Seipin-Mediated Contacts as Gatekeepers of Lipid Flux at the Endoplasmic Reticulum

Abstract

Lipid droplets (LDs) are dynamic cellular hubs of lipid metabolism. While LDs contact a plethora of organelles, they have the most intimate relationship with the endoplasmic reticulum (ER). Indeed, LDs are initially assembled at specialized ER subdomains, and recent work has unraveled an increasing array of proteins regulating ER-LD contacts. Among these, seipin, a highly conserved lipodystrophy protein critical for LD growth and adipogenesis, deserves special attention. Here, we review recent insights into the role of seipin in LD biogenesis and as a regulator of ER-LD contacts. These studies have also highlighted the evolving concept of ER and LDs as a functional continuum for lipid partitioning and pinpointed a role for seipin at the ER-LD nexus in controlling lipid flux between these compartments.

Keywords

seipin endoplasmic reticulum–lipid droplet contacts lipid droplet biogenesis

Lipid droplets (LDs) are intracellular storage organelles composed of a core of hydrophobic neutral lipids (NLs), mainly triglycerides (TAG) and sterol esters, surrounded by a phospholipid (PL) monolayer (Henne et al., 2018). Their main function is to store excess energy in the form of TAGs, but LDs also play instrumental roles in many other cellular processes, including maintaining endoplasmic reticulum (ER) homeostasis (Welte, 2015; Olzmann and Carvalho, 2019). LDs are formed in the ER, and they retain a close physical and functional relationship with it after their formation. The lipid and protein fluxes between these compartments are controlled at ER-LD contacts. Recent studies have begun to shed light on the properties of LD forming ER subdomains and ER-LD contacts. In this review, we discuss recent insights into LD biogenesis and ER-LD contacts with a special focus on the lipodystrophy protein seipin.

Seipin Structure Gives Clues to Its Function

Seipin in an oligomeric ER transmembrane (TM) protein originally identified to be mutated in Berardinelli–Seip Congenital Lipodistrophy Type 2 (BSCL2) congenital lipodystrophy (Magré et al., 2001). Distinct mutations in seipin also result in hereditary spastic paraplegias (Windpassinger et al., 2004) and a severe form of encephalopathy (Guillén-Navarro et al., 2013). Seipin is evolutionary conserved, and perturbation of seipin or its homologs results in aberrant LD morphology in numerous model systems (Szymanski et al., 2007; Fei et al., 2008; Boutet et al., 2009; Cai et al., 2015; Salo et al., 2016; Wang et al., 2016; Kornke and Maniak, 2017; Coradetti et al., 2018; Cao et al., 2019). In accordance with its role in lipodystrophy syndrome in humans, seipin is essential for adipogenesis in various organisms (Payne et al., 2008; Chen et al., 2009; Victoria et al., 2010; Cui et al., 2011; Tian et al., 2011; Ebihara et al., 2015; Mori et al., 2016).

The LD phenotype in the absence of seipin is consistent among examined systems, with a proliferation of tiny and some supersized LDs in an apparently stochastic manner (Cartwright and Goodman, 2012; Chen and Goodman, 2017). Several hypotheses have been proposed for seipin in relation to LDs: a role in controlling ER PL metabolism, sequestering NLs or regulating ER-LD contact sites. These hypotheses are not mutually exclusive, and the structure of seipin is in principle compatible with any of these. In addition to its role in LD formation, seipin has been linked to cytoskeleton remodeling, calcium handling, and lipolysis (Chen et al., 2012; Bi et al., 2014; Yang et al., 2014; Ding et al., 2018; Li et al., 2019b) with a growing number of identified interaction partners (Table 1).

Table 1.

Seipin Interacting Proteins.

Protein name	Protein function	Main evidence for interaction	Proposed function of interaction	Reference
Ldb16	Regulator of LD morphology	Co-ip: endogenous proteins in yeast	Forms a complex with seipin to regulate LDs	Cartwright et al. (2015), Wang et al. (2014), and Wolinski et al. (2015)
Promethin/LDAF1/Ldo45 Ldo16	Regulator of LD identity (yeast)	Co-ip: endogenous proteins (yeast and human cells), comigration of endogenously tagged proteins (human cells)	Forms a complex with seipin to regulate LDs	Castro et al. (2019), Chung et al. (2019), Eisenberg-Bord et al. (2018), and Teixeira et al. (2018)
GPAT3-4/Gat1-2	Catalyzes G3P to LPA conversion	Co-ip: endogenous proteins (yeast), overexpressed proteins (murine cells)	Seipin negatively regulates GPAT enzyme activity	Pagac et al. (2016) and Sim et al. (2020)
SERCA	ER calcium pump, transports Ca²⁺ into ER lumen	Co-ip: endogenous proteins (human cells), overexpressed proteins (Drosophila cells)	Seipin positively regulates SERCA activity	Bi et al. (2014)
Lipin-1	Catalyzes PA to DAG conversion	Co-ip and AFM: overexpressed proteins (murine cells)	Seipin recruits lipin to the ER membrane	Sim et al. (2012) and Talukder et al. (2015)
AGPAT-2	Catalyzes LPA to PA conversion	Co-ip and AFM: overexpressed proteins (mice cells)	Scaffolding of lipogenic enzymes	Sim et al. (2012) and Talukder et al. (2015)
Serine palmitoyltransferaseComplex, FA-elongase Tsc13	Sphingolipid synthesis	Co-ip and BiFic of overexpressed protein (yeast)	Seipin negatively regulates sphingolipid production	Su et al. (2019)
14-3-3-β	A scaffolding protein, signal transduction	Co-ip: overexpressed proteins (murine and human cells)	Cytoskeleton remodeling	Yang et al. (2014)
Perilipin-2	LD coat protein, lipolysis regulator	Co-ip: overexpressed proteins (human cells)	Seipin regulates perilipin-2 localization to LDs	Mori et al. (2016)
Perilipin-1	LD coat protein, lipolysis regulator	Co-ip and AFM: overexpressed proteins (human cells)	Seipin regulates perilipin-1 localization to LDs	Jiao et al. (2019)
Reep-1	Regulates ER morphology	Co-ip: overexpressed proteins (murine cells)	None proposed	Renvoisé et al. (2016)

ER = endoplasmic reticulum; AGPAT = 1-acylglycerol-3-phosphate-O-acyltransferase; GPAT = Glycerol-3-phosphate acyltransferase; SERCA = sarco/endoplasmic reticulum Ca²⁺-ATPase ; FA = Fatty acid; G3P = Glycerol 3-phosphate; LPA =Lysophosphatidic acid; DAG = diacylglycerol; PA = phosphatidic acid; LD = lipid droplet; AFM = Atomic force microscopy.

Structurally seipin contains two transmembrane domains, an evolutionary conserved ER luminal loop and variable N- and C-terminal cytoplasmic regions (Lundin et al., 2006; Yang et al., 2013). The luminal loop and its flanking two transmembrane domains are critical for seipin function in LD biogenesis, as they alone are sufficient to rescue defective LD formation in seipin-deficient cells (Yang et al., 2013; Wang et al., 2016). Seipin forms homo-oligomers, each containing 11 (human) or 12 (Drosophila) seipin monomers (Binns et al., 2010; Sim et al., 2012; Sui et al., 2018; Yan et al., 2018). In yeast, seipin forms foci localizing to ER-LD contact sites (Szymanski et al., 2007; Fei et al., 2008). In mammalian cells, endogenously fluorescently tagged seipin appears as homogenous, discrete, and mobile foci in the ER, a subset of which are stably associated with ER-LD contacts (Salo et al., 2016; Wang et al., 2016). Seipin oligomerization is critical for its function, as evidenced by the fact that the lipodystrophy-associated A212P-seipin mutant forms smaller oligomers and fails to rescue the aberrant LD biogenesis of seipin-deficient cells (Binns et al., 2010; Sim et al., 2013; Salo et al., 2016). The same holds true for an oligomerization deficient mutant designed based on the cryo-electron microscopy (EM) structure of seipin (Yan et al., 2018).

The structures of the luminal domain of human and Drosophila seipins have been solved by cryo-EM (Sui et al., 2018; Yan et al., 2018; Figure 1). They form a ring-like/disk-like structure of circa 15 nm in diameter, with 11 (human) or 12 (Drosophila) seipin monomers closely intertwined. Each monomer displays two notable features: several hydrophobic helices (HHs) in very close proximity to the bilayer membrane, and two beta sheets that exhibit an overall fold similar to certain lipid binding domains (Sui et al., 2018; Yan et al., 2018). The structure of the TM or cytoplasmic regions could not be solved by cryo-EM in these studies, suggesting conformational structural flexibility therein.

Figure 1.

Seipin Oligomer Structure. The TM regions were modeled into the cryo-EM structure of the luminal domains of human seipin (PDB: 6ds5; Yan et al., 2018) using PyMol software. ER = endoplasmic reticulum.

The ER luminal HHs of seipin are able to bind to a monolayer covering NLs in vitro (Sui et al., 2018) and appear to be widely conserved across species (Chapman et al., 2019). When expressed as individual peptides in cells, the HHs localize to LDs. This localization is impaired by mutating three hydrophobic residues to aspartic acid (Sui et al., 2018). A mutant seipin with corresponding amino acid substitutions still rescues seipin function in LD biogenesis, but further concomitant deletion of the N-terminus of seipin, shown previously to localize to LDs when expressed alone (Wang et al., 2016), results in a dysfunctional seipin. Hence, the presence of either LD-targeting luminal HHs or the cytoplasmic N-terminus is required for seipin function. Considering that the LD PL monolayer harbors packing defects (Bacle et al., 2017; Prévost et al., 2018; Chorlay and Thiam, 2020), these seipin helices might recognize similar packing defects induced by NL lenses in the ER (Sui et al., 2018).

The bulk of the seipin luminal region forms a beta-sandwich-like fold, structurally similar to many lipid-binding domains, including those of Niemann-Pick C2 (NPC2) and PKC (Sui et al., 2018; Yan et al., 2018).This suggests that seipin may bind lipids in the ER lumen. In vitro work suggests that both purified full-length seipin and a truncated variant containing only the putative lipid binding domain can bind anionic PLs, such as phosphatidic acid (PA; Sui et al., 2018). A mutant harboring amino acid substitutions in the hydrophobic cavity of the beta sandwich was unable to complement the seipin knockout (KO) LD phenotype (Sui et al., 2018). However, which lipids seipin interacts with in intact cells remains to be determined.

Seipin Forms an LD Assembly Complex

The most extensively studied binding partners of seipin are Ldb16 and Ldo16 and its splicing isoform Ldo 45/promethin (Table 1). In yeast, seipin forms a stable complex with Ldb16, an ER TM protein with no known mammalian homologues.Seipin is required for Ldb16 stabilization, and depletion of either Ldb16 or seipin results in indistinguishable LD phenotypes (Wang et al., 2014; Grippa et al., 2015). These LD defects can be rescued by heterologous expression of human seipin, strongly suggesting that human seipin functionally corresponds to the yeast Ldb16-seipin complex.

Yeast ER-LD-localized protein isoforms Ldo16 and Ldo45 were recently discovered to be interactors of the seipin complex (Eisenberg-Bord et al., 2018; Teixeira et al., 2018). Overexpression of Ldo45 leads to proliferation and clustering of LDs that exhibit aberrant protein targeting. This phenotype is similar to seipin KO cells, suggesting that Ldo45 and seipin may play antagonistic roles in LD formation and maintenance (Eisenberg-Bord et al., 2018; Teixeira et al., 2018). Ldo16 and Ldo45 were also found to be regulators of a subpopulation of LDs that reside in close proximity to the yeast nucleus-vacuole junctions (NVJs).During nutritional stress, this site expands and serves as a site for LD biogenesis under these conditions (Hariri et al., 2018).Indeed, the relative abundance of Ldo16/Ldo45 is regulated by cellular growth conditions, and Ldo45 is necessary for recruiting Pdr16, a lipid transfer protein, to NVJ-associated LDs (Eisenberg-Bord et al., 2018; Teixeira et al., 2018).

Based on remote homology searches, promethin (also known asTMEM159 and recently renamed as Lipid Droplet Assembly Factor 1; LDAF1 ) was proposed as a homolog of Ldo45 (Eisenberg-Bord et al., 2018). Promethin is upregulated during adipogenesis, localizes to the ER and LDs, and was found to interact with seipin (Castro et al., 2019). When expressed heterologously in yeast, human promethin was able to co-immunoprecipitate the yeast seipin complex, suggesting an evolutionary conserved interaction (Castro et al., 2019). In human cells, endogenously tagged promethin forms distinct foci in the ER and a high fraction of these comigrate with seipin foci (Chung et al., 2019). LDs were found to be formed at sites typically occupied by both seipin and promethin. Upon subsequent growth of the LDs, promethin relocates to LD surface, due to hairpin topology allowing both ER and LD localization, while seipin remains at the ER-LD junction (Chung et al., 2019).

ER Architecture Promoting LD Formation

LD biogenesis is a complex stepwise process in the ER (Pol et al., 2014). Here, only a brief description of the currently prevailing model of LD biogenesis is presented. For detailed discussion, we kindly refer the reader to recent comprehensive reviews (Chen and Goodman, 2017; Walther et al., 2017; Chapman et al., 2019; Gao et al., 2019b; Jackson, 2019; Henne et al., 2020; Renne et al., 2020).

During LD assembly, TAGs synthesized by ER-resident diacylglycerol acyltransferase (DGAT) enzymes first accumulate between the ER bilayer leaflets. However, as PL bilayers can only accommodate minute concentrations of TAG, rising local concentrations will lead to the formation of nm-sized TAG clusters through nucleation and phase separation. These form lens-like structures as predicted by simulations and observed in yeast by EM (Khandelia et al., 2010; Choudhary et al., 2015). Subsequent growth of TAG aggregates will lead to deformation of the ER bilayer and budding out of a nascent LD, with the monolayer enclosing the LD derived from the cytoplasmic leaflet of the ER (Kassan et al., 2013). These early steps are modulated by the ER PL composition via effects on ER surface tension and PL intrinsic curvature, while the directionality of LD budding to the cytosol may arise due to ER membrane asymmetry (Ben M’barek et al., 2017; Choudhary et al., 2018; Chorlay et al., 2019; Zoni et al., 2020). It therefore seems likely that the LD forming ER subdomain harbors a distinct PL milieu, although direct evidence for this is lacking.

LD formation may also be controlled by the overall ER architecture. LD biogenesis appears to occur at peripheral ER tubules in mammalian cells (Kassan et al., 2013; Joshi et al., 2018; Santinho et al., 2020). In accordance, many ER shaping proteins have been implicated in LD formation (Klemm et al., 2013; Falk et al., 2014; Papadopoulos et al., 2015). Conversely, proteins participating in LD biogenesis such as Rab18 and fat storage-inducing transmembrane 2 (FIT2) have been shown to impact ER morphology (Gerondopoulos et al., 2014; Hayes et al., 2017; Becuwe et al., 2018).Changes in ER ultrastructure have been observed also in seipin-deficient cells (Grippa et al., 2015; Salo et al., 2016), and many proposed seipin collaborators, such as Pex30/MCTP2, Ldo45/Promethin, Reep-1, and Aradopsis LDIP (Table 1), have been suggested to act as ER curvature sensors/inducers (Renvoisé et al., 2016; Pyc et al., 2017; Eisenberg-Bord et al., 2018; Joshi et al., 2018; Teixeira et al., 2018; Wang et al., 2018; Chung et al., 2019).

The relationship between ER morphology and LD nucleation was recently investigated (Santinho et al., 2020). LDs were found to be preferentially generated at ER tubules both in the presence and absence of seipin. Experiments in cells and model membranes suggested that the presence of free TAG in highly curved membranes (such as ER tubules) is energetically unfavorable compared with flat regions (such as ER sheets). This leads to either outflow of TAGs from tubules or their condensation into LDs. Accordingly, LD nucleation can be achieved in vitro by increasing membrane curvature. Together, these data suggest that the ER membrane curvature can catalyze LD assembly (Santinho et al., 2020).

A major challenge in deciphering the dynamics and kinetics of LD biogenesis is the small size and possible instability of the formed NL clusters. Indeed, while ∼40- to 60-nm sized lens-like ER-embedded structures were detected in yeast (Choudhary et al., 2015), LD-like structures in the size range of ∼30 nm were observed at seipin-marked sites in human A431 cells during LD assembly (Salo et al., 2019).Circa 40 to 50 nm vesicular structures marked by LiveDrop were detected in COS-7 cells (Li et al., 2019a), and LDs in this size range (although not necessarily made de novo) were also recently observed by cryo-electron tomography (Mahamid et al., 2019).Thus, the size limit of the smallest forming LDs is yet unknown and may vary between cell types.

Seipin Defines LD Formation Sites

Several lines of evidence suggest that seipin is a key player in early LD biogenesis and may define the ER subdomain primed for LD formation. Seipin foci are localized at LD forming sites in the ER during the earliest observable steps of LD biogenesis (Wang et al., 2016; Salo et al., 2019; Choudhary et al., 2020).Fluorescently labeled model ER-LD peptides harboring LD-targeting motifs, such as HPos and LiveDrop (Wang et al., 2016), have been used to visualize nascent LDs prior to their detection with traditional hydrophobic dyes such as BODIPY. In our hands, the bright lipophilic dye LD540 (Spandl et al., 2009) stains nascent LDs at least as early as HPos or LiveDrop. However, it is not clear whether any of these markers detect lenses or only budded-out LDs.

In A431 cells, seipin motility was found to be decreased prior to accumulation of LD540, LiveDrop, or endogenously tagged acyl-CoA synthetase 3 (ACSL3) (Salo et al., 2019), suggesting that seipin is stabilized at LD-forming sites prior to other known actors. Similar results were reported in SUM159 cells, where LDs marked by endogenously tagged Perilipin-3 emerged at seipin-defined sites (Chung et al., 2019). Seipin can also spatially define LD formation sites in the ER. Relocalization of seipin to a subdomain of the ER, the nuclear envelope, was sufficient to relocate LD biogenesis to this new site (Salo et al., 2019). Analogous results were reported when seipin was relocalized to ER-plasma membrane junctions (Chung et al., 2019). These observations not only demonstrate that seipin can define where a LD starts to develop but also suggest considerable spatial malleability in LD formation sites.

Seipin and Regulation of ER PL Metabolism

Local PL levels, especially those of PA and DAG, have been proposed to be vital for LD emergence (Adeyo et al., 2011; Cartwright et al., 2015; Wolinski et al., 2015; Ben M’barek et al., 2017; Choudhary et al., 2018). It is conceivable that seipin somehow directly modulates local PLs at LD forming sites (Yan et al., 2018). In line with this, LD budding is almost completely abolished in a yeast mutant harboring double deletion of seipin and a putative ER-shaping protein Pex30, resulting in accumulation of toxic levels of TAG in the ER (Wang et al., 2018). Moreover, this phenotype was partially restored by modulating the ER PL composition toward permissive for LD budding. However, codepletion of seipin and Pex30’s putative mammalian homologue, MCTP2 in human cells did not display the same phenotype, arguing for more redundancy in mammalian systems (Joshi et al., 2018).

Seipin might control ER PL metabolism, especially that of PA, via protein–protein interactions of major lipid synthesizing enzymes, such as AGPAT2, lipin-1, and GPATs (Sim et al., 2012; Talukder et al., 2015; Pagac et al., 2016; Gao et al., 2019b; Sim et al., 2020). Indeed, GPAT activity was altered in a number of seipin depleted systems, suggesting that seipin negatively regulates GPAT activity to inhibit aberrant expansion of LDs (Pagac et al., 2016). In line with this, defective adipogenesis due to seipin-depletion was partially restored by GPAT inhibition (Pagac et al., 2016; Gao et al., 2020). However, in a recent study, GPAT3 deficiency failed to rescue adipogenesis in seipin-deficient differentiating adipocytes (Sim et al., 2020). Based on these findings, the significance of seipin-GPAT interaction remains to be determined, but increasing evidence suggests that seipin can act as a scaffold for lipogenic enzymes.

In the absence of seipin, PA has been suggested to accumulate in the ER and play an inhibitory role on adipogenic transcriptional program (Liu et al., 2014; Gao et al., 2019b). This notion is supported by the observation of increased PA levels in several seipin depleted systems (Fei et al., 2011; Tian et al., 2011; Jiang et al., 2014; Liu et al., 2014; Pagac et al., 2016; Cao et al., 2019). However, other studies have failed to detect differences in cellular PA levels even in the presence of a robust LD phenotype, arguing that global PA handling may not be the culprit in seipin deficiency (Grippa et al., 2015; Wang et al., 2016). Rather, seipin might alter PA metabolism at a distinct subdomain of the ER. This would be in line with the observation that PA-sensing fluorescent probes accumulate at LD-associated ER foci in yeast seipin KO cells, suggesting localized PA accumulation (Grippa et al., 2015; Han et al., 2015; Wolinski et al., 2015).

In this regard, the putative PA-binding site of seipin is in the ER lumen (Yan et al., 2018) proposes a topological puzzle. While at least some of the PA-generating GPAT and AGPAT enzymes may have their active sites in the ER lumen or within the ER bilayer (Yamashita et al., 2014), the PA-utilizing lipin enzymes are soluble, cytosolic proteins (Zhang and Reue, 2017), which would necessitate PA flipping to access seipin. Intriguingly, there are indications that FIT2 might act as a lipid phosphate phosphatase in the ER luminal side (Hayes et al., 2017; Becuwe et al., 2018) and that the protein is localized at seipin-marked LD formation sites (Choudhary et al., 2020). In this scenario, seipin might function to trap PA to promote its efficient catalysis into DAG at the ER-LD contact. However, this view is challenged by the lack of robust defects in lipid biosynthesis upon acute seipin loss (Wang et al., 2016; Salo et al., 2019) and that PA appears to locally accumulate in some seipin-deficient systems.

Besides PA, studies have also linked seipin to sphingolipid metabolism and defects in acyl chain saturation (Boutet et al., 2009; Amine et al., 2017; Su et al., 2019). Further work is needed to define how these relate to the LD biogenesis and adipogenesis defects of seipin loss. The recent discovery that polyunsaturated fatty acids were found to facilitate seipin recruitment to ER-LD contacts in C. elegans is highly interesting in this regard (Cao et al., 2019).

Seipin May Sequester TAGs at LD Formation Sites

Emerging studies suggest that seipin may directly control TAG distribution in the ER and its partitioning into nascent LDs. In a yeast system where NL synthesis was held constant, LD biogenesis was severely delayed in seipin-deficient cells with a relative accumulation of TAG in ER membranes (Cartwright et al., 2015). Similarly, upon LD induction in mammalian cells in the absence of seipin, numerous tiny and growth-abortive LDs emerged from the ER, with a subset of these completely detaching from the ER, a phenomenon not observed in wild-type cells (Salo et al., 2016; Wang et al., 2016). These tiny LDs may reflect spontaneous phase separation or oiling out of excess TAG, as has been postulated to occur in protein-free systems (Deslandes et al., 2017). Thus, seipin could function to attract and foster TAG aggregates in the ER for controlled LD formation.

How could seipin sequester TAG at LD formation sites? The membrane-anchored HHs may be important for this, as they have affinity for a monolayer enclosing NLs (Sui et al., 2018). Furthermore, seipin appears to control TAG condensation in the ER (Santinho et al., 2020) and copurified seipin-promethin complexes contained TAG molecules (Chung et al., 2019). Therefore, seipin may initially attract TAG molecules in the ER bilayer and foster TAG aggregates to grow into a nascent LD. This is in line with the recently reported DGAT1 structure placing the active TAG-generating site within the ER bilayer (Sui et al., 2020; Wang et al., 2020). In the absence of seipin, efficient TAG phase separation in the ER would be delayed, leading to an increase in the ER TAG concentration, as reported (Cartwright et al., 2015; Gao et al., 2017). This increased bilayer TAG concentration eventually leads to the aberrant oiling out of LDs.

The precise role of promethin in this scenario needs to be established, as such oiling out of LDs was not observed in promethin KO cells. Instead, promethin KO cells generated fewer, but larger LDs, which still contained seipin at their ER-LD contact (Chung et al., 2019). This may suggest that seipin is de facto required to keep ER TAG concentration low enough to prevent aberrant LD formation, but promethin may aid seipin in decreasing the barrier for LD formation. Therefore, in the absence of promethin, as LDs are assembled by seipin alone, the flux to these LDs keeps the ER TAG concentration low enough to prevent aberrant LD formation, but not high enough to trigger the assembly of new LDs by seipin. This would then lead to the formation of fewer but larger LDs.

Morphology of Seipin-Mediated ER-LD Contacts

After their formation, LDs remain functionally connected to the ER via ER-LD contacts, which function to facilitate cargo exchange between the two organelles. Morphologically, ER-LD contacts can be broadly categorized into two overlapping classes: membrane bridges, wherein the LD monolayer appears continuous with the ER bilayer; and areas of close ER-LD proximity without direct membrane continuity (Figure 2A and B; Salo and Ikonen, 2019). In addition, proteinaceous tethers between the two organelles have also been observed by EM (Wang et al., 2016; Salo et al., 2019; Figure 2C). Although the functional significance of these different contact zones is not yet understood, seipin appears to be primarily involved in the formation and maintenance of the membrane bridges between ER and LDs.

Figure 2.

Seipin-Mediated ER-LD Contacts. A: Topologies of ER-LD contacts. Left panel: membranous bridges, mediated by seipin, right panel: zones of ER-LD proximity, likely mediated by various ER-LD tethers. B: Examples of ER-LD contacts from primary human fibroblasts. Left panel is adapted from Salo et al. (2019) with permission of the publisher, and right panel is from the same data set. Scale bar: 50 nm. Orange arrowhead indicates neck-like membranous bridge. C and D: Examples of the architecture of seipin mediated ER-LD contacts, adapted from Salo et al. (2019) with permission of the publisher. Scale bars for C: 100 nm, 50 nm, 20 nm, and 20 nm. Scale bars for D: 10 µm, 50 nm, 50 nm and 20 nm. C: Examplary neck-like ER-LD contact from A431 cells with seipin trapped at the NE ER subdomain. Orange arrowhead indicates neck-like membranous bridge, blue arrowheads indicate fibrous tethers. Right panel is 3D modeling of the LD (brown), NE (blue), and NE-LD contact site (red). D: CLEM of A431 cells with seipin tagged endogenously with GFPx7 and expressing the nascent LD marker LiveDrop-mCherry. The inset of the fluorescence microscopy panel (left) depicts the same LD in the same orientation as the EM images. Orange arrowhead indicates neck-like membranous bridge. Right panel depicts 3D modeling of the LD (brown), ER (yellow) and the ER-LD contact site (red). ER = endoplasmic reticulum; LD = lipid droplet; NE = nuclear envelope; CLEM = correlative light and electron microscopy.

Considering the persistent localization of seipin at ER-LD contact sites (Salo et al., 2016; Wang et al., 2016), the morphological features of seipin-mediated ER-LD contacts were recently characterized. Using correlative light and EM, seipin-mediated contacts were found to harbor a strikingly uniform, neck-like architecture (Salo et al., 2019). At these ER-LD necks, the LD monolayer appeared to fuse with the ER lumen, indicating membrane continuity between the outer leaflet of the ER and the LD monolayer (Figure 2C and D). The diameter of the region in touch with the LD was circa 15 nm, a size consistent with the structure of oligomeric seipin resolved with the cryo-EM (Sui et al., 2018; Yan et al., 2018). This suggests that seipin may physically limit the diameter of the ER-LD neck. High curvature of the neck implicates a specific PL composition, and the lipid-binding domain of seipin might be involved in stabilizing the neck. Indeed, such seipin-mediated ER-LD necks were disrupted by acute removal of seipin via auxin-induced degradation (Salo et al., 2019).

A morphologically similar ER-LD contact of circa 15 nm in diameter, with apparent membrane continuity, was also recently observed by cryo-EM tomography of HeLa cells (Figure 3E of Mahamid et al., 2019) and similar neck-like structures were previously observed in, for example, Drosophila and Cos-1 cells (Kassan et al., 2013; Wilfling et al., 2013). Membranous bridges between inner nuclear membrane and intranuclear LDs in yeast cells were also reported to be seipin-dependent (Romanauska and Köhler, 2018). Altogether, these data suggest that seipin localizes to and controls the formation of such bridges. This would be a logical consequence of early LD biogenesis occurring at seipin-marked sites: if the initial LD budding event were to take place through the seipin disk, this would constrain the dimensions of the ER-LD neck accordingly.

Figure 3.

Seipin in Human Adipocytes. A: Confocal images of human A431 WT and seipin KO cells stained with anti-seipin antibodies (Salo et al., 2019). Scale bar: 10 µm. B: Airyscan images of human adipocytes derived from visceral fat stem cells. Adipogenic differentiation was induced by a cocktail of transferrin (10 µg/ml final concentration), human insulin (Actrapid; 66 nM), cortisol (100 nM), triiodothyronine (1 nM), 3-isobutyl-1-methylxanthine (500 µM), and rosiglitazone (2 µM). After 3 days, supplements were reduced to transferrin, insulin, cortisol, and triiodothyronine with the aforementioned concentrations. After 14 days of adipogenic differentiation, cells were fixed and stained with anti-seipin antibodies followed by anti-rabbit Alexa 647 and LipidTox Green. Scale bars: 10 µm and 1 µm. LD = lipid droplet; KO = knockout; WT = wild type.

In nonadipogenic cell lines studied so far, LDs typically harbor a single distinct seipin focus stably associated with each LD (Salo et al., 2016; Wang et al., 2016) and seipin-mediated ER-LD necks appear to be long-lasting structures (stable at least on timescale of hours; Salo et al., 2019). Moreover, seipin recruitment to preexisting LDs has not been reported. These data imply that seipin does not act like a canonical membrane contact site tether but might rather remain at the topologically unique ER-LD contact site via virtue of LD biogenesis occurring through the seipin disk.

There may, however, be species–specific differences, as overexpressed C. elegans seipin was shown to form much larger (several hundreds of nanometers in diameter) peri-LD cages at sites where the tubular elements of the ER are in close proximity to the LD (Cao et al., 2019). Overexpression of C. elegans seipin promoted the formation of such structures also in Cos-7 cells and, conversely, heterologously expressed human seipin localized similarly in tubular peri-LD cages in the C. elegans intestine. Such structures have not been reported upon overexpression of human or Drosophila seipin in nonadipogenic cell lines (Salo et al., 2016; Wang et al., 2016), but seipin peri-LD cages do resemble the localization of overexpressed N-glycosylation defective N88S-seipin mutant, which also accumulates as ring-like structures flanking LDs (Hölttä-Vuori et al., 2013).

An intriguing possibility is that the peri-LD cages of C. elegans intestine, a major fat deposit in these organisms, are a consequence of LD fusion where individual LDs, initially harboring one seipin-mediated neck, fuse and the resulting structure would contain multiple such contact sites in very close proximity. One could envision this to be similar to the case in adipocytes, where a single unilocular LD forms via ripening-mediated coalescence of a high number of smaller ER-derived LDs (Paar et al., 2012; Chu et al., 2014). These giant LDs could thus contain a high number of seipin-mediated contacts. Indeed, our immunofluorescence analysis of endogenous seipin localization in differentiated human adipocyte stem cells suggests that this is the case, with numerous seipin foci juxtaposed to enlarged LDs (Figure 3). Detailed characterization of seipin-mediated ER-LD contacts at the EM level in adipocytes has not been reported.

Modulation of Seipin-Mediated ER-LD Contacts by Other Proteins

Besides seipin, a number of other proteins act at ER-LD contacts (Bohnert, 2020). These include tethering protein complexes, where one partner binds to the ER while the other contacts the LD, such as FATP1 (ER) and DGAT2 (LD; Xu et al., 2012); Rab18 (LD) and the NRZ complex and associated SNARE proteins (ER; Xu et al., 2018); and Orp2 (LD) and VAPA (ER; Weber-Boyvat et al., 2015).Snx14 is also capable of tethering LDs to the ER, by binding to both organelles simultaneously (Datta et al., 2019) and a similar topological arrangement has been proposed for DGAT2 (McFie et al., 2018).DFCP1 has also been documented to facilitate ER-LD contact formation, acting in concert with Rab18 (Gao et al., 2019a; Li et al., 2019a). In yeast cells, Pex30 and the yeast homolog of FIT2 localize to ER-LD contacts, colocalizing with seipin (Choudhary et al., 2018; Joshi et al., 2018; Wang et al., 2018). Newly discovered ER-LD contact site proteins with putative lipid transfer activity include Orp4, VPS13A, and VPS13C (Kumar et al., 2018; Du et al., 2020).

At least in the case of Rab18, Snx14, DFCP1, and VPS13A, overexpression has been shown to increase ER-LD proximity, strongly suggesting that these factors can facilitate tethering of the two organelles (Kumar et al., 2018; Xu et al., 2018; Datta et al., 2019; Li et al., 2019a). The topology of these expanded ER-LD contact zones is probably not similar to seipin-mediated ER-LD necks but rather reflects a close proximity of the two organelles without direct membrane continuity (Figure 2A). However, these factors are still likely to influence/modulate seipin-mediated ER-LD contacts. For example, DFCP1 or Rab18 perturbation decreased ER-LD proximities in seipin depleted Cos-7 cells (Li et al., 2019a). DFCP1 and Rab18 were proposed to form a complex and were shown to co-immunoprecipitate seipin, suggesting that seipin may participate in modulation of DFCP1Rab18-mediated ER-LD contacts. The phenotype of DFCP1 or Rab18 depletion, however, is distinct from that of seipin loss; overall, LD sizes were moderately reduced and LD numbers were increased. However, Rab18 and DFCP1 seem to have cell-type specific functions, as their perturbation can also induce supersized LDs in some cell types such as 3T3-L1 cells, while only minor alterations in LDs are observed in others (Jayson et al., 2018; Xu et al., 2018; Li et al., 2019a). Clearly, further investigations into the interplay between Rab18, DFCP1, and seipin are warranted.

DFCP1 has also been implicated in early LD biogenesis. Overexpressed DFCP1 labeled distinct ER-associated foci that increased in number upon fatty acid administration (Li et al., 2019a). A subset of these developed into LDs, but DFCP1 labeling preceded that of early LD marker peptides HPos or LiveDrop, suggesting that DFCP1 may detect very early NL accumulations. By EM, these DFCP1 foci appeared as small vesicles or ER-associated papillary structures. Whether seipin localizes to these DFCP1 positive regions prior to their maturation into HPos or LiveDrop-positive organelles is an interesting question not yet addressed.

The interplay of Snx14 and seipin at ER-LD contacts was also recently investigated (Datta et al., 2019). LDs were found to be supersized and tiny in Snx14 KO U2OS cells, resembling the LD phenotype upon seipin loss. Overexpressed Snx14 dramatically increased LD-ER proximity and increased TAG generation, suggesting that Snx14, similarly to its proposed yeast counterpart Mdm1, may regulate fatty acid homeostasis (Datta et al., 2019; Hariri et al., 2019). Conversely, loss of Snx14 decreased LD-ER proximity and Snx14 was thus proposed to function as an ER-LD tether, ensuring that the two organelles maintain a connection as ER-produced TAG is fluxed into the maturing LD. However, seipin overexpression failed to rescue the LD phenotype in Snx14 KO cells and vice versa, and concomitant depletion of both proteins did not aggravate the LD phenotypes. The authors therefore concluded Snx14 to function independently of seipin (Datta et al., 2019). On the other hand, considering the different topologies of Snx14 and seipin-mediated ER-LD contacts (Snx14 tethering the organelles in trans and seipin mediating an ER-LD neck with membrane continuity), Snx14 might aid in the stabilization of seipin-mediated ER-LD necks, especially during later stages of LD maturation. Indeed, Snx14 localized to LDs later than seipin, hours after initiation of LD biogenesis (Datta et al., 2019).

Seipin-Mediated Lipid Flux to LDs

The persistent localization of seipin at ER-LD contacts and a defined architecture of seipin-mediated ER-LD contacts suggests that seipin has a role in LDs beyond facilitating initial LD assembly. In support of this, depletion of seipin from mature adipocytes in vivo results in progressive lipodystrophy and decreased adiposity (Liu et al., 2014; Zhou et al., 2015). However, due to the dramatic effects on initial LD biogenesis in the absence of seipin, investigating its role in ER-LD contacts at later stages has been challenging. To overcome this obstacle, the plant-based auxin-inducible degron system (Nishimura et al., 2009; Li et al., 2019c) was recently utilized to rapidly deplete seipin in human cells. With this system seipin could be acutely removed from preexisting ER-LD contacts, revealing a role for seipin in LD maintenance (Salo et al., 2019).

Upon acute seipin removal, LDs rapidly started to become more heterogeneous in size, with larger LDs growing and acquiring more lipid cargo from the ER, and smaller LDs shrinking, apparently donating their cargo to the larger LDs. This occurred in the absence of major alterations to overall net fatty acid handling or lipolysis (Salo et al., 2019). The observed LD size changes can be accounted for by a phenomenon called Ostwald ripening. It is a molecular diffusion process by which smaller droplets continuously leak material to bigger ones through a connecting phase and wherein the directionality is governed by the higher internal pressure of smaller droplets, imposing a flux of molecules toward lower pressure, that is, bigger droplets (Thiam et al., 2013; Thiam and Forêt, 2016).

Ostwald ripening-mediated fusion is a well-known property of liquid-liquid phase-separated systems. A vast number of recent work has begun to elucidate the importance of liquid–liquid phase-separated in organization of the cellular milieu, especially with regard to membraneless organelles (Brangwynne et al., 2009, 2009; Alberti et al., 2019), but also in the context of ER-derived organelle biogenesis such as autophagosome formation (Fujioka et al., 2020).Ripening-mediated growth of LDs via direct LD-LD contacts has also been documented (Gong et al., 2011; Jüngst et al., 2013). At LD-LD interfaces, a larger LD grows via acquisition of lipids from the smaller LD in a process facilitated by Fsp27/CIDE-C (Gong et al., 2011). Interestingly, mutations in Fsp27/CIDE-C also lead to congenital lipodystrophy (Rubio-Cabezas et al., 2009) and CIDE family members are regulators of adipogenesis (Slayton et al., 2019).

However, in contrast to the Fsp27-mediated process, ripening-mediated LD size changes upon acute seipin removal take place via the ER bilayer (Salo et al., 2019). To decipher the role of seipin in this process, heterologous cell fusion experiments were conducted, revealing that in a continuous ER network, LDs with seipin at their ER-LD contact site grew, while LDs with seipin removed from their ER-LD contact shrunk. These data imply that seipin at the ER-LD neck facilitates the growth of the LD it is associated with (Figure 4A and B). This function is critical for preventing the shrinkage of smaller LDs (more prone to ripening) and facilitating their growth (Salo et al., 2019). Recent studies of C. elegans seipin also implicate a droplet-autonomous role of seipin in promoting LD growth (Cao et al., 2019).

Figure 4.

Seipin-Mediated TAG Partitioning. A: Schematic of seipin (green ellipse) function in promoting LD growth, adapted from Salo et al.(2019). B: Seipin removal evokes ripening-mediated LD size changes via the ER. C: The effect of preexisting LDs on nascent LD formation sites: seipin-mediated TAG flux to LDs decreases the nearby TAG concentration in the ER, and thus new LDs tend to be formed at sites distant to preexisting LDs. ER = endoplasmic reticulum; LD = lipid droplet; TAG = triglycerides.

How is seipin performing this LD growth promoting function? Considering the topological arrangement at seipin-mediated ER-LD necks, an intriguing possibility is that seipin controls lipid diffusion, possibly by attracting TAG molecules from within the ER bilayer and facilitating their flux to the LD and/or by controlling PL density at the LD surface (Grippa et al., 2015; Salo et al., 2019). Upon acute seipin removal, ER-LD lipid flux becomes dominated by biophysical forces favoring the growth of larger LDs over smaller ones. In essence, seipin would thus function as a lipid transfer protein. This is well in line with the proposed role of seipin during initial LD assembly.

The ER-LD Network as a Joint System

The notion that LDs may ripen via the connecting lipid phase, the ER bilayer, implies that LDs and ER can be considered as a joint system. This continuity is evident in the case of many proteins, typically harboring hairpin motifs, which exhibit dual localization on ER and LDs (Kory et al., 2016). However, also NL fluxes through the connecting ER bilayer appear to contribute to LD-LD communication. For example, during early LD biogenesis, NLs are likely synthesized at highly dispersed sites along the ER (Poppelreuther et al., 2018). Upon increased local concentrations, a NL lens may nucleate at a site marked by seipin and start to grow. The flux of NL to the growing LD, enhanced by seipin, may thus reduce nearby TAG concentration in the ER, disfavoring the formation of other LDs nearby. Thus, a preexisting LD would inhibit nearby LD formation (Figure 4C). As a consequence, LDs would be expected to form at sites more distant to one another than expected by chance, which is the case at least in human A431 cells (Salo et al., 2019). Indeed, nascent LDs are formed at highly dispersed sites in multiple cell types (Kassan et al., 2013; Chung et al., 2019).

While direct observation of NL fluxes within the ER is technically challenging, rapid advances in imaging and probe technologies (Valm et al., 2017; Adhikari et al., 2019) are likely to facilitate this in the near future. Findings from a recent study, employing elegant imaging techniques (holo-tomographic microscopy), support the notion of a fluid connection of LDs through the ER (Sandoz et al., 2019). It was found that LD growth in a given cell during fatty acid administration was highly synchronous, with both shrinking and growing of LDs happening in concert, as if sharing a common pool of lipid precursors.Furthermore, upon initial LD assembly, preexisting LDs actually initially slightly shrunk in size at the same time as numerous new LDs emerged. This suggests that preexisting LDs unload material into a common lipid pool, the ER (Sandoz et al., 2019).

Concluding Remarks

Overall, these are exciting times for seipin and LD biology. Through concentrated efforts of cell biological, structural, and biophysical endeavors, we are beginning to get clues of the molecular mechanisms underlying LD biogenesis in the ER-LD network. Bridging these emerging concepts to in vivo models of fat storage will likely yield powerful insight and new ways to tackle lipid imbalance beyond that related to the loss of seipin function.

Footnotes

Acknowledgements

We thank Prof. Hannele Yki-Järvinen, Helsinki University Hospital and University of Helsinki, for collaboration in providing human visceral fat biopsies and Dr. Xavier Prasanna for providing the structures for Figure 1. We acknowledge the use of HiLIFE and Biocenter Finland imaging core facilities (Biomedicum Imaging Unit and Electron Microscopy Unit). Figures 1A, 2A and 3C were generated with the help of BioRender.com.

Declaration of Conflicting Interests

The author(s) declared no potential conflicts of interest with respect to the research, authorship, and/or publication of this article.

Funding

The author(s) disclosed receipt of the following financial support for the research, authorship, and/or publication of this article: This work was supported by Sigrid Juséliuksen Säätiö, Jane ja Aatos Erkon Säätiö, Helsingin Yliopisto, and Academy of Finland (grant no. 307415 and 312491).

ORCID iDs

Veijo T. Salo

Maarit Hölttä-Vuori

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Seipin-Mediated Contacts as Gatekeepers of Lipid Flux at the Endoplasmic Reticulum–Lipid Droplet Nexus

Abstract

Keywords

Seipin Structure Gives Clues to Its Function

Seipin Forms an LD Assembly Complex

ER Architecture Promoting LD Formation

Seipin Defines LD Formation Sites

Seipin and Regulation of ER PL Metabolism

Seipin May Sequester TAGs at LD Formation Sites

Morphology of Seipin-Mediated ER-LD Contacts

Modulation of Seipin-Mediated ER-LD Contacts by Other Proteins

Seipin-Mediated Lipid Flux to LDs

The ER-LD Network as a Joint System

Concluding Remarks

Footnotes

Acknowledgements

Declaration of Conflicting Interests

Funding

ORCID iDs

References