression of either long (462aa) or short (398aa) form of human SEIPIN or the 443aa mouse SEIPIN reverses supersized LD phenotype. In addition, N88S and S90L, but not A212P, expression 14

ACCEPTED MANUSCRIPT rescue the defects in yeast [20]. In a genome wide analyses of sterol-lipid storage and trafficking, ldb16Δ strain has been found to contain reduced number of LDs [83]. Although there is little

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similarity in primary structure or domain organization between Fld1p and Ldb16p, ldb16Δ was

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shown to phenocopy fld1Δ in regard to the accumulation of two types of LDs: small aggregated or supersized, though ldb16Δ more frequently develop small LDs in cluster [24]. Similar to that in fld1Δ, supersized LD phenotype in ldb16Δ can be reversed by addition of inositol, but not choline

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nor ethanolamine. Fld1p and Ldb16p physically interact and function together at ER-LD contact sites

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[21, 24, 25]. Interestingly, human SEIPIN can restore LD defects in both ldb16Δ and fld1Δ ldb16Δ strains, whereas Fld1p fails to complement Ldb16p deficiency [24]. There is no recognizable Ldb16p

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homologue in higher eukaryotes; it was speculated that Fld1p and Ldb16p converge on SEIPIN

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during evolution, whereby Fld1p is transformed into two transmembrane helices and ER luminal

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loop, while Ldb16p constitutes part of ER luminal region and the C-terminal tail. The CGL2 causative mutants, A212P, Y187C, L91P and R275X fail to rescue LD defects in either fld1Δ or

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ldb16Δ, suggesting some potential links between LD abnormality and defective adipogenesis [24]. Of note, the first 280a.a. conserved region of human SEIPIN is able to restore LD defects in both fld1Δ and ldb16Δ strains, indicating the complementation does not require the C-terminal region of mammalian SEIPIN[20, 24].

Aberrant LD morphology has also been observed in other SEIPIN deficient models. SEIPIN-/lymphoblastoid cells and fibroblasts derived from CGL2 patients exhibit increased number of smaller and sometimes aggregated LDs [22, 84]. Global knockout (KO) of SEIPIN in mice results in a CGL phenotype and small unilocular LDs in white adipocytes [85-87]; whereas adipose-specific 15

ACCEPTED MANUSCRIPT KO (ASKO) of SEIPIN causes progressive loss of WAT, accompanied by enlarged unilocular LDs in white adipocytes [88]. The smaller LD size in the remnant WAT of global KO mice is attributed to an

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insufficient differentiation of white adipocytes, while increased LD size in ASKO cells recapitulates

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LD function of SEIPIN in yeast and other cells. Gigantic LDs are also present in SEIPIN deficient testes, where SEIPIN is normally highly expressed [89], as well as adipocytes differentiated from Seipin-/- MEFs [86, 90]. In response to prolonged oleate exposure, SEIPIN deficient cells tend to

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develop supersized LDs, as seen in A431 cells and CGL2-derived fibroblasts [23], Drosophila S2

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cells [91] and 3T3-L1 preadipocytes [21]. How does SEIPIN deficiency cause such drastic changes in LD size? To date, three possible mechanisms have been proposed: 1) by regulating PL

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composition on LD surface, 2) by regulating ER homeostasis and 3) by regulating LD biogenesis at

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ER-LD contact sites.

SEIPIN and phospholipids

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Genome wide RNA interference screen in Drosophila S2 cells identified that PC content is correlated with the size and abundance of LDs [41]. Treatment with oleate increases the need of PLs, in particular PC, to coat the neutral lipids in growing LDs. Accordingly, CTP:phosphocholine cytidylyltransferase 1 (CCT1), the rate-limiting enzyme in PC synthesis, translocate to LD surface at times of PC shortage to help generate more PC. PE is the precursor of PC synthesis via PEMT pathway. Increased PE content enhances membrane curvature, and thus may drive droplet fusion into supersized LDs. In addition, suppressing PC synthesis diverts more DAG to TAG synthesis, leading to a decreased surface-to-volume ratio, which is another reason for the formation of supersized LDs under PC deficiency [41]. In support of this, a genome wide screen in yeast also identified key 16

ACCEPTED MANUSCRIPT enzymes (Cho2p and Opi3p) and transcription factors (Ino2p and Ino4p) in PC synthesis as critical regulators of LD growth [43]. Deletion of either resulted in the formation of supersized LDs, which

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can be reversed by addition of choline in growth medium [43]. Indeed, fld1Δ and ldb16Δ strains tend

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to accumulate small LDs during logarithmic growth, a condition when PL synthesis is active, whereas to form supersized LDs as cells approach stationary phase, when PL synthesis is reduced [24]. Nevertheless, some observations in the fld1Δ strain don’t necessarily agree with the PC content

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and surface-to-volume ratio theories [43]: addition of choline to fld1Δ cells has little impact on

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supersized LD phenotype; when grown in YPD medium, supersized LDs no longer appear in cho2Δ, ino2Δ and ino4Δ cells, even though decreased PL-to-TAG ratio persists. Instead, it was demonstrated

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that the increased PA content is a common feature of cho2Δ, opi3Δ, ino2Δ and ino4Δ strains [43]. PA

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is a cone-shaped lipid that alters membrane curvature, and has been shown to promote both

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SNARE-dependent and -independent membrane fusion events [92, 93]. Although total cellular PA content was unaltered, fld1Δ cells exhibit an increased PA in the microsomal fraction [21, 43]. PA

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accumulation in the ER adjacent to growing LDs has also been visualized by fluorescent probes in both fld1Δ and ldb16Δ strains, which can be reversed by expression of human SEIPIN [94]. Consistent with the notion that localized PA accumulation drives LD fusion and growth, deletion of fld1 results in enrichment of PA probes in the proximity of nuclear envelope and aberrant expansion of nuclear ER [95]. The clustered LDs are transformed to supersized LDs when PC synthesis is compromised, indicating that Fld1p affects PA homeostasis at LD-forming subdomain of the nuclear envelope.

In Drosophila, dSEIPIN (CG9904) null mutants are viable and fertile with no noticeable behavioral 17

ACCEPTED MANUSCRIPT defects [96], but they exhibit larger LDs in larval salivary gland [21]. Notably, the content of total PA and a collection of PA species are increased in dSEIPIN null mutant larvae as compared to WT

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counterparts [96]. In 3T3-L1 preadipocytes, knocking down SEIPIN dramatically increases total

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cellular PA content in the presence of AGPAT2 overexpression that favors PA production [97]. Although overexpression of SEIPIN has little impact on microsomal PA in 3T3-L1 preadipocytes prior to adipogenic induction [21], it profoundly decreases total cellular PA on the day 2 of adipocyte

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differentiation [97]. The testes in both global SEIPIN KO and germ cell specific KO mice contain a

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higher total PA content and higher ratios of PA to PLs, as compared to WT littermates [89]. Together, data from various organisms suggest that localized increase in PA is the underlying reason for the

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development of abnormal LDs in SEIPIN deficient cells.

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The mechanism whereby SEIPIN/Fld1p regulates PA metabolism remains obscure. Lipin-1, the enzyme that diverts PA into DAG, was reported to interact with SEIPIN. Knocking down lipin-1

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leads to formation of supersized LDs, even though TAG storage is reduced [38, 43]. On the day 2 of 3T3-L1 adipocyte differentiation, SEIPIN knockdown reduces the quantity of lipin-1 bound to the ER, and increases local PA accumulation. The interacting domain resides in the C- and/or N- termini, but not ER luminal loop, of SEIPIN. SEIPIN mutants lacking C-termini failed to repress PA production as compared to the WT [97]. In yeast cells, fld1Δ results in aggregation of Pah1p, yeast ortholog of lipin-1, along with increased PA puncta at the ER adjacent to growing LDs [94]. However, the hypothesis that SEIPIN anchors and facilitates lipin-1 function at ER-LD junctions might overlook existing physiological evidence. For example, depleting lipin-1 in mouse adipocytes reduces LD size [98], whereas depleting SEIPIN in mouse adipocytes increases LD size [88]. Also, 18

ACCEPTED MANUSCRIPT loss of SEIPIN function cause much less severe changes in yeast and mammals as compared to a loss of PAP activity [99]. In addition, a recent study failed to detect a SEIPIN-lipin-1 interaction [21].

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Finally, a loss of Pah1 dramatically reduced the amount of cellular TAG whereas a loss of Fld1/Sei1

might interact with lipin-1 requires further studies.

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had a rather limited impact on TAG synthesis in yeast [20, 100]. Therefore, whether and how SEIPIN

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AGPAT2 is the disease locus of type 1 CGL [67, 101]. AGPAT2 is a member of enzymes possessing

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lysophosphatidic acid acyltransferase (LPAAT) activity, which includes AGPAT family (AGPAT1-11), and others such as CGI-58 and endophilin [102]. In comparison with other AGPATs, AGPAT2 is the

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most abundant isoform expressed in adipose tissue [103]. AGPAT2 converts LPA to PA, sitting

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upstream of lipins in the biosynthetic pathway of PLs and TAGs (Figure 1)[10, 31, 38]. Both long

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and short forms of human SEIPIN were shown to interact with human AGPAT2, which, in contrast to Lipin-1-SEIPIN interaction, relies on the luminal loop and the first transmembrane domain (adjacent

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to N-terminus) [104]. Atomic force microscopy reveals that both AGPAT2 and Lipin-1 can directly and simultaneously interact with SEIPIN dodecamer, but there is no report on any functional consequence of this scaffold, including the impact on localized PA accumulation. Also, the functional interactions between SEIPIN and AGPAT2 have yet to be fully investigated.

In a more recent study, Fld1p/Sei1p interacting partners have been screened in the yeast Saccharomyces cerevisiae, using affinity isolation and tandem mass spectrometry analyses. The most prominent Fld1p partner is the Gat1p, a yeast GPAT [21]. As rate-limiting step in PA biosynthesis, GPATs convert glycerol-3-phosphate to LPA. Mammals express four GPAT isoforms; GPAT1 and 19

ACCEPTED MANUSCRIPT GPAT2 are present on the outer mitochondrial membrane, while GPAT3 and GPAT4 are present on the ER membrane [2]. SEIPIN is shown to interact with both ER GPATs in 3T3-L1 preadipocytes,

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which can be weakened by T78A missense mutation and is independent of its C-terminal extension.

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Importantly, in yeast cells, mouse embryonic fibroblasts (MEFs), 3T3-L1 preadipocytes and mouse testes, GPAT activity was significantly increased when SEIPIN/Fld1p was deleted. As a consequence, microsomal PA level is elevated in SEIPIN deficient cells. Conversely, enforced expression of

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ER-located GPAT isoforms in either 3T3-L1 preadipocytes or yeast increases microsomal PA level,

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which is compromised by co-expressing SEIPIN/Fld1p [21]. Accordingly, LD size is positively correlated to microsomal PA level under these genetic manipulations. In contrast to the interactions

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with Lipin-1 and AGPAT2 whereby SEIPIN was proposed to facilitate their functions, the interaction

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between SEIPIN and ER-located GPATs leads to inhibition of GPAT function (Figure 1) [21]. These

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findings together favor the concept that aberrant LDs in SEIPIN deficient cells arise from increased

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activities of ER GPATs and local PA abundance.

SEIPIN and ER homeostasis High resolution fluorescent microscopy and electron microscopy revealed that the mobility of LDs within fld1Δ cell and their transfer into daughter cells are both greatly impaired, possibly due to the aberrant ER structure [105]. Defective LD mobility and inheritance cause TAG accumulation, followed by abnormal LD growth. The ER is also the major intracellular calcium store, and impairments in calcium homeostasis lead to ER stress, cellular dysfunction or even death [106-108]. The sarco/endoplasmic reticulum Ca2+-ATPase (SERCA) pumps cytosolic calcium into the ER lumen and maintains a high calcium concentration gradient between the ER lumen and the cytosol at 20

ACCEPTED MANUSCRIPT rest. SERCA is found to interact with SEIPIN in both Drosophila larvae and cultured human cell lines [109]. The absence of dSEIPIN causes a reduced calcium pump activity and a loss of dSERCA

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function. The loop region is required for the dSEIPIN oligomerization and interaction with dSERCA.

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Knocking down dSERCA reduces LD size in fat cells, but it can’t be rescued by overexpression of dSEIPIN, which places dSERCA downstream of dSEIPIN [109]. A recent study also reported a widening of ER lumen in SEIPIN knockout cells [23], however, exactly how SEIPIN may regulate

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ER homeostasis is not clear. It is conceivable that changes in phospholipids can alter general ER

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function and calcium homeostasis [110].

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SEIPIN and LD maturation at ER-LD contact sites

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SEIPIN and its orthologs may function at a discrete step in LD biogenesis. Fld1p is found to localize

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to ER-LD junction nearly a decade ago [22], where it forms a complex with another ER membrane protein Ldb16p [21, 24, 25]. Both overexpressed and endogenous human SEIPIN are also shown to

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localize to ER-LD contact sites [23, 26]. LD surface monolayer is in continuous with the cytoplasmic leaflet of the ER bilayer, but it has distinct biophysical properties and composition of proteins and lipids. The complex of Fld1p and Ldb16p is postulated to establish a diffusion barrier between two compartments, and thus regulate LD dynamics [25]. In early stage of LD biogenesis, fld1Δ drastically impedes de novo LD formation, accompanied by ER membrane accumulation of unpacked TAGs [111]. Deletion of 14aa in N-terminus of Fld1p causes both supersized LD phenotype and blockade in initiating LD biogenesis, suggesting these two processes may be dissectible [111]. In Drosophila S2 cells, oligomers of dSEIPIN form highly mobile foci at contact sites of the ER and small, nascent LDs, where they might facilitate lipid transfer from ER to nascent 21

ACCEPTED MANUSCRIPT LDs [26]. Without dSEIPIN, these nascent LDs fail to grow, resulting in massive accumulation of premature LDs. dSEIPIN regulates a more distal step in the maturation of nascent LDs and facilitates

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their expansion, rather than initial lens formation and TAG droplets budding in the initiation of LD

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biogenesis. These data explain why clustering of small LDs appears in SEIPIN deficient cells. After prolonged exposure to oleate, dSEIPIN null mutants elicit aberrant dGPAT4 targeting to ~72% LDs, in contrast to dGPAT4 coating only ~8% LDs in WT cells. Concurrently, dSEIPIN null LDs are

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deficient in PLs, along with an increased CCT1 association. These data explain why a subset of LDs,

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characterized as eLDs, can be fast transformed into supersized LDs in dSEIPIN null S2 cells [26]. Similar findings were also shown in SEIPIN KO A431 cells using CRISPR/Cas9 [23]. After 3 days

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of delipidation, 1h oleate treatment induces formation of numerous small LDs in varying sizes in

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SEIPIN KO cells, indicative of delayed LD maturation. Normal LD growth can be restored by

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re-introducing WT SEIPIN, but not A212P. In contrast to dSEIPIN null S2 cells where nascent LDs unable to grow accumulate in contact sites with the ER, SEIPIN KO A431 cells exhibit highly

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mobile small LDs with almost no contact with the ER. SEIPIN KO impairs lipid cargo transport and protein trafficking from the ER to LDs. Even with prolonged exposure to oleate, a subset of small LDs behaves in the same manner. In contrast, all the large LDs keep connectivity with the ER, with less mobility, which explains the heterogeneity of LD morphology in SEIPIN deficient cells [23]. However, the exact role of SEIPIN at ER-LD contacts remains to be established, as SEIPIN is not required for establishing such contacts. It is also conceivable that disrupted ER phospholipid homeostasis due to SEIPIN deficiency may affect the integrity of ER-LD contacts, as well as the lipid transfer from ER to LDs.

22

ACCEPTED MANUSCRIPT SEIPIN in adipogenesis SEIPIN has distinct tissue distribution pattern in different organisms. In human, SEIPIN is highly

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expressed in brain and testes, followed by pancreas and kidney. Although SEIPIN is not abundant in

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adipose tissue, null or missense mutations in SEIPIN lead to the most severe form of lipodystrophy, CGL2 in humans [72]. The distribution of SEIPIN in mouse displays another pattern, with highest expression in testes, WAT and BAT, moderate in skeletal muscle and adrenal gland, and low in brain

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[112]. In response to adipogenic stimulation at terminal phase, SEIPIN mRNA level is increased by

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several folds in both mouse and human primary adipocytes as well as cultured cell lines, such as C3H10T1/2, 3T3-L1 and MEFs; whereas SEIPIN level is unchanged in the earlier determination

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phase [112, 113]. Indeed, knocking down SEIPIN drastically impairs the induction of “master”

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transcription factors, such as PPARγ and C/EBPα, and their target genes; whereas it does not alter

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determination phase markers, such as C/EBPβ, C/EBPδ and BMP4 [112, 113]. This finding has also

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been seen in Seipin-/- MEFs and stromal vascular cells [86, 90].

Seipin-/- mice recapitulate most features of CGL2 patients, exhibiting severe lipodystrophy, with at least 90% loss of WAT. The remnant fat can be detected in some visceral and subcutaneous locations, such as inguinal, epididymal or mesenteric regions [85, 86, 90, 114]. Due largely to the loss of WAT, adipokines such as adiponectin and leptin are reduced dramatically, while ectopic fat extensively accumulates in non-adipose tissues, leading to severe insulin resistance and hepatic steatosis [85, 86, 90]. In Seipin-/- mice, the remaining epididymal and subcutaneous WATs consist almost entirely of small immature adipocytes, most of which contain brightly eosinophilic cytoplasm, loaded with smaller unilocular LDs [85, 86]. The BAT mass is also greatly reduced in Seipin-/- mice. The 23

ACCEPTED MANUSCRIPT remaining BAT exhibit few small LDs interspersed with occasional giant LD, instead of multilocular LD phenotype [86]. An essential role for SEIPIN in human adipogenesis was recently demonstrated

However, two recent studies reported that SEIPIN is not required for brown

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mutations[115].

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by using induced pluripotent stem (iPS) cells from patients with CGL harboring BSCL2/SEIPIN

adipogenesis[116, 117]. It is quite surprising that SEIPIN deficiency exerts different effects on white and brown adipogenesis. Additional studies in vitro and in vivo are required to unveil the molecular

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basis for such differential effects.

To assess the role of SEIPIN in mature adipocytes, adipose-specific seipin knockout mice (ASKO

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mice) were generated by applying the aP2 promotor to drive Cre recombinase expression, and thus

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SEIPIN is deleted after formation of fat depots because aP2 is expressed at late stages of adipocyte

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differentiation [88]. ASKO mice display progressive loss of WAT by ~25% at 3 months, ~50% at 6 months and ~75% at 10 months. The adipocytes in ASKO epididymal and subcutaneous WAT

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become more hypertrophic over the time, while the LD size in these cells turns to be vastly variable. ASKO BAT also undergoes a progressive loss, but less severe than WAT. Strikingly, the remnant brown adipocytes contain gigantic, white adipocyte–like LDs [88]. In a novel mouse model, where SEIPIN is conditionally deleted by tamoxifen treatment in adult mice at 8-10 weeks old, SEIPIN deletion does not cause acute loss of WAT immediately after tamoxifen administration, but it induces dramatic and progressive fat loss in the following 12 weeks, akin to the ASKO mice [118]. These acquired KO mice are resistant to diet-induced obesity and dyslipidemia, but tend to develop insulin resistance and hepatic steatosis. In agreement with the general conclusion from the two studies above that SEIPIN is required for the maintenance of mature adipocytes, a recent cellular model also 24

ACCEPTED MANUSCRIPT showed progressive loss of mature adipocytes under inducible SEIPIN deficiency [119].

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Generation of Seipin-/- mouse models provides further insights into exactly how SEIPIN deficiency

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impairs adipogenesis. To date, two mechanisms have been proposed: 1) by regulating adipocyte lipolysis or 2) by regulating PPARγ activation. SEIPIN and adipocyte lipolysis

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Chen et al attribute the impaired adipogenesis to unbridled cyclic AMP (cAMP)-dependent

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PKA-activated lipolysis during the adipocyte differentiation [86]. The enhanced lipolysis underlies loss of LDs and silencing of adipogenic transcription factors in differentiating Seipin-/- MEFs, as

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adipogenic defects can be rescued by treatment with TAG lipase inhibitor, E600, but not by PPARγ

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agonist, pioglitazone. Notably, basal, but not stimulated, lipolysis increased in Seipin-/- cells [86]. In

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partial agreement with this, PPARγ agonist, rosiglitazone fails to correct the increased basal lipolysis in Seipin-/- MEFs. However, contrary to Chen et al.’s work, rosiglitazone was shown to be able to

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restore multiple/well-defined pattern of LDs, mRNA expression of PPARγ and Adipoq, and adipocyte differentiation in mature Seipin-/- MEF adipocytes [90]. In addition, the effect of SEIPIN in adipocyte lipolysis is still under debate. In ASKO mice, basal lipolytic rate is comparable to that of WT counterparts; whereas β3-adrenergic stimulated lipolysis is decreased in vivo, which contributes to the enlargement of LDs as well as white adipocyte hypertrophy [88]. Stimulated lipolysis is also impaired in epididymal WAT explant ex vivo, as reflected by a decreased release of FFA and glycerol, and defective HSL phosphorylation [88]. In support of this, adipose-specific SEIPIN (short form of human SEIPIN) transgenic mice exhibit increased lipolysis at both basal and stimulated states, which is accompanied with reduced adipocyte volume and size [120]. In the 25

ACCEPTED MANUSCRIPT tamoxifen-induced SEIPIN-/- adult mice, basal lipolysis is minimally elevated, but responses to lipolytic stimulation are defective, which is attributed to the downregulation of the β3 adrenergic

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receptor [118]. These findings together suggest SEIPIN can regulate adipocyte lipolysis, although its

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role seems changing over the course of adipogenesis. The pathophysiological role of lipolysis in the development and maintenance of adipose tissue also remains obscure. Therefore, exactly how SEIPIN regulates lipolysis, and whether SEIPIN regulates adipogenesis through its role in lipolysis,

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await further investigation.

SEIPIN and PPARγ activation

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It was shown that restoration of adipogenic transcription program by PPARγ can rescue adipocyte

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differentiation and mitigate metabolic disorders in SEIPIN deficient models [90]. TZDs including

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pioglitazone and rosiglitazone are a group of pharmacological PPARγ agonists. Pioglitazone induces the expression of adipogenic marker aP2 in MEFs from CGL2 patients [121], and it also improves

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insulin resistance and hepatic steatosis in global SEIPIN-/- mice [90]. We have also observed that rosiglitazone can partially restore defective adipocyte differentiation in SEIPIN deficient 3T3-L1 cells and MEFs (unpublished). In ASKO mice, although rosiglitazone treatment failed to normalize body weight or total WAT mass, it significantly increased WAT mass in subcutaneous and inguinal areas and drove the formation of newly differentiated adipocytes in subcutaneous fat [88]. Clinically, TZDs have been demonstrated to promote fat recovery in HIV-induced acquired lipodystrophy [122, 123]. Pagac et al further elucidated how SEIPIN deficiency impairs adipogenesis via PPARγ [21]. In 3T3-L1 cells, SEIPIN interacts with ER GPATs, GPAT3 and GPAT4, which are rate-limiting enzyme in PA biosynthesis [21]. Upon adipocyte differentiation, GPAT3 mRNA is elevated by ~60 fold, 26

ACCEPTED MANUSCRIPT whereas GPAT4 induction is fairly modest. Knocking down of GPAT3, but not GPAT4, profoundly decreases total GPAT activity, followed by inhibition of lipid accumulation and decreased expression

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of adipogenic markers in 3T3-L1 adipocytes [124]. Paradoxically, low GPAT activity during the first

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few hours of adipocyte differentiation allows PPARγ to be fully active. Overexpression of GPAT3 elevates microsomal PA level and blocks adipogenesis, which can be restored by co-overexpression of SEIPIN; while knocking down GPAT3 rescues microsomal PA accumulation and adipogenic

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defects in SEIPIN knockdown 3T3-L1 cells. GPAT inhibitor (formula: C21H26CINO4S) has also

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been demonstrated to partially rescue adipogenesis in Seipin-/- MEF adipocytes [21]. In addition to leptin and TZDs, GPAT inhibitor might become a promising therapeutic option for the treatment of

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CGL2. These data suggest that PA may serve as a strong antagonist of PPARγ. In support of this, PA

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was shown to be a competitive inhibitor of PPARγ, and a PA derivative, cyclic PA is an endogenous

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antagonist of PPARγ, which blocks adipogenic differentiation of 3T3-L1 cells [62]. Moreover, many conditions that increase PA by disrupting the glycerol-3-phosphate pathway, including depleting

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AGPAT2, Lipin-1 or CDS1/2, or overexpressing GPAT, are all associated with defective adipogenesis and severe lipodystrophy [21, 38, 44, 71]. Therefore, SEIPIN may regulate adipogenesis through its role in regulating PA distribution/metabolism.

Hypothesis and future directions The development of adipocytes/adipogenesis and the growth of LDs are often coupled, however, they are mechanistically distinct processes: adipogenesis entails the formation of adipocytes from stem cells in mammals, whereas LD growth is a cellular process that is conserved from bacteria to man. Results from recent studies strongly suggest that adipogenesis and LD biogenesis/growth may 27

ACCEPTED MANUSCRIPT share some common factors. SEIPIN, lipin-1, CDS1 and AGPAT2, conserved from yeast to man, can all regulate the maturation of both LDs and adipocytes: loss of SEIPIN, lipin-1, CDS1 or AGPAT2

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can completely block adipogenesis; aberrant LD morphology has also been observed in cells lacking

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SEIPIN, lipin-1 CDS1, and AGPAT2 ([38, 43], and our unpublished data). Lipin/PAP, CDS1, and AGPAT2 are well known enzymes that mediate the synthesis of phospholipids and/or TAGs (Figure 1). The molecular function of SEIPIN is not clear, although there is ample genetic and biochemical

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evidence implicating SEIPIN in the metabolism of phospholipids, especially that of phosphatidic

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acid (PA). We therefore propose the following scheme/mechanism linking LD growth and adipogenesis (Figure 3): a loss of SEIPIN, lipin-1, CDS1 or AGPAT2 function leads to general and/or

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localized changes in the level of lipid intermediates, particularly an increase of phosphatidic acid

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(PA). In preadipocytes, PA may serve as a strong antagonist of PPARγ, thereby blocking

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adipogenesis. The fact that aberrant LDs were observed in the nuclei of SEIPIN-deficient yeast cells suggests that SEIPIN can impact nuclear functions [111]. In all cell types including yeast, PA may

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affect LD budding and/or growth, leading to the formation of abnormal LDs. Under such a scheme, LD growth and adipogenesis are connected through phospholipids.

To fully understand the role of phospholipids in LD biogenesis/growth and adipogenesis, the following challenges require immediate attention in the near future: 1. Reliable methods need to be developed to accurately detect and quantitate localized changes of membrane lipids, such as phospholipids, DAG, cholesterol and sphingolipids. 2. The atomic structures of the enzymes and proteins regulating lipid synthesis and storage need to be resolved. Recent development in cryo-electron microscopy (Cryo-EM) technology offers endless opportunities to lipid researchers. In 28

ACCEPTED MANUSCRIPT the case of phosphatidic acid (PA) and PPARγ, it would be important to crystalize PPARγ together with PA to determine whether and exactly how PA may modulate PPARγ function. 3. It is also

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important to examine changes in the nuclear envelope and the nuclei when the genes mentioned

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above are disrupted. Lipin-1 has been shown to play an important role in gene regulation, which requires its enzymatic activity [99]. Likewise, SEIPIN, GPATs, AGPAT2 or CDS1/2 may also impact the transcriptional regulation of genes through lipids. It is hoped that these and other studies will

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enable a better mechanistic appreciation of how our cells and body store fat, which may lead to

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effective therapeutic strategies against obesity and its related disorders.

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Acknowledgements

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We sincerely apologize to those whose work could not be cited or discussed here due to limitations in

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space and scope. This work was supported by grants 1057852 and 1078117 from the National

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Health and Medical Research Council (NHMRC), Australia, and by grant DP130100457 from the Australian Research Council. H. Yang is a Senior Research Fellow of the NHMRC (1058237). L.

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Sun is supported by Singapore NRF fellowship: NRF-2011NRF-NRFF001-025 and by Singapore

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National Research Foundation CBRG grant NMRC/CBRG/0101/2016.

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ACCEPTED MANUSCRIPT References [1] A.L. Robbins, D.B. Savage, The genetics of lipid storage and human lipodystrophies, Trends in molecular medicine, 21 (2015) 433-438.

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[2] R.A. Coleman, D.G. Mashek, Mammalian triacylglycerol metabolism: synthesis, lipolysis, and signaling, Chemical reviews, 111 (2011) 6359-6386.

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[3] M.A. Welte, Expanding roles for lipid droplets, Current biology : CB, 25 (2015) R470-481. cell biology, 14 (2013) 775-786.

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[4] A.R. Thiam, R.V. Farese, Jr., T.C. Walther, The biophysics and cell biology of lipid droplets, Nature reviews. Molecular [5] H.F. Hashemi, J.M. Goodman, The life cycle of lipid droplets, Current opinion in cell biology, 33 (2015) 119-124. [6] S.D. Kohlwein, Obese and anorexic yeasts: experimental models to understand the metabolic syndrome and lipotoxicity, Biochimica et biophysica acta, 1801 (2010) 222-229.

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[7] B. Gustafson, S. Hedjazifar, S. Gogg, A. Hammarstedt, U. Smith, Insulin resistance and impaired adipogenesis, Trends in endocrinology and metabolism: TEM, 26 (2015) 193-200.

[8] J.S. Tan, C.J. Seow, V.J. Goh, D.L. Silver, Recent advances in understanding proteins involved in lipid droplet formation,

MA

growth and fusion, Journal of genetics and genomics = Yi chuan xue bao, 41 (2014) 251-259. [9] R.V. Farese, Jr., T.C. Walther, Lipid droplets finally get a little R-E-S-P-E-C-T, Cell, 139 (2009) 855-860. [10] W. Fei, X. Du, H. Yang, Seipin, adipogenesis and lipid droplets, Trends in endocrinology and metabolism: TEM, 22 (2011) 204-210.

D

[11] K.K. Buhman, H.C. Chen, R.V. Farese, Jr., The enzymes of neutral lipid synthesis, The Journal of biological chemistry,

TE

276 (2001) 40369-40372.

[12] C.W. Wang, Lipid droplet dynamics in budding yeast, Cellular and molecular life sciences : CMLS, 72 (2015) 2677-2695.

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[13] A. Pol, S.P. Gross, R.G. Parton, Review: biogenesis of the multifunctional lipid droplet: lipids, proteins, and sites, The Journal of cell biology, 204 (2014) 635-646. [14] Y. Ohsaki, M. Suzuki, T. Fujimoto, Open questions in lipid droplet biology, Chemistry & biology, 21 (2014) 86-96. [15] F. Wilfling, H. Wang, J.T. Haas, N. Krahmer, T.J. Gould, A. Uchida, J.X. Cheng, M. Graham, R. Christiano, F. Frohlich, X.

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Liu, K.K. Buhman, R.A. Coleman, J. Bewersdorf, R.V. Farese, Jr., T.C. Walther, Triacylglycerol synthesis enzymes mediate lipid droplet growth by relocalizing from the ER to lipid droplets, Developmental cell, 24 (2013) 384-399. [16] D.A. Gross, C. Zhan, D.L. Silver, Direct binding of triglyceride to fat storage-inducing transmembrane proteins 1 and 2 is important for lipid droplet formation, Proc Natl Acad Sci U S A, 108 (2011) 19581-19586. [17] V. Choudhary, N. Ojha, A. Golden, W.A. Prinz, A conserved family of proteins facilitates nascent lipid droplet budding from the ER, The Journal of cell biology, 211 (2015) 261-271. [18] H. Yang, A. Galea, V. Sytnyk, M. Crossley, Controlling the size of lipid droplets: lipid and protein factors, Current opinion in cell biology, 24 (2012) 509-516. [19] N. Jacquier, V. Choudhary, M. Mari, A. Toulmay, F. Reggiori, R. Schneiter, Lipid droplets are functionally connected to the endoplasmic reticulum in Saccharomyces cerevisiae, Journal of cell science, 124 (2011) 2424-2437. [20] W. Fei, G. Shui, B. Gaeta, X. Du, L. Kuerschner, P. Li, A.J. Brown, M.R. Wenk, R.G. Parton, H. Yang, Fld1p, a functional homologue of human seipin, regulates the size of lipid droplets in yeast, The Journal of cell biology, 180 (2008) 473-482. [21] M. Pagac, D.E. Cooper, Y. Qi, I.E. Lukmantara, H.Y. Mak, Z. Wu, Y. Tian, Z. Liu, M. Lei, X. Du, C. Ferguson, D. Kotevski, P. Sadowski, W. Chen, S. Boroda, T.E. Harris, G. Liu, R.G. Parton, X. Huang, R.A. Coleman, H. Yang, SEIPIN Regulates Lipid Droplet Expansion and Adipocyte Development by Modulating the Activity of Glycerol-3-phosphate Acyltransferase, Cell reports, 17 (2016) 1546-1559. [22] K.M. Szymanski, D. Binns, R. Bartz, N.V. Grishin, W.P. Li, A.K. Agarwal, A. Garg, R.G. Anderson, J.M. Goodman, The

31

ACCEPTED MANUSCRIPT lipodystrophy protein seipin is found at endoplasmic reticulum lipid droplet junctions and is important for droplet morphology, Proc Natl Acad Sci U S A, 104 (2007) 20890-20895. [23] V.T. Salo, I. Belevich, S. Li, L. Karhinen, H. Vihinen, C. Vigouroux, J. Magre, C. Thiele, M. Holtta-Vuori, E. Jokitalo, E. Ikonen, Seipin regulates ER-lipid droplet contacts and cargo delivery, The EMBO journal, 35 (2016) 2699-2716.

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[24] C.W. Wang, Y.H. Miao, Y.S. Chang, Control of lipid droplet size in budding yeast requires the collaboration between Fld1 and Ldb16, Journal of cell science, 127 (2014) 1214-1228.

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[25] A. Grippa, L. Buxo, G. Mora, C. Funaya, F.Z. Idrissi, F. Mancuso, R. Gomez, J. Muntanya, E. Sabido, P. Carvalho, The seipin complex Fld1/Ldb16 stabilizes ER-lipid droplet contact sites, The Journal of cell biology, 211 (2015) 829-844.

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[26] H. Wang, M. Becuwe, B.E. Housden, C. Chitraju, A.J. Porras, M.M. Graham, X.N. Liu, A.R. Thiam, D.B. Savage, A.K. Agarwal, A. Garg, M.J. Olarte, Q. Lin, F. Frohlich, H.K. Hannibal-Bach, S. Upadhyayula, N. Perrimon, T. Kirchhausen, C.S. Ejsing, T.C. Walther, R.V. Farese, Seipin is required for converting nascent to mature lipid droplets, eLife, 5 (2016). [27] J. Gong, Z. Sun, L. Wu, W. Xu, N. Schieber, D. Xu, G. Shui, H. Yang, R.G. Parton, P. Li, Fsp27 promotes lipid droplet

NU

growth by lipid exchange and transfer at lipid droplet contact sites, The Journal of cell biology, 195 (2011) 953-963. [28] L. Zhou, S.Y. Park, L. Xu, X. Xia, J. Ye, L. Su, K.H. Jeong, J.H. Hur, H. Oh, Y. Tamori, C.M. Zingaretti, S. Cinti, J. Argente, M. Yu, L. Wu, S. Ju, F. Guan, H. Yang, C.S. Choi, D.B. Savage, P. Li, Insulin resistance and white adipose tissue

MA

inflammation are uncoupled in energetically challenged Fsp27-deficient mice, Nature communications, 6 (2015) 5949. [29] L. Xu, L. Zhou, P. Li, CIDE Proteins and Lipid Metabolism, Arteriosclerosis, thrombosis, and vascular biology, 32 (2012) 1094-1098. dietary fatty acids, Hepatology, (2012).

D

[30] L. Zhou, L. Xu, J. Ye, D. Li, W. Wang, X. Li, L. Wu, H. Wang, F. Guan, P. Li, Cidea promotes hepatic steatosis by sensing

TE

[31] J.P. Fernandez-Murray, C.R. McMaster, Lipid synthesis and membrane contact sites: a crossroads for cellular physiology, Journal of lipid research, 57 (2016) 1789-1805. [32] N.E. Wolins, B.K. Quaynor, J.R. Skinner, M.J. Schoenfish, A. Tzekov, P.E. Bickel, S3-12, Adipophilin, and TIP47 package

CE P

lipid in adipocytes, The Journal of biological chemistry, 280 (2005) 19146-19155. [33] N. Jacquier, S. Mishra, V. Choudhary, R. Schneiter, Expression of oleosin and perilipins in yeast promotes formation of lipid droplets from the endoplasmic reticulum, Journal of cell science, 126 (2013) 5198-5209. [34] Z. Sun, J. Gong, H. Wu, W. Xu, L. Wu, D. Xu, J. Gao, J.W. Wu, H. Yang, M. Yang, P. Li, Perilipin1 promotes unilocular

AC

lipid droplet formation through the activation of Fsp27 in adipocytes, Nature communications, 4 (2013) 1594. [35] M. Schweiger, R. Schreiber, G. Haemmerle, A. Lass, C. Fledelius, P. Jacobsen, H. Tornqvist, R. Zechner, R. Zimmermann, Adipose triglyceride lipase and hormone-sensitive lipase are the major enzymes in adipose tissue triacylglycerol catabolism, The Journal of biological chemistry, 281 (2006) 40236-40241. [36] A. Lass, R. Zimmermann, M. Oberer, R. Zechner, Lipolysis - a highly regulated multi-enzyme complex mediates the catabolism of cellular fat stores, Progress in lipid research, 50 (2011) 14-27. [37] D.A. Gross, D.L. Silver, Cytosolic lipid droplets: from mechanisms of fat storage to disease, Critical reviews in biochemistry and molecular biology, 49 (2014) 304-326. [38] Y. Qi, T.S. Kapterian, X. Du, Q. Ma, W. Fei, Y. Zhang, X. Huang, I.W. Dawes, H. Yang, CDP-diacylglycerol synthases regulate the growth of lipid droplets and adipocyte development, Journal of lipid research, 57 (2016) 767-780. [39] R. Bartz, W.H. Li, B. Venables, J.K. Zehmer, M.R. Roth, R. Welti, R.G. Anderson, P. Liu, K.D. Chapman, Lipidomics reveals that adiposomes store ether lipids and mediate phospholipid traffic, Journal of lipid research, 48 (2007) 837-847. [40] K. Grillitsch, M. Connerth, H. Kofeler, T.N. Arrey, B. Rietschel, B. Wagner, M. Karas, G. Daum, Lipid particles/droplets of the yeast Saccharomyces cerevisiae revisited: lipidome meets proteome, Biochimica et biophysica acta, 1811 (2011) 1165-1176. [41] Y. Guo, T.C. Walther, M. Rao, N. Stuurman, G. Goshima, K. Terayama, J.S. Wong, R.D. Vale, P. Walter, R.V. Farese,

32

ACCEPTED MANUSCRIPT Functional genomic screen reveals genes involved in lipid-droplet formation and utilization, Nature, 453 (2008) 657-661. [42] N. Krahmer, Y. Guo, F. Wilfling, M. Hilger, S. Lingrell, K. Heger, H.W. Newman, M. Schmidt-Supprian, D.E. Vance, M. Mann, R.V. Farese, Jr., T.C. Walther, Phosphatidylcholine synthesis for lipid droplet expansion is mediated by localized activation of CTP:phosphocholine cytidylyltransferase, Cell Metab, 14 (2011) 504-515.

T

[43] W. Fei, G. Shui, Y. Zhang, N. Krahmer, C. Ferguson, T.S. Kapterian, R.C. Lin, I.W. Dawes, A.J. Brown, P. Li, X. Huang, R.G. Parton, M.R. Wenk, T.C. Walther, H. Yang, A role for phosphatidic Acid in the formation of "supersized" lipid droplets,

IP

PLoS genetics, 7 (2011) e1002201.

[44] V.A. Cortes, D.E. Curtis, S. Sukumaran, X. Shao, V. Parameswara, S. Rashid, A.R. Smith, J. Ren, V. Esser, R.E. Hammer,

SC R

A.K. Agarwal, J.D. Horton, A. Garg, Molecular mechanisms of hepatic steatosis and insulin resistance in the AGPAT2-deficient mouse model of congenital generalized lipodystrophy, Cell Metab, 9 (2009) 165-176. [45] S.E. Gale, A. Frolov, X. Han, P.E. Bickel, L. Cao, A. Bowcock, J.E. Schaffer, D.S. Ory, A regulatory role for 1-acylglycerol-3-phosphate-O-acyltransferase 2 in adipocyte differentiation, The Journal of biological chemistry, 281

NU

(2006) 11082-11089.

[46] D. Barneda, J. Planas-Iglesias, M.L. Gaspar, D. Mohammadyani, S. Prasannan, D. Dormann, G.S. Han, S.A. Jesch, G.M. Carman, V. Kagan, M.G. Parker, N.T. Ktistakis, J. Klein-Seetharaman, A.M. Dixon, S.A. Henry, M. Christian, The brown

MA

adipocyte protein CIDEA promotes lipid droplet fusion via a phosphatidic acid-binding amphipathic helix, eLife, 4 (2015). [47] E.D. Rosen, B.M. Spiegelman, What we talk about when we talk about fat, Cell, 156 (2014) 20-44. [48] J.H. Stern, J.M. Rutkowski, P.E. Scherer, Adiponectin, Leptin, and Fatty Acids in the Maintenance of Metabolic Homeostasis through Adipose Tissue Crosstalk, Cell Metab, 23 (2016) 770-784.

D

[49] P. Cohen, B.M. Spiegelman, Brown and Beige Fat: Molecular Parts of a Thermogenic Machine, Diabetes, 64 (2015)

TE

2346-2351.

[50] S. Gesta, Y.H. Tseng, C.R. Kahn, Developmental origin of fat: tracking obesity to its source, Cell, 131 (2007) 242-256. [51] N. Patni, A. Garg, Congenital generalized lipodystrophies--new insights into metabolic dysfunction, Nature reviews.

CE P

Endocrinology, 11 (2015) 522-534.

[52] S.M. Grundy, Adipose tissue and metabolic syndrome: too much, too little or neither, European journal of clinical investigation, 45 (2015) 1209-1217.

[53] J.Y. Kim, E. van de Wall, M. Laplante, A. Azzara, M.E. Trujillo, S.M. Hofmann, T. Schraw, J.L. Durand, H. Li, G. Li, L.A.

AC

Jelicks, M.F. Mehler, D.Y. Hui, Y. Deshaies, G.I. Shulman, G.J. Schwartz, P.E. Scherer, Obesity-associated improvements in metabolic profile through expansion of adipose tissue, The Journal of clinical investigation, 117 (2007) 2621-2637. [54] V. Peirce, S. Carobbio, A. Vidal-Puig, The different shades of fat, Nature, 510 (2014) 76-83. [55] P. Tontonoz, B.M. Spiegelman, Fat and beyond: the diverse biology of PPARgamma, Annu Rev Biochem, 77 (2008) 289-312. [56] M.I. Lefterova, Y. Zhang, D.J. Steger, M. Schupp, J. Schug, A. Cristancho, D. Feng, D. Zhuo, C.J. Stoeckert, Jr., X.S. Liu, M.A. Lazar, PPARgamma and C/EBP factors orchestrate adipocyte biology via adjacent binding on a genome-wide scale, Genes & development, 22 (2008) 2941-2952. [57] Q.A. Wang, C. Tao, L. Jiang, M. Shao, R. Ye, Y. Zhu, R. Gordillo, A. Ali, Y. Lian, W.L. Holland, R.K. Gupta, P.E. Scherer, Distinct regulatory mechanisms governing embryonic versus adult adipocyte maturation, Nature cell biology, 17 (2015) 1099-1111. [58] F. Wang, S.E. Mullican, J.R. DiSpirito, L.C. Peed, M.A. Lazar, Lipoatrophy and severe metabolic disturbance in mice with fat-specific deletion of PPARgamma, Proc Natl Acad Sci U S A, 110 (2013) 18656-18661. [59] D. Bishop-Bailey, J. Wray, Peroxisome proliferator-activated receptors: a critical review on endogenous pathways for ligand generation, Prostaglandins & other lipid mediators, 71 (2003) 1-22. [60] Y. Oishi-Tanaka, C.K. Glass, A new role for cyclic phosphatidic acid as a PPARgamma antagonist, Cell Metab, 12 (2010) 207-208.

33

ACCEPTED MANUSCRIPT [61] W.L. Chou, L.M. Chuang, C.C. Chou, A.H. Wang, J.A. Lawson, G.A. FitzGerald, Z.F. Chang, Identification of a novel prostaglandin reductase reveals the involvement of prostaglandin E2 catabolism in regulation of peroxisome proliferator-activated receptor gamma activation, The Journal of biological chemistry, 282 (2007) 18162-18172. [62] T. Tsukahara, R. Tsukahara, Y. Fujiwara, J. Yue, Y. Cheng, H. Guo, A. Bolen, C. Zhang, L. Balazs, F. Re, G. Du, M.A.

T

Frohman, D.L. Baker, A.L. Parrill, A. Uchiyama, T. Kobayashi, K. Murakami-Murofushi, G. Tigyi, Phospholipase D2-dependent inhibition of the nuclear hormone receptor PPARgamma by cyclic phosphatidic acid, Molecular cell, 39

IP

(2010) 421-432.

[63] P. Tontonoz, E. Hu, B.M. Spiegelman, Stimulation of adipogenesis in fibroblasts by PPAR gamma 2, a lipid-activated

SC R

transcription factor, Cell, 79 (1994) 1147-1156.

[64] N. Petrovic, T.B. Walden, I.G. Shabalina, J.A. Timmons, B. Cannon, J. Nedergaard, Chronic peroxisome proliferator-activated receptor gamma (PPARgamma) activation of epididymally derived white adipocyte cultures reveals a population of thermogenically competent, UCP1-containing adipocytes molecularly distinct from classic brown

NU

adipocytes, The Journal of biological chemistry, 285 (2010) 7153-7164.

[65] J.E. Digby, C.T. Montague, C.P. Sewter, L. Sanders, W.O. Wilkison, S. O'Rahilly, J.B. Prins, Thiazolidinedione exposure increases the expression of uncoupling protein 1 in cultured human preadipocytes, Diabetes, 47 (1998) 138-141.

MA

[66] E. Hu, J.B. Kim, P. Sarraf, B.M. Spiegelman, Inhibition of adipogenesis through MAP kinase-mediated phosphorylation of PPARgamma, Science, 274 (1996) 2100-2103. [67] A. Garg, Clinical review#: Lipodystrophies: genetic and acquired body fat disorders, The Journal of clinical endocrinology and metabolism, 96 (2011) 3313-3325.

D

[68] T. Nolis, Exploring the pathophysiology behind the more common genetic and acquired lipodystrophies, Journal of

TE

human genetics, 59 (2014) 16-23.

[69] W.V. Brown, A. Garg, P. Gorden, R. Shamburek, JCL roundtable: Diagnosis and clinical management of lipodystrophy, Journal of clinical lipidology, 10 (2016) 728-736.

CE P

[70] M. Peterfy, J. Phan, P. Xu, K. Reue, Lipodystrophy in the fld mouse results from mutation of a new gene encoding a nuclear protein, lipin, Nat Genet, 27 (2001) 121-124. [71] P. Zhang, K. Takeuchi, L.S. Csaki, K. Reue, Lipin-1 phosphatidic phosphatase activity modulates phosphatidate levels to promote peroxisome proliferator-activated receptor gamma (PPARgamma) gene expression during adipogenesis, The

AC

Journal of biological chemistry, 287 (2012) 3485-3494. [72] J. Magre, M. Delepine, E. Khallouf, T. Gedde-Dahl, Jr., L. Van Maldergem, E. Sobel, J. Papp, M. Meier, A. Megarbane, A. Bachy, A. Verloes, F.H. d'Abronzo, E. Seemanova, R. Assan, N. Baudic, C. Bourut, P. Czernichow, F. Huet, F. Grigorescu, M. de Kerdanet, D. Lacombe, P. Labrune, M. Lanza, H. Loret, F. Matsuda, J. Navarro, A. Nivelon-Chevalier, M. Polak, J.J. Robert, P. Tric, N. Tubiana-Rufi, C. Vigouroux, J. Weissenbach, S. Savasta, J.A. Maassen, O. Trygstad, P. Bogalho, P. Freitas, J.L. Medina, F. Bonnicci, B.I. Joffe, G. Loyson, V.R. Panz, F.J. Raal, S. O'Rahilly, T. Stephenson, C.R. Kahn, M. Lathrop, J. Capeau, Identification of the gene altered in Berardinelli-Seip congenital lipodystrophy on chromosome 11q13, Nat Genet, 28 (2001) 365-370. [73] A. Garg, Acquired and inherited lipodystrophies, The New England journal of medicine, 350 (2004) 1220-1234. [74] C. Windpassinger, M. Auer-Grumbach, J. Irobi, H. Patel, E. Petek, G. Horl, R. Malli, J.A. Reed, I. Dierick, N. Verpoorten, T.T. Warner, C. Proukakis, P. Van den Bergh, C. Verellen, L. Van Maldergem, L. Merlini, P. De Jonghe, V. Timmerman, A.H. Crosby, K. Wagner, Heterozygous missense mutations in BSCL2 are associated with distal hereditary motor neuropathy and Silver syndrome, Nat Genet, 36 (2004) 271-276. [75] C. Lundin, R. Nordstrom, K. Wagner, C. Windpassinger, H. Andersson, G. von Heijne, I. Nilsson, Membrane topology of the human seipin protein, FEBS Lett, 580 (2006) 2281-2284. [76] A.K. Agarwal, A. Garg, Seipin: a mysterious protein, Trends in molecular medicine, 10 (2004) 440-444. [77] H. Kim, K. Melen, G. von Heijne, Topology models for 37 Saccharomyces cerevisiae membrane proteins based on

34

ACCEPTED MANUSCRIPT C-terminal reporter fusions and predictions, The Journal of biological chemistry, 278 (2003) 10208-10213. [78] D. Ito, T. Fujisawa, H. Iida, N. Suzuki, Characterization of seipin/BSCL2, a protein associated with spastic paraplegia 17, Neurobiol Dis, 31 (2008) 266-277. [79] D. Binns, S. Lee, C.L. Hilton, Q.X. Jiang, J.M. Goodman, Seipin is a discrete homooligomer, Biochemistry, 49 (2010)

T

10747-10755. [80] M.F. Sim, M.M. Talukder, R.J. Dennis, S. O'Rahilly, J.M. Edwardson, J.J. Rochford, Analysis of naturally occurring

IP

mutations in the human lipodystrophy protein seipin reveals multiple potential pathogenic mechanisms, Diabetologia, 56 (2013) 2498-2506.

SC R

[81] A.R. Thiam, M. Beller, The why, when and how of lipid droplet diversity, Journal of cell science, 130 (2017) 315-324. [82] B.K. Straub, P. Stoeffel, H. Heid, R. Zimbelmann, P. Schirmacher, Differential pattern of lipid droplet-associated proteins and de novo perilipin expression in hepatocyte steatogenesis, Hepatology, 47 (2008) 1936-1946. [83] W. Fei, G. Alfaro, B.P. Muthusamy, Z. Klaassen, T.R. Graham, H. Yang, C.T. Beh, Genome-wide analysis of sterol-lipid

NU

storage and trafficking in Saccharomyces cerevisiae, Eukaryot Cell, 7 (2008) 401-414. [84] E. Boutet, H. El Mourabit, M. Prot, M. Nemani, E. Khallouf, O. Colard, M. Maurice, A.M. Durand-Schneider, Y. Chretien, S. Gres, C. Wolf, J.S. Saulnier-Blache, J. Capeau, J. Magre, Seipin deficiency alters fatty acid Delta9 desaturation

MA

and lipid droplet formation in Berardinelli-Seip congenital lipodystrophy, Biochimie, 91 (2009) 796-803. [85] X. Cui, Y. Wang, Y. Tang, Y. Liu, L. Zhao, J. Deng, G. Xu, X. Peng, S. Ju, G. Liu, H. Yang, Seipin ablation in mice results in severe generalized lipodystrophy, Human molecular genetics, 20 (2011) 3022-3030. [86] W. Chen, B. Chang, P. Saha, S.M. Hartig, L. Li, V.T. Reddy, Y. Yang, V. Yechoor, M.A. Mancini, L. Chan, Berardinelli-seip

D

congenital lipodystrophy 2/seipin is a cell-autonomous regulator of lipolysis essential for adipocyte differentiation,

TE

Molecular and cellular biology, 32 (2012) 1099-1111.

[87] X.G. Peng, S. Ju, F. Fang, Y. Wang, K. Fang, X. Cui, G. Liu, P. Li, H. Mao, G.J. Teng, Comparison of brown and white adipose tissue fat fractions in ob, seipin, and Fsp27 gene knockout mice by chemical shift-selective imaging and (1)H-MR

CE P

spectroscopy, American journal of physiology. Endocrinology and metabolism, 304 (2013) E160-167. [88] L. Liu, Q. Jiang, X. Wang, Y. Zhang, R.C. Lin, S.M. Lam, G. Shui, L. Zhou, P. Li, Y. Wang, X. Cui, M. Gao, L. Zhang, Y. Lv, G. Xu, G. Liu, D. Zhao, H. Yang, Adipose-specific knockout of SEIPIN/BSCL2 results in progressive lipodystrophy, Diabetes, 63 (2014) 2320-2331.

AC

[89] M. Jiang, M. Gao, C. Wu, H. He, X. Guo, Z. Zhou, H. Yang, X. Xiao, G. Liu, J. Sha, Lack of testicular seipin causes teratozoospermia syndrome in men, Proc Natl Acad Sci U S A, 111 (2014) 7054-7059. [90] X. Prieur, L. Dollet, M. Takahashi, M. Nemani, B. Pillot, C. Le May, C. Mounier, H. Takigawa-Imamura, D. Zelenika, F. Matsuda, B. Feve, J. Capeau, M. Lathrop, P. Costet, B. Cariou, J. Magre, Thiazolidinediones partially reverse the metabolic disturbances observed in Bscl2/seipin-deficient mice, Diabetologia, 56 (2013) 1813-1825. [91] M. Shao, J. Ishibashi, C.M. Kusminski, Q.A. Wang, C. Hepler, L. Vishvanath, K.A. MacPherson, S.B. Spurgin, K. Sun, W.L. Holland, P. Seale, R.K. Gupta, Zfp423 Maintains White Adipocyte Identity through Suppression of the Beige Cell Thermogenic Gene Program, Cell Metab, 23 (2016) 1167-1184. [92] S.Y. Choi, P. Huang, G.M. Jenkins, D.C. Chan, J. Schiller, M.A. Frohman, A common lipid links Mfn-mediated mitochondrial fusion and SNARE-regulated exocytosis, Nature cell biology, 8 (2006) 1255-1262. [93] N. Vitale, A.S. Caumont, S. Chasserot-Golaz, G. Du, S. Wu, V.A. Sciorra, A.J. Morris, M.A. Frohman, M.F. Bader, Phospholipase D1: a key factor for the exocytotic machinery in neuroendocrine cells, The EMBO journal, 20 (2001) 2424-2434. [94] S. Han, D.D. Binns, Y.F. Chang, J.M. Goodman, Dissecting seipin function: the localized accumulation of phosphatidic acid at ER/LD junctions in the absence of seipin is suppressed by Sei1p(DeltaNterm) only in combination with Ldb16p, BMC cell biology, 16 (2015) 29. [95] H. Wolinski, H.F. Hofbauer, K. Hellauer, A. Cristobal-Sarramian, D. Kolb, M. Radulovic, O.L. Knittelfelder, G.N.

35

ACCEPTED MANUSCRIPT Rechberger, S.D. Kohlwein, Seipin is involved in the regulation of phosphatidic acid metabolism at a subdomain of the nuclear envelope in yeast, Biochimica et biophysica acta, 1851 (2015) 1450-1464. [96] Y. Tian, J. Bi, G. Shui, Z. Liu, Y. Xiang, Y. Liu, M.R. Wenk, H. Yang, X. Huang, Tissue-autonomous function of Drosophila seipin in preventing ectopic lipid droplet formation, PLoS genetics, 7 (2011) e1001364.

T

[97] M.F. Sim, R.J. Dennis, E.M. Aubry, N. Ramanathan, H. Sembongi, V. Saudek, D. Ito, S. O'Rahilly, S. Siniossoglou, J.J. Rochford, The human lipodystrophy protein seipin is an ER membrane adaptor for the adipogenic PA phosphatase lipin 1,

IP

Molecular metabolism, 2 (2012) 38-46.

[98] K. Nadra, J.J. Medard, J.D. Mul, G.S. Han, S. Gres, M. Pende, D. Metzger, P. Chambon, E. Cuppen, J.S. Saulnier-Blache,

SC R

G.M. Carman, B. Desvergne, R. Chrast, Cell autonomous lipin 1 function is essential for development and maintenance of white and brown adipose tissue, Molecular and cellular biology, 32 (2012) 4794-4810. [99] L.S. Csaki, J.R. Dwyer, L.G. Fong, P. Tontonoz, S.G. Young, K. Reue, Lipins, lipinopathies, and the modulation of cellular lipid storage and signaling, Progress in lipid research, 52 (2013) 305-316.

NU

[100] G.S. Han, W.I. Wu, G.M. Carman, The Saccharomyces cerevisiae Lipin homolog is a Mg2+-dependent phosphatidate phosphatase enzyme, The Journal of biological chemistry, 281 (2006) 9210-9218. [101] A.K. Agarwal, V. Simha, E.A. Oral, S.A. Moran, P. Gorden, S. O'Rahilly, Z. Zaidi, F. Gurakan, S.A. Arslanian, A. Klar, A.

MA

Ricker, N.H. White, L. Bindl, K. Herbst, K. Kennel, S.B. Patel, L. Al-Gazali, A. Garg, Phenotypic and genetic heterogeneity in congenital generalized lipodystrophy, The Journal of clinical endocrinology and metabolism, 88 (2003) 4840-4847. [102] C.A. Nagle, E.L. Klett, R.A. Coleman, Hepatic triacylglycerol accumulation and insulin resistance, Journal of lipid research, 50 Suppl (2009) S74-79.

D

[103] A.K. Agarwal, Lysophospholipid acyltransferases: 1-acylglycerol-3-phosphate O-acyltransferases. From discovery to

TE

disease, Current opinion in lipidology, 23 (2012) 290-302. [104] M.M. Talukder, M.F. Sim, S. O'Rahilly, J.M. Edwardson, J.J. Rochford, Seipin oligomers can interact directly with AGPAT2 and lipin 1, physically scaffolding critical regulators of adipogenesis, Molecular metabolism, 4 (2015) 199-209.

CE P

[105] H. Wolinski, D. Kolb, S. Hermann, R.I. Koning, S.D. Kohlwein, A role for seipin in lipid droplet dynamics and inheritance in yeast, Journal of cell science, 124 (2011) 3894-3904. [106] Y. Qi, W. Wang, J. Chen, L. Dai, D. Kaczorowski, X. Gao, P. Xia, Sphingosine Kinase 1 Protects Hepatocytes from Lipotoxicity via Down-regulation of IRE1alpha Protein Expression, The Journal of biological chemistry, 290 (2015)

AC

23282-23290.

[107] Y. Qi, P. Xia, Cellular inhibitor of apoptosis protein-1 (cIAP1) plays a critical role in beta-cell survival under endoplasmic reticulum stress: promoting ubiquitination and degradation of C/EBP homologous protein (CHOP), The Journal of biological chemistry, 287 (2012) 32236-32245. [108] D. Mekahli, G. Bultynck, J.B. Parys, H. De Smedt, L. Missiaen, Endoplasmic-reticulum calcium depletion and disease, Cold Spring Harbor perspectives in biology, 3 (2011). [109] J. Bi, W. Wang, Z. Liu, X. Huang, Q. Jiang, G. Liu, Y. Wang, X. Huang, Seipin promotes adipose tissue fat storage through the ER Ca(2)(+)-ATPase SERCA, Cell Metab, 19 (2014) 861-871. [110] S. Fu, L. Yang, P. Li, O. Hofmann, L. Dicker, W. Hide, X. Lin, S.M. Watkins, A.R. Ivanov, G.S. Hotamisligil, Aberrant lipid metabolism disrupts calcium homeostasis causing liver endoplasmic reticulum stress in obesity, Nature, 473 (2011) 528-531. [111] B.R. Cartwright, D.D. Binns, C.L. Hilton, S. Han, Q. Gao, J.M. Goodman, Seipin performs dissectible functions in promoting lipid droplet biogenesis and regulating droplet morphology, Molecular biology of the cell, 26 (2015) 726-739. [112] W. Chen, V.K. Yechoor, B.H. Chang, M.V. Li, K.L. March, L. Chan, The human lipodystrophy gene product Berardinelli-Seip congenital lipodystrophy 2/seipin plays a key role in adipocyte differentiation, Endocrinology, 150 (2009) 4552-4561. [113] V.A. Payne, N. Grimsey, A. Tuthill, S. Virtue, S.L. Gray, E. Dalla Nora, R.K. Semple, S. O'Rahilly, J.J. Rochford, The

36

ACCEPTED MANUSCRIPT human lipodystrophy gene BSCL2/seipin may be essential for normal adipocyte differentiation, Diabetes, 57 (2008) 2055-2060. [114] L. Dollet, J. Magre, B. Cariou, X. Prieur, Function of seipin: new insights from Bscl2/seipin knockout mouse models, Biochimie, 96 (2014) 166-172.

T

[115] E. Mori, J. Fujikura, M. Noguchi, K. Nakao, M. Matsubara, M. Sone, D. Taura, T. Kusakabe, K. Ebihara, T. Tanaka, K. Hosoda, K. Takahashi, I. Asaka, N. Inagaki, K. Nakao, Impaired adipogenic capacity in induced pluripotent stem cells from

IP

lipodystrophic patients with BSCL2 mutations, Metabolism: clinical and experimental, 65 (2016) 543-556. [116] H. Zhou, S.M. Black, T.W. Benson, N.L. Weintraub, W. Chen, Berardinelli-Seip Congenital Lipodystrophy 2/Seipin Is

SC R

Not Required for Brown Adipogenesis but Regulates Brown Adipose Tissue Development and Function, Molecular and cellular biology, 36 (2016) 2027-2038.

[117] L. Dollet, J. Magre, M. Joubert, C. Le May, A. Ayer, L. Arnaud, C. Pecqueur, V. Blouin, B. Cariou, X. Prieur, Seipin deficiency alters brown adipose tissue thermogenesis and insulin sensitivity in a non-cell autonomous mode, Scientific

NU

reports, 6 (2016) 35487.

[118] H. Zhou, X. Lei, T. Benson, J. Mintz, X. Xu, R.B. Harris, N.L. Weintraub, X. Wang, W. Chen, Berardinelli-Seip congenital lipodystrophy 2 regulates adipocyte lipolysis, browning, and energy balance in adult animals, Journal of lipid

MA

research, 56 (2015) 1912-1925.

[119] L. Dollet, C. Levrel, T. Coskun, S. Le Lay, C. Le May, A. Ayer, Q. Venara, A.C. Adams, R.E. Gimeno, J. Magre, B. Cariou, X. Prieur, FGF21 Improves the Adipocyte Dysfunction Related to Seipin Deficiency, Diabetes, 65 (2016) 3410-3417. [120] X. Cui, Y. Wang, L. Meng, W. Fei, J. Deng, G. Xu, X. Peng, S. Ju, L. Zhang, G. Liu, L. Zhao, H. Yang, Overexpression of a

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short human seipin/BSCL2 isoform in mouse adipose tissue results in mild lipodystrophy, American journal of physiology.

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Endocrinology and metabolism, 302 (2012) E705-713. [121] B. Victoria, J.M. Cabezas-Agricola, B. Gonzalez-Mendez, G. Lattanzi, R. Del Coco, L. Loidi, F. Barreiro, C. Calvo, J. Lado-Abeal, D. Araujo-Vilar, Reduced adipogenic gene expression in fibroblasts from a patient with type 2 congenital

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generalized lipodystrophy, Diabetic medicine : a journal of the British Diabetic Association, 27 (2010) 1178-1187. [122] J.M. Raboud, C. Diong, A. Carr, S. Grinspoon, K. Mulligan, J. Sutinen, W. Rozenbaum, R.B. Cavalcanti, H. Wand, D. Costagliola, S. Walmsley, A meta-analysis of six placebo-controlled trials of thiazolidinedione therapy for HIV lipoatrophy, HIV clinical trials, 11 (2010) 39-50.

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[123] M. Tungsiripat, D.E. Bejjani, N. Rizk, A. O'Riordan M, A.C. Ross, C. Hileman, N. Storer, D. Harrill, G.A. McComsey, Rosiglitazone improves lipoatrophy in patients receiving thymidine-sparing regimens, AIDS, 24 (2010) 1291-1298. [124] D. Shan, J.L. Li, L. Wu, D. Li, J. Hurov, J.F. Tobin, R.E. Gimeno, J. Cao, GPAT3 and GPAT4 are regulated by insulin-stimulated phosphorylation and play distinct roles in adipogenesis, Journal of lipid research, 51 (2010) 1971-1981.

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ACCEPTED MANUSCRIPT Figure legends Figure 1. The glycerol-3-phosphate pathway. Enzymes implicated in adipogenesis are highlighted in

O-acyltransferase;

CDS:

CDP-DAG

synthase;

GPAT,

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1-acylglycerol-3-phosphate

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green. SEIPIN/Fld1p/Sei1p may also regulate this pathway through inhibiting GPAT. AGPAT:

glycerol-3-phosphate acyltransferase; PAP, phosphatidic acid phosphatase.

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Figure 2. The schematic structures of yeast Fld1p/Sei1p and human SEIPIN. Missense mutations that

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cause generalized lipodystrophy and neuropathy are indicated in green, and red, respectively.

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Figure 3. Hypothesis. Phospholipids, such as phosphatidic acid, may regulate both adipogenesis and

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the expansion of lipid droplets. A loss of AGPAT2, CDS1, lipin-1 or possibly SEIPIN function can

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increase the amount of phosphatidic acid at the endoplasmic reticulum and nuclear envelope, which may antagonize PPARγ function in differentiating preadipocytes, thereby blocking adipogenesis. In

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all cell types including yeast, the increased phosphatidic acid may affect the budding, growth and/or fusion of lipid droplets.

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Figure 1

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Figure 2

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Figure 3

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ACCEPTED MANUSCRIPT Highlights

Phospholipids regulate both lipid droplets growth and adipogenesis

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Enzymes regulating phosphatidic acid metabolism are implicated in adipogenesis

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SEIPIN regulates the metabolism and/or distribution of phosphatidic acid

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