Aymeric Chorlay

Luca Monticelli

Joana Veríssimo Ferreira

Kalthoum Ben M'barek

Dalila Ajjaji

Sihui Wang

Errin E Johnson

Rainer Beck

Mohyeddine Omrane

Mathias Beller

Mohyeddine Ormane

Pedro Carvalho

Abdou Rachid Thiam
email: thiam@ens.fr

Membrane Asymmetry Imposes Directionality on Lipid Droplet Emergence from the ER

ou non, émanant des établissements d'enseignement et de recherche français ou étrangers, des laboratoires publics ou privés.

Introduction

The endoplasmic reticulum (ER) is the cellular factory of lipids and membrane proteins, which are subsequently distributed to other organelles [START_REF] Schwarz | The endoplasmic reticulum: structure, function and response to cellular signaling[END_REF]. This anabolic function of the ER is essential for maintaining organelle and cellular homeostasis, particularly when cells enter energy storage regimes. In these periods, excess energy is converted into neutral lipids, which upon synthesis are stored into ER-derived organelles called lipid droplets (LDs), also referred to as oil bodies or lipid bodies for plants [START_REF] Martin | Lipid droplets: a unified view of a dynamic organelle[END_REF]Parton, 2006, Murphy and[START_REF] Murphy | Mechanisms of lipid-body formation[END_REF]. The core of these droplets contains neutral lipids and is covered by a phospholipid monolayer derived from the ER bilayer, likely from its cytoplasmic leaflet [START_REF] Fujimoto | Not just fat: the structure and function of the lipid droplet[END_REF]. It seems therefore essential to replenish the ER with new phospholipids to keep its integrity and proper function during lipogenesis. This is consistent with the trigger of de novo phospholipid synthesis by the activation of sterol regulatory element-binding protein genes in response to lipogenesis [START_REF] Walker | A conserved SREBP-1/phosphatidylcholine feedback circuit regulates lipogenesis in metazoans[END_REF][START_REF] Krahmer | Phosphatidylcholine synthesis for lipid droplet expansion is mediated by localized activation of CTP:phosphocholine cytidylyltransferase[END_REF][START_REF] Aitchison | Nuclear-localized CTP:phosphocholine cytidylyltransferase alpha regulates phosphatidylcholine synthesis required for lipid droplet biogenesis[END_REF]. These concordant events reveal a close relationship between LD biogenesis and ER homeostasis through phospholipid production [START_REF] Lagace | The role of phospholipids in the biological activity and structure of the endoplasmic reticulum[END_REF][START_REF] Aitchison | Nuclear-localized CTP:phosphocholine cytidylyltransferase alpha regulates phosphatidylcholine synthesis required for lipid droplet biogenesis[END_REF][START_REF] Vevea | Role for Lipid Droplet Biogenesis and Microlipophagy in Adaptation to Lipid Imbalance in Yeast[END_REF][START_REF] Ben M'barek | ER Membrane Phospholipids and Surface Tension Control Cellular Lipid Droplet Formation[END_REF]. The consequences on LD biogenesis under conditions of limited phospholipid supply are poorly understood.

LDs also play important roles in several other biological processes, such as viral infections, embryonic development and neurodegeneration [START_REF] Welte | Expanding Roles for Lipid Droplets[END_REF]. These functions are tightly related to how LDs are formed [START_REF] Pol | Review: biogenesis of the multifunctional lipid droplet: lipids, proteins, and sites[END_REF][START_REF] Thiam | The why, when and how of lipid droplet diversity[END_REF][START_REF] Henne | The assembly of lipid droplets and their roles in challenged cells[END_REF]. Neutral lipids synthesized in the ER at low concentration are likely dispersed and mobile in the space between the membrane leaflets. At elevated levels, these lipids can spontaneously demix to nucleate a lipid lens [START_REF] Khandelia | Triglyceride blisters in lipid bilayers: implications for lipid droplet biogenesis and the mobile lipid signal in cancer cell membranes[END_REF][START_REF] Ben M'barek | ER Membrane Phospholipids and Surface Tension Control Cellular Lipid Droplet Formation[END_REF]. The lens grows by accumulating more lipids and eventually forms a spherical LD that buds off or emerges from the bilayer [START_REF] Thiam | The physics of lipid droplet nucleation, growth and budding[END_REF][START_REF] Pol | Review: biogenesis of the multifunctional lipid droplet: lipids, proteins, and sites[END_REF][START_REF] Choudhary | Architecture of Lipid Droplets in Endoplasmic Reticulum Is Determined by Phospholipid Intrinsic Curvature[END_REF]. Throughout these steps toward LD maturation, proteins are dynamically targeted to the LD surface to control the LD function: they target LDs from the cytosol [START_REF] Londos | Perilipin: unique proteins associated with intracellular neutral lipid droplets in adipocytes and steroidogenic cells[END_REF], the ER membrane [START_REF] Wilfling | Arf1/COPI machinery acts directly on lipid droplets and enables their connection to the ER for protein targeting[END_REF], Wilfling et al., 2013[START_REF] Jacquier | Lipid droplets are functionally connected to the endoplasmic reticulum in Saccharomyces cerevisiae[END_REF], or possibly the ER lumen [START_REF] Ohsaki | Biogenesis of cytoplasmic lipid droplets: from the lipid ester globule in the membrane to the visible structure[END_REF]. The amount of LD surface area that is in contact with the lumen and the cytosol will reflect LD proteome and subsequent function. Most findings so far suggest that the surface of the majority of mature LDs is almost entirely exposed to the cytosol.

The emergence of LDs in the cytosol is crucial for metabolism because most regulatory proteins and enzymes that release lipids from LDs reside in the cytosol. LDs interact with nearly all cytoplasmic organelles to trade lipids and proteins [START_REF] Olzmann | Dynamics and functions of lipid droplets[END_REF]. Also, they serve as a reservoir for certain proteins such as histones during fly development [START_REF] Welte | Expanding Roles for Lipid Droplets[END_REF], or aggregationprone proteins that are targeted for degradation [START_REF] Vevea | Role for Lipid Droplet Biogenesis and Microlipophagy in Adaptation to Lipid Imbalance in Yeast[END_REF]. Finally, as most proteins involved in lipid secretion reside in the ER lumen, abnormal accessibility of LDs to ER luminal proteins will likely alter the secretion of very low-density lipoproteins [START_REF] Ohsaki | Biogenesis of cytoplasmic lipid droplets: from the lipid ester globule in the membrane to the visible structure[END_REF].

Factors controlling the directionality of LD budding are still poorly understood [START_REF] Henne | The assembly of lipid droplets and their roles in challenged cells[END_REF]. Seipin is an integral ER protein that forms a scaffold around emerging LDs and is important for proper LD morphology [START_REF] Yan | Human SEIPIN Binds Anionic Phospholipids[END_REF][START_REF] Sui | Cryo-electron microscopy structure of the lipid droplet-formation protein seipin[END_REF]. However, the depletion of Seipin does not hamper the trafficking of proteins residing on the cytosolic face of the ER, such as GPAT4, ADRP or LiveDrop, to the LD surface (Salo et al., 2016;[START_REF] Wang | Seipin is required for converting nascent to mature lipid droplets[END_REF]. This suggests that LDs still bud toward the cytosol in the absence of Seipin, which is therefore not essential to controlling the directionality of LD budding. In contrast, depletion of Fat storage-inducing transmembrane protein 2 (FIT2) results in the budding defects of a LD population [START_REF] Choudhary | A conserved family of proteins facilitates nascent lipid droplet budding from the ER[END_REF]. FIT2 is another integral ER membrane possibly having phosphate phosphatase activity on the ER luminal side [START_REF] Becuwe | FIT2 is a lipid phosphate phosphatase crucial for endoplasmic reticulum homeostasis[END_REF][START_REF] Hayes | Fat storage-inducing transmembrane (FIT or FITM) proteins are related to lipid phosphatase/phosphotransferase enzymes[END_REF]. It would transform phospholipids into diacylglycerol, which are molecules soluble in the neutral lipid phase [START_REF] Thiam | The Energy of COPI for Budding Membranes[END_REF]. FIT2 activity would thus result in depleting phospholipids from the ER luminal leaflet. Thereupon, FIT2 generates at least transiently membrane asymmetries that may determine the directionality of LD budding. Such contribution of FIT2 is appealing as a droplet in a bilayer has equivalent chances to emerge on both sides of the bilayer, unless that a symmetry breaking is introduced (Chorlay and[START_REF] Chorlay | An Asymmetry in Monolayer Tension Regulates Lipid Droplet Budding Direction[END_REF][START_REF] Thiam | The physics of lipid droplet nucleation, growth and budding[END_REF]).

An asymmetry in phospholipid composition of the ER membrane can indeed control the directionality of LD emergence. For example, the recruitment of positively curved lipids, i.e. with bulky hydrophilic head, onto one of the monolayer leaflets will favor LD emergence toward this leaflet [START_REF] Choudhary | Architecture of Lipid Droplets in Endoplasmic Reticulum Is Determined by Phospholipid Intrinsic Curvature[END_REF][START_REF] Chorlay | An Asymmetry in Monolayer Tension Regulates Lipid Droplet Budding Direction[END_REF][START_REF] Zanghellini | Phospholipid demixing and the birth of a lipid droplet[END_REF]. Another source of asymmetry possibly modulating the direction of LD formation is membrane curvature. Curved membranes have a higher pressure in their concave region. Thus, when a LD forms, the internal pressure would tend to push the LD outside the lumen [START_REF] Thiam | The physics of lipid droplet nucleation, growth and budding[END_REF]. The basic tubular structure of the ER membrane could hence modulate the direction of LD budding. Whether membrane curvature determines the directionality of LD emergence is not fully understood. Finally, another possible asymmetry is between the LD monolayer surface tensions [START_REF] Chorlay | An Asymmetry in Monolayer Tension Regulates Lipid Droplet Budding Direction[END_REF], which reflects an asymmetry in the surface coverage between the cytosolic and luminal monolayers surrounding LDs. Soon after nascent LDs appear in the ER (with a diameter over ~ 30 nm), the LD monolayer surface tension should indeed dominate spontaneous curvature and bending effects, and control LD shape. In this size range, we previously worked with macroscopic model LDs and identified that an asymmetry in monolayer surface tension is sufficient to control the emergence of an artificial LD [START_REF] Chorlay | An Asymmetry in Monolayer Tension Regulates Lipid Droplet Budding Direction[END_REF].

Here, we test the hypothesis that monolayer surface tension asymmetry is sufficient to guarantee that newly made and growing LDs become fully exposed to the cytosol. We investigate the presence of imbalances in monolayer surface tension and how they contribute to LD emergence. We find that artificial LDs, herein referred to as aLDs, emerge towards the membrane monolayer side offering the better phospholipid surface coverage, i.e. having the higher phospholipid surface density and hence the lower surface tension. The monolayer coverage is also improved by the insertion of proteins; the strong binding of proteins to the aLD monolayer promotes retention of the droplet into the proteincontaining reservoir. Our data support that systematic LD emergence towards the cytosol requires a phospholipid refill to the cytosolic leaflet of the ER membrane. Such refill ensures keeping a good coverage (low surface tension) of this monolayer. In cells experiencing a limited capacity of phospholipid supply, we observe an aberrant ER morphology concomitant with an increase in the number of LDs in large contact with the ER lumen.

Results

Oil-water surface tension is diminished by covering the interface between the two liquids. In the context of LD biogenesis, the system presents two oil-water interfaces: between the neutral lipid phase and the cytosol, and between the neutral lipids and the ER lumen. An asymmetry in the coverage between these interfaces will yield an asymmetry in surface tension. A monolayer surface coverage is improved by the adsorption of surfactants, which, in cells, are mainly proteins and lipids (Figure 1A). Our model predicts that the recruitment of these membrane constituents to one of the monolayers can improve this monolayer surface coating and be sufficient to promote emergence of the droplet. To test our prediction, we used the droplet-embedded vesicle system (DEV), consisting of giant unilamellar vesicles (GUVs) in which an aLD are incorporated between the leaflets of the bilayer (Figure 1B) (Chorlay andThiam, 2018, Ben M'barek et al., 2017), mimicking the topology of an LD in the ER bilayer. We characterized the degree of emergence of an aLD from the bilayer by comparing the inner and outer surface areas of the droplet (Figure 1C). The asymmetry in the monolayer coverage, and more precisely in surface tension, is reflected by the angles formed by the droplet with the bilayer [START_REF] Chorlay | An Asymmetry in Monolayer Tension Regulates Lipid Droplet Budding Direction[END_REF], which defines the coverage asymmetry parameter (Figure 1C, Star Methods: Determination of shape and position).

Insertion of proteins promotes emergence of lipid droplets from a bilayer

Our model predicts that the insertion of a protein into either monolayer leaflet of an ER-enclosed LD will affect the direction of LD emergence. We tested the effect of different proteins on DEVs made with 1,2-dioleoyl-sn-glycero-3-phosphocholine (here referred to as PC) and containing triolein oil, which is the major cellular neutral lipid.

We started with Arf1, which regulates lipid metabolism and binds to cellular LDs from the cytosol [START_REF] Wilfling | Arf1/COPI machinery acts directly on lipid droplets and enables their connection to the ER for protein targeting[END_REF][START_REF] Guo | Functional genomic screen reveals genes involved in lipid-droplet formation and utilization[END_REF], Thiam et al., 2013a). Arf1 uses an N-terminal myristoyl group and an N-terminal amphipathic helix to bind to membranes. These motifs are shielded within Arf1 when the protein is soluble in the cytosol, and deployed when Arf1 is activated by a guanine nucleotide exchange factor and in the presence of GTP [START_REF] Antonny | N-terminal hydrophobic residues of the G-protein ADP-ribosylation factor-1 insert into membrane phospholipids upon GDP to GTP exchange[END_REF]; the nucleotide exchange is also promoted by reducing Mg 2+ concentration down to µM level by chelating it with EDTA [START_REF] Bigay | Lipid packing sensed by ArfGAP1 couples COPI coat disassembly to membrane bilayer curvature[END_REF]. We can thus dynamically switch on binding and record the impact on aLD emergence. When we added Arf1-Alexa488 and EDTA to DEVs, we observed no protein recruitment, as expected (Figure 1D, Figure S1A). Only when GTP was subsequently added, Arf1 was recruited to the membranes (Figure 1D, Figure S1A). Recruitment of Arf1 was concomitant to the full emergence of the artificial LD to the protein solution (Figure 1D, Figure S1B). Following Arf1 binding, the droplet outer area increased and its inner one decreased (Figure 1D). Consistent with this observation, the outer monolayer coverage was improved (Figure 1H), supporting that binding induced aLD emergence by improving the monolayer coverage.

Most proteins use amphipathic helices (AHs) to associate with the LD surface. The presence of highly hydrophobic residues such as phenylalanine, tyrosine or tryptophan, on the hydrophobic face of AHs increases membrane association. We used an AH peptide from caveolin 1 (Cav aa159-178 with an NBD tag), which binds to LDs and targets other proteins to cellular LDs [START_REF] Ostermeyer | Accumulation of caveolin in the endoplasmic reticulum redirects the protein to lipid storage droplets[END_REF][START_REF] Kassan | Acyl-CoA synthetase 3 promotes lipid droplet biogenesis in ER microdomains[END_REF]. This peptide is particularly rich in hydrophobic amino acids (Figure S1E).

When it was added to DEVs, we observed an increase of the peptide signal on the DEV membrane (Figure 1E). This binding was concomitant with the emergence of the droplet into the protein solution (Figure 1E) and the improvement of the external monolayer coverage.

We next worked with an AH that lacks highly hydrophobic residues. We chose alpha-Synuclein-Alexa488, which is localized to the surface of LDs [START_REF] Cole | Lipid droplet binding and oligomerization properties of the Parkinson's disease protein alpha-synuclein[END_REF] but which binds in vitro only to droplet monolayers with poor phospholipid coating (Thiam et al., 2013a)(Figure S1C,F). Here, alpha-Synuclein failed to bind DEVs and the droplets did not emerge (Figure 1F); the inner and outer droplet areas remained constant and no improvement of the external monolayer coverage was detected (Figure 1F,H). These results support that only AH-containing proteins that are able to strongly associate with monolayers can promote LD emergence.

We finally investigated the effect of perilipins (Plin), which are major LD-associated proteins.

Plins bind to LDs in part by using an 11-mer AH repeat [START_REF] Brasaemle | Thematic review series: adipocyte biology. The perilipin family of structural lipid droplet proteins: stabilization of lipid droplets and control of lipolysis[END_REF]. To test whether this domain alone can mediate LD emergence, we examined the effect of Plin1 11-mer AH repeat, Plin1 93-192 [START_REF] Rowe | Conserved amphipathic helices mediate lipid droplet targeting of perilipins 1-3[END_REF], and a fragment of this domain also displaying an AH motif, Plin1 108-137 (Figure S1E). We found that these peptides did not enable aLD emergence (Figure 1G,J, Figure S1D).

The failure of these peptides in promoting emergence is probably due to inefficient binding to a droplet monolayer which is continuous with a phospholipid bilayer reservoir (see below). Compared to Cav 159-178 AH, the Plin fragments are less dense in hydrophobic residues (Figure S1E), which could explain (at least partly) their inefficient binding [START_REF] Prevost | Mechanism and Determinants of Amphipathic Helix-Containing Protein Targeting to Lipid Droplets[END_REF]. Alternatively, this Plin domain might simply require specific lipids or membrane curvature for binding; other domains of the full-length protein, or protein partners, are required to achieve the strong membrane association of the full-length protein [START_REF] Ajjaji | Dual binding motifs underpin the hierarchical association of perilipins1-3 with lipid droplets[END_REF].

Since many proteins target LDs from the cytosol, we decided to put artificial DEVs in contact with a cytosolic extract from hepatocyte cells. We observed systematic outbound emergence of the droplets into the cytosolic medium (Figure 1G, Figure S1D). This experimental condition closely mimics the cytosolic emergence of cellular LDs mediated by cytosolic components. Note the small decrease of the GUV size because of water evaporation (Figure 1G); this decrease only affects the bilayer tension and does not introduce membrane asymmetry [START_REF] Chorlay | An Asymmetry in Monolayer Tension Regulates Lipid Droplet Budding Direction[END_REF].

To further challenge our hypothesis, we tested the effect of the addition of Apolipoprotein B (ApoB) on LD emergence. ApoB is localized in the ER lumen where it was shown to retain LDs following lipid accumulation [START_REF] Ohsaki | Lipid droplets are arrested in the ER membrane by tight binding of lipidated apolipoprotein B-100[END_REF]. Here, we indeed found that the addition of ApoB promoted aLD emergence (Figure 1G, Figure S1D) towards the ApoB-containing medium. In cells, ApoB would thus promote the emergence of LDs into the ER lumen. As a control, we finally also tested for an effect of bovine serum albumin, which is not an LD protein, and observed no emergence (Figure 1F).

These observations support our hypothesis that the insertion of proteins in a LD monolayer promotes LD emergence by decreasing the monolayer surface tension, through improving its surface coverage. To confirm the link between protein binding and surface tension, we measured for selected examples the changes in surface tension upon addition of the protein using the micropipette aspiration technique (Figure 1G). Upon addition of ApoB (which improved the monolayer coverage and induced aLD emergence) (Figure 1H, Figure S1D), we observed a decrease in the surface tension of the external monolayer (Figure 1I); on the contrary, in the case of Plin1 93-192 (or proteins which did not induce budding), no change in the external tension was measured (Figure 1I). Emergence of aLDs induced by protein binding is therefore mediated by the decrease in monolayer tension (Figure 1J).

In summary, the asymmetric insertion of proteins to an aLD surface promotes emergence toward the protein-enriched monolayer. This emergence results from a decrease of this monolayer surface tension due to an improvement of its surface coverage. In cells, controlling LD emergence would require regulating the access of proteins to the luminal and cytosolic sides of nascent LDs.

Overexpression of proteins that strongly bind to LDs from these compartments will affect LD emergence.

Lipid droplets emerge toward the membrane monolayer side bearing an excess of phospholipids

Monolayer surface coverage can be also improved by increasing the surface density of phospholipids (Figure 1A). Phospholipids can be expected to be more effective than proteins at decreasing tension, by virtue of their higher flexibility and ability to form well-packed interfaces. For example, we measured the minimal surface tension reachable in a monolayer made of PC and found that it is lower than that obtained with ApoB, or with previously studied proteins (Figure S2A). This analysis suggests that the monolayer coverage can be improved more effectively by adding phospholipids than by inserting proteins. The addition of phospholipids could be thus more effective at promoting LD emergence.

We decided to asymmetrically increase the amount of phospholipids in DEVs, for which the outer monolayer is more accessible. However, increasing the number of PC molecules by direct supply is difficult, because PC forms in solution relatively stable and non-fusogenic bilayer structures.

Instead, we chose to test the addition of Lyso-PC because it forms micelles, which spontaneously adsorb onto membranes. To prove this idea, we first made Lyso-PC micelles containing fluorescent NBD-PE (1%, w/w) and added them to PC GUVs (Figure 2A). Over time, after the micelles arrived in the vicinity of the GUVs, the phospholipids were recruited to the membrane, as reported by the increase of the NBD signal (Figure 2A,C, Figure S2B). To know whether the phospholipids were only recruited to the external monolayer, we added dithionite to the bulk solution. Dithionite quenches the NBD signal and can only access the outer leaflet of the GUV, as it does not cross the membrane [START_REF] Laouini | Preparation, characterization and applications of liposomes: state of the art[END_REF]. After few minutes, the NDB signal disappeared completely (Figure 2B,C, Figure S2C), supporting that phospholipids were mainly recruited to the external GUV monolayer. We hence decided to use this technique to increase the amount of phospholipids in the outer monolayer of DEVs.

We took DEVs in which the aLD monolayers had no asymmetry in phospholipid composition.

We next added Lyso-PC micelles to increase the amount of phospholipids in the outer monolayer surface. We systematically observed that the aLDs emerged toward the outside (Figure 2D,E), exactly as with the insertion of proteins. This observation confirmed that aLD emergence happens to the monolayer with the better surface coverage, here mediated by a phospholipid supply from the bulk solution.

It is worth noting that Lyso-PC has a positive spontaneous curvature, which, in principle, can have an effect on LD emergence at nanometric scales, as recently proposed [START_REF] Choudhary | Architecture of Lipid Droplets in Endoplasmic Reticulum Is Determined by Phospholipid Intrinsic Curvature[END_REF].

However, in our macroscopic scale (Figure 2D), spontaneous curvature effects should be dominated by surface tension [START_REF] Thiam | The physics of lipid droplet nucleation, growth and budding[END_REF]. We hypothesize that aLD emergence observed here is not due to Lyso-PC positive curvature, but to its ability to improve the external monolayer surface coverage. To demonstrate that the asymmetry in monolayer phospholipid coverage is the main factor that mediates emergence, we used molecular dynamics simulations. Compared to experiments, in simulations it is much easier increase or decrease the amount of phospholipid selectively in each leaflet. We performed simulations of phospholipid bilayers exclusively made of PC (~75 nm lateral size) and bearing a ~35 nm-sized triolein droplet (Figure 2F); note that PC has a slightly negative spontaneous curvature [START_REF] Zimmerberg | How proteins produce cellular membrane curvature[END_REF], opposite to the pronounced positive curvature of Lyso-PC. Starting from a completely symmetric system (equal number of PC molecules on both leaflets), we performed simulations with an increasing phospholipid imbalance, up to 11 % excess of PC molecules in the upper monolayer (Figure 2G, Figure S2D). We observed that a 4 % imbalance in the phospholipid count is sufficient to induce a significant asymmetry in droplet shape. The asymmetry grows as the lipid imbalance increases, and it causes the aLD to emerge towards the PC-enriched monolayer (Figure 2G,H, Figure S2D). These results support that LDs can emerge from the ER bilayer simply by maintaining an excess of phospholipids on one monolayer side, without using phospholipids with positive spontaneous curvature, which fully agrees with our in vitro data (Figure 2A-E).

It is important to note that generation of high membrane coverage asymmetry cannot be maintained in the bilayer, as it would otherwise destabilize the membrane, both in vitro and in silico.

Instead, the surplus of phospholipids is absorbed by the increasing surface of the emerging droplet.

This process is almost instantaneous as it happens in much less than a second, based on our previous work [START_REF] Chorlay | An Asymmetry in Monolayer Tension Regulates Lipid Droplet Budding Direction[END_REF]. During LD budding in vivo, possible lipid flipping which happens over minutes [START_REF] Pomorski | Lipid flippases and their biological functions[END_REF] would probably not be fast enough to reset the bilayer symmetry (see STAR Methods: Time scale of LD emergence).

Membrane curvature facilitates lipid droplet budding but a supplement of phospholipids remains essential to guarantee full emergence

The ER consists of relatively flat cisternae and high curvature tubules, with diameters in the range of 30-100 nm [START_REF] West | A 3D analysis of yeast ER structure reveals how ER domains are organized by membrane curvature[END_REF]. The DEV system does not fully capture the shape nor the scale of such tubules; also, due to the large size of DEVs, the effect of membrane rigidity on LD shape is supposed to be negligible. Nonetheless, by conserving droplet-to-membrane aspect ratios, our DEV system can be seen as an upscale of a section of a LD forming on a tube (Figure 2I left). We thus decided to use it to explore possible contributions of bilayer membrane curvature on LD emergence.

We made DEVs presenting various droplet/GUV aspect ratios, by varying the aLD and GUV sizes. For each case, we measured the fraction of the droplet area exposed to the exterior (A ext ). This surface fraction was represented against the ratio of the droplet-to-GUV size (Figure 2I), which merely represents the curvature of the GUV bilayer membrane relatively to the droplet. Droplets which were small relative to the GUV size were equally exposed to both membrane sides, A ext = A int = 50% (Figure 2I, lower black circles). In contrast, larger droplets had an external area larger than the inner one, A ext > 50%, indicating that they emerged more towards the exterior (Figure 2I, upper black circles). These results suggest that membrane curvature favors the outward emergence of large droplets. However, we never observed full emergence of large droplets in the DEVs, i.e. A ext = 100%, unless phospholipids or proteins were added to the external leaflet (Figure 2I, red circles). This suggests that curvature promotes outward budding but the addition of phospholipids to the external leaflet is necessary to guarantee full emergence.

To further understand our data, we used a basic theoretical model computing the balance of forces acting on a droplet in a bilayer [START_REF] Chorlay | An Asymmetry in Monolayer Tension Regulates Lipid Droplet Budding Direction[END_REF])(Star Methods : Determination of shape and position). This model well fitted with our experimental data (Figure 2I, black line) confirmed that the effect of phospholipid surface density is stronger than the effect of curvature in determining the direction of emergence (Figure 2J,K, Figure S2E,F). It is worth noting that, if no phospholipid is added in the system, the externalization of growing droplets would induce stretching of the external monolayer. This monolayer, not supplemented with phospholipids, would reach a lower coverage (higher surface tension) than the inner one, which, in turn, would promote inward budding (Figure S2G). In this situation, the droplets will tend to remain in contact with the lumen despite the effect of curvature.

In conclusion, our data support that when a LD forms and grows, the ER membrane curvature helps to preset the direction of emergence of the LD, i.e. in the cytosol. However, the supplement of phospholipids, and eventually of proteins, to the cytosolic monolayer would be required and be more efficient at promoting the full emergence of the LD into the cytosol. In case of phospholipid deficiency on this leaflet, droplets would fail to properly emerge and remain in contact with the ER lumen (Figure S2G).

Altogether, our results in Figures 1 and2 suggest that in close systems, i.e. where there is no excess or addition of phospholipids or proteins, the droplet emergence essentially follows leaflet asymmetry. Emergence occurs toward the monolayer with the better surface coverage, which is improved by the insertion of proteins and phospholipids. In open systems, such as in cells, the order of events is not known, so asymmetry in leaflet coverage may be induced by multiple events. In any case, our in vitro observations predict that LD exclusive formation into the cytosol requires a regulation of phospholipid density and protein binding in of each monolayer.

Stimulated LD production in Saccharomyces cerevisiae cells by oleic acid is associated with aberrant ER morphology

Under basal conditions, the ER cytosolic side probably has enough proteins and phospholipids to impose directionality on LD budding. However, during high lipogenesis regimes, the demands for LD surface coating may become limiting. The emergence of many LDs will deplete phospholipids from the cytosolic leaflet of the ER membrane. In this case, if phospholipids are not present in large excess, quickly replenished or replaced by other surfactant molecules (e.g. proteins), LD emergence (Figure S2G) and ER morphology could be both affected.

We considered cellular model systems in which the ER membrane surface can be easily visualized and characterized. We chose to work with the Saccharomyces cerevisiae cells because most of their ER surface is essentially composed of two domains, the cortical and nuclear ER [START_REF] West | A 3D analysis of yeast ER structure reveals how ER domains are organized by membrane curvature[END_REF]. The total apparent surface of these two ER domains can be approximately determined by considering them as spherical surfaces, which is an overestimation of the ER surface based on quantifications from 3D ultrastructure imaging [START_REF] Wei | High-resolution three-dimensional reconstruction of a whole yeast cell using focused-ion beam scanning electron microscopy[END_REF]. We reasoned that if too many LDs were to form in these cells, they would mobilize phospholipids from the ER cytosolic leaflet and the ER membrane might find a way to compensate for this asymmetric usage of phospholipids.

Yeast strains expressing the ER marker protein Sec63 at endogenous levels, tagged with the monomeric fluorescent protein neon green (Sec63-mNG), were grown in oleate-containing media (0.1%) for 6 to 24hr, in order to produce numerous and big enough LDs to quantify by fluorescence (LDs were stained by monodansylpentane, MDH). Under these conditions we observed increased numbers of cells with aberrant ER membranes, as compared to non-oleate loaded cells (Figure 3A,B).

Unusual ER membrane signals appeared in the cytoplasm in the form of lumps or threads. The cortical ER, which seemed to be discontinuous or disappeared, was more affected than the nuclear ER, which was sometimes shrunk or distorted (Figure 3B, Figure S3F). These phenotypes were observed irrespective whether LD production was stimulated during exponential or stationary phases (Figure S3A-D). Interestingly, cells with aberrant ER very often contained larger numbers of LDs (Figure 3C, Figure S3E), suggestive of a link between the LD number/surface and ER morphology defects. By estimating the amount of total LD surface over the cortical and nuclear ER surface, we found that this ratio was less than 2% in most cells cultured in regular media (no oleate) (Figure 3D)consistent with values from tomogram reconstitution data of Saccharomyces cerevisiae cells [START_REF] Wei | High-resolution three-dimensional reconstruction of a whole yeast cell using focused-ion beam scanning electron microscopy[END_REF][START_REF] West | A 3D analysis of yeast ER structure reveals how ER domains are organized by membrane curvature[END_REF]. In contrast, in cells grown in oleate-containing media, this ratio was more than 22 % (Figure 3D).

Assuming that the LDs were all formed toward the cytosol in the oleate loaded cells, the ER membrane would be depleted of 22 % phospholipids on its cytosolic leaflet, which would generate a huge asymmetry on the membrane. Indeed, a pure bilayer can only withstand an asymmetry of less than 1 % in phospholipids (Segrest et al., 2015, Farge and[START_REF] Farge | Shape changes of giant liposomes induced by an asymmetric transmembrane distribution of phospholipids[END_REF]. Otherwise, the membrane will undergo spontaneous reshaping to release the stress created by this asymmetry (Segrest et al., 2015, Farge and[START_REF] Farge | Shape changes of giant liposomes induced by an asymmetric transmembrane distribution of phospholipids[END_REF]. With the potential membrane asymmetry of 22 % induced by LD formation, such spontaneous membrane reshaping mechanism could explain, at least in part, the observed loss of the normal ER morphology.

Phospholipids may be refilled to the ER cytosolic leaflet during the formation of LDs in non-fed cells, through the activity of scramblases [START_REF] Pomorski | Lipid somersaults: Uncovering the mechanisms of protein-mediated lipid flipping[END_REF]. In contrast, the fed cells were probably challenged beyond their capacity of phospholipid supply for covering the LDs. Thus, they would likely respond to this demand by remodeling their ER membrane. Consistent with this analysis, cells fed under our working conditions were previously reported to reach a ~1.3 fold increase of their total phospholipid [START_REF] Grillitsch | Lipid particles/droplets of the yeast Saccharomyces cerevisiae revisited: lipidome meets proteome[END_REF]. Our quantifications from thin layer chromatography showed that the total phospholipid level barely changed (Figure S3G). Overall, these observations suggest that LD formation in oleate feeding conditions, the phospholipid levels were not sufficient to overcome the demand.

We reasoned that increasing phospholipid production under lipogenic conditions would rescue the aberrant ER phenotype. In yeast, inositol is a major component regulating phospholipid biosynthesis (Carman and Henry, 1989). It is a precursor of the synthesis of PI, a main lipid of yeast ER membranes. The enzymes executing PI synthesis are located on the cytosolic membrane side of the ER (Carman and Han, 2011). The addition of inositol to yeast cells led to a five-fold increase in PI as well as overall cellular phospholipid levels [START_REF] Gaspar | Inositol induces a profound alteration in the pattern and rate of synthesis and turnover of membrane lipids in Saccharomyces cerevisiae[END_REF], (Figure 3SH); only PC levels were slightly decreased, as reported before [START_REF] Gaspar | Inositol induces a profound alteration in the pattern and rate of synthesis and turnover of membrane lipids in Saccharomyces cerevisiae[END_REF] (Figure 3SH). We cultured yeast cells in the presence of oleate and inositol, and quantified the amount of total phospholipid per microgram proteins. The overall phospholipid level was increased by ~ 30 % (Figure 3F) and we found a significant increase of PI (Figure 3SH). Interestingly, we found that the addition of inositol rescued the aberrant ER phenotype (Figure 3E,G). In this condition, we also noticed that cells presenting a normal ER morphology tolerated a higher number of LDs than the control cells, i.e. without inositol addition but in the presence of OA (Figure 3H). Based on our model, these observations support that the cells treated with inositol, which induced an increase in total phospholipids, were better set with phospholipids to cover the number/surface of generated LDs. This overall rescue of the aberrant ER phenotype is probably mediated by the presence on the ER cytosolic monolayer of additional phospholipids, probably PI (Figure S3H), which contributed to the coverage of the LDs.

Finally, we tried to accentuate the aberrant ER phenotype, by aiming at decreasing the amount of phospholipids available in the cell. We deleted the choline-phosphate cytidylyltransferase gene (PCT1), the rate-limiting enzyme in the Kennedy pathway of PC biosynthesis [START_REF] Deslandes | Turnover of phosphatidylcholine in Saccharomyces cerevisiae. The role of the CDP-choline pathway[END_REF]. However, no striking differences were observed between WT and pct1∆, except at 6hr after OA treatment where pct1∆ cells showed a stronger increase of aberrant ER frequency (Figure S3D). This is probably because the large asymmetric phospholipid usage introduced by the numerous LDs formed was already too strong and therefore no further ER membrane reorganization could be induced. As an alternative, the contribution of PCT1 to PC synthesis under our working condition could be smaller as compared to pathways involving e.g. Cho2 and Opi3.

In summary, the formation of multiple LDs correlated with the appearance of an aberrant ER membrane. This phenotype was rescued by the addition of inositol, which led to an increase in phospholipid levels. These results support that increasing phospholipid levels during lipogenesis ensures the formation of multiple LDs in the cytosol and keeping the ER morphology intact.

Proper LD emergence is facilitated by phospholipid synthesis in yeast cells

The formation of multiple LDs depletes phospholipids from the cytosolic leaflet of the ER membrane. If the ER cannot meet this phospholipid demand on the cytosolic monolayer its luminal monolayer could become better covered with phospholipids, at least transiently. Therefore, we expect that in such conditions some LDs fail to fully emerge in the cytosol, and, instead, remain in contact with the ER lumen.

In the yeast cells fed with oleate to stimulate LD production, we observed increased numbers of LDs sharing large contacts with the ER membrane marker (Figure 4A, left, Figure 4B). This phenotype was much more pronounced in cells with aberrant ER. In these cells, the ER sometimes adopted a cup shape structure, which embedded the LD, or surrounded the LD on the observed plane (Figure 4A,arrows). This phenotype was more pronounced in pct1∆ cells (Figure S4A). Note that theses LDs were not in the nucleus, which remained intact in most cases (Figure 4A, Figure S4A,B).

These data support a failure in LD emergence in the cytosol, which would be consistent with our model of the need of phospholipid refill on the cytosolic monolayer for proper LD emergence (Figure 2). In fact, by the addition of inositol, which increased phospholipid levels and reduced the aberrant ER phenotype (Figure 3E,F,G), we found that the fraction of LDs which were in large contact with the ER membrane was significantly decreased (Figure 4A, right, Figure 4B).

To visualize the topology of the ER-associated LDs, we performed electron microscopy (EM)

imaging of high pressure frozen (HPF) cells. In control (non-oleate loaded) cells, most LDs were not in large contact with the ER (Figure 4C,top). Even though the ER membrane was sometimes close to LDs in these cells, there was a perfectly distinguishable free space between the two organelles (Figure 4C top , black arrowhead). In oleate-loaded cells, which developed an aberrant ER phenotype, LDs were larger than in control cells and were often molded by ER membranes (Figure 4C). The ER lumen, marked by a membrane region devoid of ribosomes, seemed to be in contact with the LDs. In some cases, the focal section enabled us to catch structures that looked like an unzipping of the ER bilayer at the LD edge (see Figure 4C, blue arrowhead). These phenotypes might look like mere membrane wrapping phenomena, but could be rather signature of a defect in LD emergence, as theoretically predicted (Figure 2SG). This conclusion is consistent with previous studies, performed exactly under our working conditions, which showed that the LDs were accessible to proteins expressed in the ER lumen, by both fluorescence and EM imaging [START_REF] Mishra | Mature lipid droplets are accessible to ER luminal proteins[END_REF]. To better visualize the membrane, we opted for a chemical fixation of the cells instead of HPF. With this method, membranes were better labeled and darker (Figure 4D white arrowhead, Figure S4C). This approach enabled us to identify in oleate-loaded cells section planes where LDs were in contact with the ER lumen, as previously seen in yeast cells lacking FIT2 [START_REF] Choudhary | Architecture of Lipid Droplets in Endoplasmic Reticulum Is Determined by Phospholipid Intrinsic Curvature[END_REF][START_REF] Choudhary | A conserved family of proteins facilitates nascent lipid droplet budding from the ER[END_REF]. On other focal sections, we observed also phenotypes where the ER membrane formed stacks on a portion of the LDs (Figure 4E, Figure S4D), and sometimes we could catch what seemed to be an unzipping of the membrane at the LD edge (Figure 4E, black arrowhead).

LDs remain more in contact with the ER in Drosophila cells with reduced CCT1 activity

We decided to test our model in a different cell type by manipulating phospholipid levels. We worked with Drosophila Kc167 cells because the synthesis of PC in these cells essentially happens through the Kennedy pathway. Contrary to PCT1 inhibition in yeast, PC level in Drosophila cells is markedly reduced by inhibiting the rate-limiting enzyme in PC synthesis, the choline-phosphate cytidylyltransferase enzyme 1 (CCT1) [START_REF] Krahmer | Phosphatidylcholine synthesis for lipid droplet expansion is mediated by localized activation of CTP:phosphocholine cytidylyltransferase[END_REF][START_REF] Aitchison | Nuclear-localized CTP:phosphocholine cytidylyltransferase alpha regulates phosphatidylcholine synthesis required for lipid droplet biogenesis[END_REF]. Cells expressing the GFP-tagged ER marker Sec61β were treated with dsRNA targeting the CCT1 encoding transcript (Figure S5A), and then they were fed with oleate. We observed an increase in the number of large LDs (Figure S5C,D), a signature of PC deficiency in CCT1 depletion [START_REF] Krahmer | Phosphatidylcholine synthesis for lipid droplet expansion is mediated by localized activation of CTP:phosphocholine cytidylyltransferase[END_REF][START_REF] Aitchison | Nuclear-localized CTP:phosphocholine cytidylyltransferase alpha regulates phosphatidylcholine synthesis required for lipid droplet biogenesis[END_REF]. As compared to yeast cells, the ER formed in the fly cells a complex 3D network, which was a limitation to properly characterize ER membrane morphology and to quantify the LD-to-ER surface ratio. Nonetheless, by using super-resolution structured illumination microscopy, we found in was reminiscent of what a LD-associated protein signal would look like. This observation revealed that these LDs were in close contact with the ER lumen. We next performed EM imaging and found that CCT1 depletion led to significantly more ER-connected LDs (Figure 5B). We also found another significant phenotype where the LDs had regions devoid of ribosomes and delineated by membranes, which were probably the ER. This phenotype is reminiscent of the one observed in yeast (Figure 4C),

where LDs are probably in contact with the ER lumen.

Model LDs equally emerge on both membrane sides if phospholipids are not replenished

To further understand the ER-associated LD phenotypes, we opted for the DEV set-up to mimic the absence of a phospholipid refilling mechanism to the cytosolic monolayer, under conditions where many LDs are generated. We made PC DEVs with many small and symmetrically positioned artificial LDs (Figure 6A). Under these conditions, the aLD-to-bilayer surface ratio was ~ 2% (Figure 6A), close to LD/ER surface ratios in yeast (Figure 3D). We then decreased the DEV tension, by deflating them with a hypertonic buffer added to the aqueous medium, to promote spontaneous emergence of the droplets (Deslandes et al., 2017, Chorlay and[START_REF] Chorlay | An Asymmetry in Monolayer Tension Regulates Lipid Droplet Budding Direction[END_REF]. By following droplets of the same DEV, we observed that they fully emerged and, interestingly, into both sides of the bilayer (Figure 6B,C; Figure S6A,B). These results were highly reproducible and the budding events into the internal and external sides were statistically equal (Figure 6D).

This even distribution of emerged artificial LDs across the bilayer probably arose from a compensatory mechanism of surface expansion (Figure S6C). The external and internal monolayers of the bilayer had initially approximately the same phospholipid coverage. When an artificial LD emerged, e.g. towards the exterior, it expanded the external leaflet surface area, while the amount of phospholipids remained constant on the time scale of droplet budding (because flip-flop rates are extremely slow for PC [START_REF] Tieleman | Lipids out of equilibrium: energetics of desorption and pore mediated flip-flop[END_REF]Marrink, 2006, Nakano et al., 2009)). Expansion of the external leaflet reduced this monolayer phospholipid coverage as compared to the internal monolayer; therefore emergence of the second droplet was favored toward the opposite leaflet (Figure S6C).

Emergence of the second droplet re-equilibrated the surface coverage between the two monolayers, and so on and so forth. This feedback mechanism can explain the increased number of LDs that stayed in contact with the ER lumen (Figure 4,5): in ER membranes unable to provide enough phospholipids to the cytoplasmic monolayer, the emergence of an LD population into the cytosol will tend expose of the rest of the LDs to the lumen.

To fully prove this emergence feedback mechanism, we made a PC DEV with one large artificial LD (Figure 6E), with a droplet-to-bilayer surface of 22% (Figure 6F), close to the situation where many LDs are formed in yeast cells (Figure 3D). The aLD was symmetrically positioned in the bilayer (Figure 6E,G). With a micropipette, we suctioned out a small portion of the droplet and removed it, which mimics emergence from the ER of a given total aLD volume or surface (Figure 6E, Figure S6D). This operation removed phospholipids from the outer monolayer, reducing its coverage as compared to the inner one. As expected, the remaining total aLD volume becomes more exposed toward the side with the better phospholipid coverage, i.e. the luminal side (Figure 6E,G). Emergence to the luminal side was evident after slightly decreasing the bilayer tension or internal pressure (previously increased by the ablation step) (Figure 6E,G,H); note that simply decreasing this tension or pressure without the induced phospholipid coverage asymmetry was not solely sufficient to induce such emergence (Figure S6H). A cross section of a 3D visualization of the final system demonstrating the internal emergence is shown in Figure 6H and Figure S6F. In conclusion, this experiment is an evidence that, in the absence of phospholipid replenishment, the budding of a droplet population toward on membrane side will promote the emergence of the rest of the droplets to the opposite side.

Finally, we estimated that the difference in monolayer coverage that induced the aLD internal emergence was about 9 %, corresponding to an imbalance of monolayer surface tension of less than 8 % (Figure 6I, Figure S6G). Because our experimental condition reproduced LD/ER surface ratios found in yeast cells boosted with LDs, our findings suggest that maintaining an excess of around 9 % of phospholipids on the cytosolic monolayers of the ER is sufficient to guarantee proper LD formation in the cytosol during high lipogenesis regimes.

Discussion

In the present work, we show that LDs bud off and emerge towards the membrane side with higher monolayer leaflet coverage. LDs rapidly absorb this asymmetry by emerging on the side of higher coverage. Because in vivo LDs mainly emerge to the cytosol, our results suggest that, during lipogenesis, the cytosolic monolayer coverage should be transiently kept higher than the luminal one.

We shed light on the mechanisms that maintain such tension imbalance.

The strong adsorption of proteins from the cytosol improves the coverage of the monolayer surface, by masking possible phospholipid packing defects of the monolayer [START_REF] Bacle | Interdigitation between Triglycerides and Lipids Modulates Surface Properties of Lipid Droplets[END_REF][START_REF] Prevost | Mechanism and Determinants of Amphipathic Helix-Containing Protein Targeting to Lipid Droplets[END_REF], Thiam et al., 2013b). Thereupon adsorbed proteins lower the surface tension of the cytosolic monolayer and imposes LD emergence in the cytosol (Figure 7A). This directional emergence could be also mediated by ER-embedded proteins that target LDs. Most of these proteins are indeed strongly attached to the LD cytosolic monolayer by using AHs and/or hairpin motifs [START_REF] Kassan | Acyl-CoA synthetase 3 promotes lipid droplet biogenesis in ER microdomains[END_REF][START_REF] Pol | Review: biogenesis of the multifunctional lipid droplet: lipids, proteins, and sites[END_REF], Wilfling et al., 2013[START_REF] Jacquier | Expression of oleosin and perilipins in yeast promotes formation of lipid droplets from the endoplasmic reticulum[END_REF][START_REF] Huang | Unique Motifs and Length of Hairpin in Oleosin Target the Cytosolic Side of Endoplasmic Reticulum and Budding Lipid Droplet[END_REF].

More generally, the strong binding of any protein to the monolayer of a nascent LD will tend to retain the LD into this monolayer compartment (Figure 7B). Thus, proteins that would strongly bind LDs from the cytosol and the ER lumen would compete for LD emergence. Depending on the binding affinity of these proteins, a LD could be trapped in the ER membrane with various degrees of contact with the ER lumen and the cytosol (Figure 7B). This probably explains how in hepatocytes ApoB100 retains some mature LDs in the ER lumen, due to its irreversible binding to the LD surface (Figure S1F). In these cells, ApoB binds only to some LDs to possibly make them accessible to ER luminal proteins, which is important for the secretion of very low-density lipoprotein [START_REF] Ohsaki | Lipid droplets are arrested in the ER membrane by tight binding of lipidated apolipoprotein B-100[END_REF][START_REF] Ohsaki | Biogenesis of cytoplasmic lipid droplets: from the lipid ester globule in the membrane to the visible structure[END_REF]. Similarly, LD retention in the ER lumen is also likely caused by the overexpression of Perilipin constructs targeted to the ER lumen. Indeed, in such cases LDs become accessible to ER luminal proteins [START_REF] Mishra | Mature lipid droplets are accessible to ER luminal proteins[END_REF]. Thus, the overexpression or down-regulation of any protein that is able to tightly bind to the LD surface will affect LD emergence (Figure 7B). For LDs to fully emerge into the cytosol, proteins with high affinity for LD surface should not get access to LD from the ER lumen. One possible way to fence out proteins, especially those coming from the luminal side, is provided by some specific proteins localized to LD biogenesis sites such as Seipin [START_REF] Wang | Seipin is required for converting nascent to mature lipid droplets[END_REF][START_REF] Chen | The Collaborative Work of Droplet Assembly[END_REF][START_REF] Thiam | The why, when and how of lipid droplet diversity[END_REF][START_REF] Grippa | The seipin complex Fld1/Ldb16 stabilizes ER-lipid droplet contact sites[END_REF].

The release of multiple ER-derived LDs into the cytosol will reduce the amount of phospholipids in the ER cytosolic leaflet. Hence, this leaflet needs to be refilled to guarantee that all LDs bud in the same direction while preserving ER morphology (Figure 7C). Otherwise, chances are that LDs get in large contact with in the ER lumen (Figure 7D). In Saccharomyces cerevisiae and Drosophila cells deficient in phospholipids or PCT1/ CCT1 enzymes, many LDs indeed failed to fully emerge in the cytosol.

Excess phospholipids on the cytosolic monolayer will make the ER asymmetric, at least under lipogenesis conditions during which phospholipid synthesis is triggered on the cytosolic ER leaflet [START_REF] Lagace | The role of phospholipids in the biological activity and structure of the endoplasmic reticulum[END_REF][START_REF] Walker | A conserved SREBP-1/phosphatidylcholine feedback circuit regulates lipogenesis in metazoans[END_REF][START_REF] Van Meer | Membrane lipids: where they are and how they behave[END_REF]. Thus, the idea that the ER has a symmetric membrane needs to be re-considered. During lipogenesis conditions, the ER may actually display regions where an asymmetry in phospholipid type and density is generated for controlling LD emergence side. For example, early lipidomic studies of ER-derived microsomes from rat liver have shown that the ER outer leaflet is more enriched in PC and less in phosphatidylethanolamine [START_REF] Higgins | Asymmetry of the phospholipid bilayer of rat liver endoplasmic reticulum[END_REF]. Also, phospholipids are provided in an asymmetric fashion by the fusion with the ER of retrograde transport vesicles. These vesicles are derived from the Golgi, that is asymmetrical in lipid composition [START_REF] Van Meer | Membrane lipids: where they are and how they behave[END_REF], and have more and different phospholipids on their outer monolayer. Finally, the exclusive phosphate phosphatase activity of FIT2 in the ER luminal membrane side suggests at least a transient asymmetry of the membrane in phosphatidic acid.

Regarding the asymmetry in phospholipid density, various mechanisms can enable the refill of the cytosolic leaflet with phospholipids. De novo phospholipid biosynthesis is probably the primary mechanism fulfilling such need [START_REF] Lagace | The role of phospholipids in the biological activity and structure of the endoplasmic reticulum[END_REF]Ridgway, 2013, van Meer et al., 2008). Enzymes of the Kennedy pathway for de novo PC biosynthesis are indeed important for LD emergence [START_REF] Lagace | The role of phospholipids in the biological activity and structure of the endoplasmic reticulum[END_REF][START_REF] Krahmer | Phosphatidylcholine synthesis for lipid droplet expansion is mediated by localized activation of CTP:phosphocholine cytidylyltransferase[END_REF][START_REF] Aitchison | Nuclear-localized CTP:phosphocholine cytidylyltransferase alpha regulates phosphatidylcholine synthesis required for lipid droplet biogenesis[END_REF] (Figure S4A, Figure 5). Alternatively, ERembedded flippases [START_REF] Bishop | Assembly of the endoplasmic reticulum phospholipid bilayer: the phosphatidylcholine transporter[END_REF] might transfer phospholipids from the ER luminal to the cytosolic monolayer to control LD emergence. The unselective insertion of proteins into the ER cytosolic monolayer may also increase the phospholipid density in this monolayer [START_REF] Segrest | Surface Density-Induced Pleating of a Lipid Monolayer Drives Nascent High-Density Lipoprotein Assembly[END_REF]. Finally, FIT2 is also a potential factor promoting ER membrane asymmetry in phospholipid density. By its presumable phosphate phosphatase activity in the ER luminal monolayer [START_REF] Becuwe | FIT2 is a lipid phosphate phosphatase crucial for endoplasmic reticulum homeostasis[END_REF][START_REF] Hayes | Fat storage-inducing transmembrane (FIT or FITM) proteins are related to lipid phosphatase/phosphotransferase enzymes[END_REF], FIT2 may convert phospholipids of this monolayer into diacylglycerols.

These molecules are soluble in the neutral lipid phases and will be released from the luminal monolayer into the lipid phase; they could be next reconverted into phospholipids on the cytosolic monolayer. Hence, FIT2 activity would result in a net phospholipid transfer from the ER luminal leaflet to the cytosolic one. Such action would maintain LD emergence towards the cytosol.

Finally, membrane curvature may also help LD emergence (Figure 2I-K). However, the emergence of LDs in the cytosol will deplete the cytosolic monolayer of phospholipids, unless these are refilled. This would introduce an asymmetry in monolayer phospholipid coverage that would favor inward emergence. These antagonist effects would perturb the tubular membrane topology and later formed LDs would remain in contact with the ER lumen. This situation will induce phenotypes where growing LDs, which are more exposed to the lumen, become tightly apposed to the ER membrane. Hence, the often-reported ER-wrapped phenotype may be a signature of a defect in LD emergence.

In conclusion, our findings bring insights on mechanisms of cytosolic LD biogenesis, and their link with ER morphology and phospholipid biosynthesis. Our data support that the ER membrane undergoes dynamic asymmetrical remodeling during lipogenesis periods, so as to guarantee LD formation in the cytosol, which is critical for LD proteome and subsequent biological functions.

Figures Captions Figure 1: The insertion of proteins promotes artificial LD emergence

A) The surface tension of an oil/water interface is decreased by improving the surface coverage with surfactants, such as proteins and phospholipids. J) Summary of the effect of the protein solutions on aLD emergence and membrane coverage asymmetry (decrease of external surface tension).

See also supplementary Figure S1. See also supplementary Figure S2. See also supplementary Figure S3. See also supplementary Figure S4. LDs surface over the GUV bilayer apparent area (See Figure S6B). Scale bar is 10 µm. Error bar is the measurement precision.

B) Example of two LDs in the same DEV that emerge on different directions upon decreasing the DEV bilayer tension. Scale Bar is 2.5 µm. An extremely deflated GUV with aLDs emerged on both bilayer side is shown in Figure S6A).

C) The external area variation ("Aext var") is shown for the two aLDs that emerged on different direction in (B). Error bar is the measurement precision.

D) Example and quantification of the number of aLDs that emerged in the interior and exterior of DEVs. Scale bar is 10 µm. T-test showed ns difference; N = 100. Data are represented as mean ±SD.

E) Mimic of the budding of a total lipid droplet number/surface from a bilayer. The aLD of a DEV is suctioned on the exterior and ablated (see Figure S6D) to mimic emergence of a certain LD volume/surface from the bilayer. The remaining artificial LD is displaced in the bilayer lumen. Deflating the DEV to reduce its surface tension led to almost complete exposing the remained artificial LD into the lumen (full data see Figure S6E). Scale bar is 10 µm.

F) Quantification of the total aLD surface over the GUV bilayer initial apparent area. Error bar is the measurement precision.

G) The aLD external area variation corresponding to E. Error bar is the measurement precision.

H) 3D reconstitution of the inward emerged aLD (see Figure S6F). Scale bar is 10 µm.

I) Quantification, based on the ablated aLD surface, of the relative change in the phospholipid number between the internal and external monolayers. Figure S6G shows the related external/internal monolayer tension generated. Error bar is the measurement precision.

See also supplementary Figure S6. All GUVs were 99.5 % DOPC 0.5% (w/w) Rhodamine-DOPE. GUVs were prepared by electroformation (Thiam et al., 2013a). Phospholipids and mixtures thereof in chloroform at 0.5 µM were dried on an indium tin oxide (ITO) coated glass plate. The lipid film was desiccated for 1 h. The chamber was sealed with another ITO coated glass plate. The lipids were then rehydrated with a sucrose solution (275+-15 mOsm). Electro-formation is performed using 100 Hz AC voltage at 1.0 to 1.4 Vpp and maintained for at least 1 h. This low voltage was used to avoid hydrolysis of water and dissolution of the titanium ions on the glass plate. GUVs were either stored in the chamber at 4 °C overnight or directly collected with a Pasteur pipette.

STAR★METHODS KEY RESOURCES

To prepare the artificial lipid droplets (aLDs), 5 µL of the oil was added to 45 µL of HKM buffer. The mixture was sonicated. The diameter of the resulting droplets is on the order of a few hundred nanometers. To make DEV, GUVs were then incubated with the LDs for 5 min. The GUV-LD mixture was then placed on a glass coverslip pretreated with 10 % (w/w) BSA and washed three times with buffer.

Determination of the degree of LD emergence in the DEV experimental system

We characterized the degree of emergence of the aLD from the bilayer by comparing the inner and outer surface areas of the droplets (Figure 1C). This was assessed by considering the inner and outer surfaces of the LD as spherical caps. All micrographs of in vitro systems were made on a Carl ZEISS LSM 800. Glass coverslips: Menzel Glaser (24x36mm #0).

Determination of the coverage asymmetry.

The asymmetry in the monolayer coverage, is directly related to the surface tension asymmetry which is reflected by the angles formed by the droplet with the bilayer (α and θ, see Figure 1C). More precisely we defined "coverage asymmetry" parameter as (α -θ)/180.

If the two LD monolayers present the same coverage, i.e. the same surface tension, α and θ are equal, and "coverage asymmetry" parameter is 0. When the external coverage is increased, i.e. external surface tension decreases, α increases, and θ diminishes (Chorlay and Thiam 2018, star methods: Determination of the shape and position of an LD in a GUV bilayer ). This results in an increasing of the "coverage asymmetry" parameter. In the limit of a drastic improvement of the external monolayer coverage, provoking a full emergence of the LD, θ is equal to zero, α to 180 degrees and the "coverage asymmetry" parameter is equal to 1. O the opposite configuration, i.e. strong improvement of the external monolayer coverage, If the LD would emerge in the opposite direction, and the "coverage asymmetry" parameter would the be -1.

Determination of the shape and position of an LD in a GUV bilayer (adapted from Chorlay and Thiam Biophysical Journal 2018)

Considering that the aLD's shape is driven by the equilibrium of the three surface tensions (bilayer , external monolayer and internal monolayer ) (Eq. 1), can be projected on the bilayer axis and lead to (Eq. 2).

0

(1) cos cos

(2) sin sin

Moreover, the conserved volume of the LD and the GUV gives two other equations (Eq. 3,4):

4 Π 3 ! 2 cos θ δ 3 cos θ δ sin θ δ 2 cos α δ 3 cos α δ sin α δ # (3) $%& 4 Π 3 ! 2 cos δ 3 cos δ sin δ - 2 cos α δ 3 cos α δ sin α δ #
(4) In order to determine the shape of the droplet, we only need three parameters: alpha, theta and delta, as a function of the tensions , and .

Rearranging these equations leads to a set of 3 equations (Eq. 5,6,7) that can be solved numerically.

The following equations (5, 6) Gives alpha and theta:

cos θ ( ( ( 2 5 cos α ( ( ( 2 6
The last parameter delta is determined numerically, keeping the volumes constant (Eq. 7):

$%& 2 cos θ δ 3 cos θ δ sin θ δ 2 cos α δ 3 cos α δ sin α δ 2 cos δ 3 cos δ sin δ - 2 cos α δ 3 cos α δ sin α δ (7)
The simulation displayed in Fig 2I,J are obtain using this mathematical model and varying the Vld/Vguv parameter.

Proteins production and purification

Cytosol extract was obtained from hepatocyte Huh7.5 cells: confluent monolayers of Huh7.5 cells were placed on ice, washed twice with ice-cold PBS at pH 7.4, and then incubated with 10 mM Tris/HCl, pH 7.4 buffer for 5 minutes on ice. The cells were scraped into a homogenization buffer comprising 10 mM Tris/HCl, 1 mM EGTA, 0.5 mM EDTA and 0.25 M sucrose, pH 7.4, which also contained Complete TM protease inhibitors. The cells were homogenized by twenty in/out passages within a syringe fin needle. The homogenate was transferred to ultracentrifugation tube (2 ml) and were centrifuged for 1 h at 100 000 G at 4°C. The resulting supernatant was used as a cytosol extract.

Fluorescently labeled Arf1 was generated using an Arf1-variant in which the single cysteine residue of Arf1 was replaced with serine, and the C-terminal lysine was replaced with cysteine, yielding Arf1-C159S-K181C. Published work has demonstrated that exchanging the C-terminal lysine of the small GTPase with a Cys-residue, and subsequent fluorescent labeling (using thiol-reactive dyes on Cys181), does not inhibit Arf1-function [START_REF] Beck | Membrane curvature induced by Arf1-GTP is essential for vesicle formation[END_REF]. In short, human Arf1-C159S-K181C and yeast N-myristoyltransferase were coexpressed in Escherichia coli supplied with BSA-loaded myristate. Cell lysates were subjected to 35% ammonium sulfate, and the precipitate, enriched in myristoylated Arf1, was further purified by DEAE-ion exchange. Eluted fractions of interest were concentrated in spin-column filters with a 10-kD cutoff (Millipore), and fluorescently labeled using Cy3maleimide (GE Healtcare) according to the manufacturer's protocol. To remove excess dye, samples were purified by gel filtration using a Superdex 75 column (GE Healthcare).

Protein enrichment of the DEV external leaflet

A DEV hold by a glass micropipette is observed by confocal microscopy. 5 µL of protein solution is injected in the 100 µL bulk phase of the DEV solution. The resulting effective concentrations were respectively: Arf1 (1µM); CAV159-178 (5µM); Alpha-Synuclein (5.5µM); ApolipoproteinB (0.1 µM); Plin1 fragment 93-192 (4.7µM); BSA (75µM); The DEV is then followed for ~ 40 min. Solutions were injected at similar spots from the pipette.

Micromanipulation & surface tension measurements by micro aspiration

Micro-pipettes were made from capillaries drawn out with a Sutter Instruments pipette-puller. They were used to manipulate the LD-embedded GUVs in order to get a side view of the system. The pipettes were incubated for 30 min in a 5% bovine serum albumin solution (BSA) before use, in order to prevent droplets and membranes from adhering to the glass. Additionally, surface tensions were measured and modulated using the same pipettes. As shown in (Figure 1I), the micromanipulation of the external LD monolayer enables the measurement of the eternal monolayer surface tension. Using Laplace's law, and the measurement of the pipette inner diameter, droplet diameter, and suction pressure, the surface tension of the interface can be determined:

∆Psuc 2 / 1 1 1 Rd 4
where ∆Psuc, Rp and Rd are the suction pressure, the pipette radius and the droplet external radius.

The suction was carried out using a syringe.

Pendent droplet interfacial tension measurements

A pendent droplet tensiometer designed by Teclis Instruments (Longessaigne, France) was used to measure the interfacial tension of oil/water interfaces. All experiments were conducted at room temperature. To create triolein/buffer interfaces, triolein drops (from 5 to 16.0µL) were formed at the tip of a J-needle submerged in 5 mL of HKM buffer.

Triolein-protein surface tension measurement: 5 µl of the solution containing the proteins is added to the bulk buffer while the tension of a triolein/buffer interface is measured. The resulting concentrations of proteins were: Alpha-Synuclein (0.1µM); BSA (1.5µM); ApolipoproteinB (2nM); Plin1 108-137 (1µM); Plin1 fragment 93-192 (0.09µM). When tension stabilizes, the interface is compressed. The tension at maximum coverage was determined when wrinkles appeared on the surface of the droplet. If no wrinkles were observed, the compression was stopped, the max packing tension was determined after waiting for the relaxation and re-stabilization of the tension.

Triolein-phospholipid surface tension measurement: to create phospholipid monolayers at the interface, phospholipids were dissolved in the triolein (0.01% w/w). After forming a droplet, the PLs relocate at the interface. The Droplet is then compressed until a tension plateau is reached. This plateau of tension is the tension corresponding to the maximum PL coverage, and evoked as the minimal tension reachable.

Lyso-PC enrichment to the GUV external leaflet

A micellar solution of lyso-PC (18:1) in HKM at 10x the CMC is prepared by desiccating the phospholipids dissolved in chloroform and rehydrating them in HKM buffer. In order to get a fluorescent signal, 1% (w/w) of NBD-PE is added to the lyso-PC. The resulting solution is then vortexed and sonicated for 5 min, three times. 5 to 20 µL of this micellar solution is then directly added to the 100 µL GUVs solution already placed under the microscope. This system is then observed with and without LDs embedded in the GUVs.

Molecular Dynamics (MD) simulations

We built lipid bilayer systems consisting of dioleoylphosphatidylcholine (DOPC) containing triolein (TO) lenses starting from a homogeneous distribution of TO in a DOPC bilayer. All systems featured the same number of TO molecules (7500) and a variable number of DOPC lipids, starting from a symmetric system with 9072 DOPC molecules per monolayer and reducing progressively the number of DOPC molecules in one leaflet, in steps of 100 lipids, to reach a maximum imbalance of 1000 lipids (11% imbalance). For each system, we carried out molecular dynamics (MD) simulations of at least 2 µs, and used the final snapshot as a starting point to build the subsequent stage. The reference symmetric system (9072 lipids in each bilayer) was simulated for 30 µs, and the final system with 11% imbalance was simulated for 10 µs. All systems were simulated at the coarse-grained level, using the MARTINI force field (v2.0, lipid version 2015-06 except for triolein, optimized specifically for the present work). All simulations were carried out with GROMACS v5.1.4, using a leap-frog stochastic dynamics integrator (sd option in GROMACS), with a time step of 20 fs. Non-bonded interactions were calculated with a cutoff of 1.1 nm, using a Verlet neighborlist scheme (Verlet buffer tolerance of 10 -6 kJ mol -1 ps -1 per atom); electrostatic interactions were calculated with a dielectric constant of 15 and shifted from 0 nm, Lennard-Jones were shifted from 0.9 nm. All simulations were run at constant temperature (300 K) and pressure using semi-isotropic pressure coupling (Parrinello-Rahman barostat, time constant of 20 ps, 1 bar in xy and z dimensions), resulting in overall zero surface tension in the system.

Time scale of LD emergence and phospholipid flipping

In a bilayer containing a lipid droplet, any asymmetry in monolayer coverage is spontaneously absorbed (below a second) by the re-positioning of the LD in the bilayer (i.e. its emergence). Indeed, micro-manipulation experiments showed that it is a mechanical equilibrium that is reached quicker than we can see (∆t < 40ms, from our imaging frame rate) [START_REF] Chorlay | An Asymmetry in Monolayer Tension Regulates Lipid Droplet Budding Direction[END_REF]).

An asymmetry can be generating in ER by inserting phospholipids on the outer leaflet. This excess of phospholipids has thus to be detected by a LD for emerging. To assess the time scale of this event, let's consider a 1m long ER and a speed of propagation of the lateral pressure (i.e. phospholipid density) on a monolayer of 1 m/s, despite it has been estimated around 100 m/s (Griesbauer et al. 2009). In this setting, a LD will sense a fabricated PL in few 10 ms. Consequently the increase of the lateral pressure induced by this new phospholipid will be instantaneously detected by the droplet, which will therefore emerge on this side of higher lateral pressure.

Based on these above points, the time frame to rectify a bilayer asymmetry by the re-positioning of a lipid droplet is in the tenth ms range. This time is much smaller than the flip-flop time of a phospholipid in a bilayer which is in the order of minutes as reported in vitro and in vivo [START_REF] Pomorski | Lipid flippases and their biological functions[END_REF], which could rectify the asymmetry.

Altogether, if an asymmetry is created between a bilayer leaflets, a lipid droplet will almost instantaneously expend toward the denser monolayer side and absorb the asymmetry. The more asymmetry created the more the droplet will bulge toward the monolayer with excess phospholipids. The flipping of phospholipids will not be fast enough to rectify the asymmetry between the leaflets.

Lipid droplet induction and fluorescence microscopy of yeast cells

Yeast cells grown as described in the EXPERIMENTAL MODEL AND SUBJECT DETAILS section. For LD induction, cells were grown to exponential growth phase, pelleted, and resuspended in YPD + 0.1% oleic acid (Sigma, #O1008) where indicated. In inositol supplementated experiments, cells were grown in synthetic media lacking inositol, in presence (0.1%) or absence of oleate. After 18h growth, cells were harvested by centrifugation and each culture was resuspended and diluted in the same media in presence of absence of 75µM of inositol. Cells were imaged at 0h, 6h and 24h upon inositol addition. Similar timepoints were collected for lipid analysis. Fluorescence microscopy was performed at room temperature using the Zeiss Cell Observer HS equipped with a CMOS camera (Hamamatsu ORCA-Flash4.0), controlled by 3i Slidebook6.0 software. A Plan-APOCHROMAT 100x 1.4 objective was used. mNeonGreen and MDH signals were detected using the GFP and DAPI filters respectively, at 600ms exposure.

Electron microscopy experiments in yeast cells

Yeast cells grown as described above were subjected to high pressure freezing and processed as described in [START_REF] Wang | Seipin and the membrane-shaping protein Pex30 cooperate in organelle budding from the endoplasmic reticulum[END_REF]. Alternatively, 10 ODs of exponentially growing cells were subjected to chemical fixation in 1% glutaraldehyde, 0.2% PFA in 40mM potassium phosphate pH 7 for 12-16h at 4C. Fixed cells were further processed as described in [START_REF] Choudhary | A conserved family of proteins facilitates nascent lipid droplet budding from the ER[END_REF]. All grids were imaged in a FEI Tecnai 12 TEM operated at 120 kV using a Gatan OneView digital camera.

Thin Layer Chromatography and Lipid Detection

Lipids were extracted and separated on silica TLC plates (Merck) either in chloroform/methanol/acetone/acetic acid/water (100:20:40:20:10) for phospholipids or in petroleum ether/diethyl ether/acetic acid (90:10:1) for neutral lipids, and detected by iodine vapor staining. Bands were identified by comparison with standards. The quantification of the bands were done using Fiji.

dsRNAs construction for Drosophila experiments dsRNAs targeting CCT1 were prepared according to standard procedures (e.g. Beller et al. PLoS 2008). In brief, we amplified targeting sequences pre-computed from the Drosophila RNAi screening center (www.flyrnai.org) from genomic DNA using oligonucleotides fused to a T7 RNA polymerase promotor (lower case in the oligonucleotide sequences). For CCT1 we used the oligonucleotides corresponding to DRSC08157: fwd: gtaatacgactcactataggATCGGAAGTTCAGGATCAGC , rev: gtaatacgactcactataggCCCGCTGAAGAATTGCTG, or DRSC27869: fwd: gtaatacgactcactataggGATGGCCACCTCATCGATAC, rev: gtaatacgactcactataggCTTTGTGAGATGGTGCGTGT Following PCR amplification we performed an in vitro transcription reaction with the TranscriptAid T7 High Yield Transcription Kit (Thermo Fisher Scientific) or the HiScribe T7 High Yield RNA Synthesis Kit (New England Bioloabs) and purified the dsRNA molecules using the Qiagen RNeasy Mini Kit according to the manufacturer's instructions. The dsRNA was stored at -80°C until use.

Structured illumination microscopy of Drosophila experiments

For the targeting experiment analyzed by structured illumination microscopy we seeded 1 mL of 2.5 x 10[5] cells/mL Drosophila Kc167 cells in wells of 12-well tissue culture plates (Sarstedt, Numbrecht, Germany). Cells were imaged using a Zeiss Elyra structured illumination microscope and a Plan-Apochromat 63x/1.4 oil objective with Z-stacks. Cells were grown in Schneider's Drosophila medium with L-Glutamine including 10% heat inactivated fetal calf serum (medium and serum from PAN Biotech, Aidenbach, Germany) with Penicillin/Streptomycin. Following a 6 to 8 hours recovery phase we transfected the cells with 500 ng of the GFP-tagged Sec61beta expression construct (expression is driven by an actin promotor; kind gift from Tobias Walther) and 500 ng of the respective dsRNA or a water only control using the Effectene transfection reagent from Qiagen (Hilden, Germany) according to the manufacturer's instructions. After three days the cells were serum starved overnight (serum free medium with 0.2% fatty acid free bovine serum albumin) to deplete the cells of already existing LDs. On the next day, the cells were resuspended and transferred to high precision cover slips and allowed to adhere for one hour. Subsequently, the medium was replaced with medium containing 600 µM oleic acid containing 1 µg/mL BODIPY558/568 labeled C12 fatty acids (Thermo/Molecular probes) and incubated for six hours. Subsequently the cells were fixed with 4% paraformaldehyde and embedded in Mowiol containing antifades.

qPCR confirmation of CCT1 knock-down in Drosophila experiments

We isolated RNA from the frozen cells using the rNeasy Mini Kit (Qiagen). qPCR was performed according to standard procedures with oligos for rp49 (fwd: ATCGGTTACGGATCGAACAA, rev: GACAATCTCCTTGCGCTTCT) and CCT1 (fwd: GGAAGCGGACCTACGAGATA and rev: GTGCCCTGATCCTGAACTT).

Electron microscopy experiments of Drosophila cells

We added 30 µg of dsRNA or water only as a control to 6-well plates. Subsequently, we added 1 mL of 2 x 10[5] cells/mL Drosophila Kc167 cells in serum free Schneider's Drosophila medium. After 45 min incubation, we added 3 mL of complete Schneider's medium and incubated for four days to allow for knock-down of the target by RNA interference. Subsequently, we added 600 µM OA to the cells and incubated for six hours. Cells from one well per sample were washed with PBS and frozen at -80°C to allow later RNA extraction for qPCR analysis. Cells from a second well of each sample were washed with 0.1 M phosphate buffer and afterwards fixed with 2.5 % glutaraldehyde in 0.1 M phosphate buffer for 30 min. After another wash with 0.1 M phosphate buffer and the cells were mixed with 3 % low melting agar in PBS. Agar was cut into small 1 mm square blocks, washed three times with 0.1 M sodium cacodylate buffer before incubation in 1 % osmium, 10 mg/mL potassium ferrocyanide in 0.1 M sodium cacodylate buffer, followed by three more washed in the cacodylate buffer and an ethanol dehydration series. After 100 % ethanol was added, there were two more washes with epon:ethanol (1:1) and epon:ethanol (3:1), respectively. Subsequently, the blocks were incubated with fresh epon at 4 °C over night, and two more hours at room temperature, followed by curing at 60 °C for 48 to 72 hours. Ultrathin sections (70 µm) were cut from the blocks using an ultramicrotome (Leica, UC7) and stained with 1.5% uranyl acetate for 20 min and Reynolds lead citrate solution for 3 min. Electron micrographs were recorded with a JEM-2100 Plus Transmission Electron Microscope (JEOL) operating at 80 kV with a Camera OneView 4k 32 bit (Gatan) and the software DigitalMicrograph (Gatan).

Modulation of GUVs bilayer tension using osmotic pressure

GUV membranes are water-permeable, thus an osmolarity difference between the lumen and the exterior of the GUV results in a deflation or swelling of the GUV. This increases or decreases the bilayer tension. It is then possible to increase the bilayer tension osmotically with a hypotonic external buffer (e.g. adding MQ water to the HKM buffer surrounding the GUV; standard procedure is 10 µL of MQ water added to the 100 µL of HKM buffer). The bilayer tension can be decreased using a hypertonic external buffer (5 µL of 1M NaCl solution added to the 100 µL of HKM buffer).

Calculation of the amount of removed phospholipids

When mechanically depleting the external leaflet of the GUV-LD system (see Figure 6E), we remove a portion of the PLs by ablating a part of the external monolayer. Assuming that the repartition of the PLs is homogenous on the leaflets, the number of PLs removed from the external leaflet is proportional to the ablated area of leaflet. Then the percentage of removed PLs can be calculated by: area of the removed tongue over total external leaflet area. The internal leaflet remains intact, so the number of PLs on this leaflet is constant.

QUANTIFICATION AND STATISTICAL ANALYSIS

Quantification methods in yeast Aberrant ER: Yeast cells were enumerated according to their ER morphology using a Z projection of Z-stack images. If the cortical and nuclear ER were clearly defined, the cell was considered regular, if unusual ER membrane signals appeared in the cytoplasm, forming of lumps or threads, the cell was considered as aberrant. Number of LD in Aberrant /Regular cells: Two sets of cells were selected according to the morphology of their ER (see above). Ambiguous cells were discarded. In these two populations the number of LD was quantified in approximately 40 cells in each subset. LD in large contact with the ER: the proportion of cells presenting at least one event of LD presenting a close contact with ER (on half of its surface or more) was quantified over the total number of cell.

Quantification in Drosophila cells

LD in close contact with the ER: the number of LD surrounded or more than semi surrounded in the ER is quantified over the total number of LD in the cells. LD size distribution: After preforming a Z projection with the Z-stack images of the LD and a thresholding, a Matlab program calculated the surface of the individual LDs. The statistics on the LD radius was maid (considering the LD as spherically shaped).

Statistical analysis

The statistical comparisons where made using a non-parametric t-test (GraphPad Prism 7.0a; *** indicates p < 0.0001 ** indicates p < 0,001 * indicates p < 0.01). Unless mentioned, all values shown in the text and figures are mean ± S.D.

Data and software Availabity

All simulations were carried out with GROMACS v5.1.4.

  CCT1-depleted cells similar LD phenotypes as in yeast (Figure4A,C,D; FigureS4A,C): as compared to WT control cells, many LDs were much more often in close contact with the ER membrane marker (Figure5A), as visualized through z-stacks. For many LDs, there were focal planes where the ER membrane marker protein was extremely close to the LD surface (Figure5A, see right examples), and

B

  ) Top: Schematic of an artificial LD (aLD) embedded in a bilayer as during LD formation. Bottom: principle of the droplet-embedded vesicle (DEV) system, made by an aLD incorporated into the intermonolayer space of a giant unilamellar vesicle (GUV).C) LD emergence is characterized by the fraction of its external and internal surface areas. The imbalance of monolayer tension between the interior and exterior, reflecting the asymmetry in surface coverage, defines the coverage asymmetry parameter (see STAR Methods). D) Arf1 binding to DEVs promotes artificial LD extraction. Left: a PC DEV with a triolein droplet is made and Arf1-Alexa488 added to the external medium. Arf1 DEV-recruitment occurred only with the subsequent addition of GTP. The artificial LD emerged towards the exterior upon Arf1 binding. The glass pipette was used for softly maintaining the DEV under the observatory region during injection; the pipette was motionless; images were rotated for visualization. Top right: external and internal surface areas corresponding to the pictures shown on the left. Bottom right: corresponding time course displaying the protein recruitment effect on external Area ("Aext var" represent the external area variation). Arf1 recruitment on the bilayer and droplet surface is displayed in Figure S1A. Additional images of fully emerged aLDs following Arf1 binding are displayed in Figure S1B. Scale bar is 10 µm. E) The fragment of Caveolin 159-178, used in Hpos construct which targets LDs (Kassan et al., 2013), bound to DEVs and promoted aLD extraction. Left: Cav 159-178-NBD was added to the medium of a PC DEV containing triolein droplets. The artificial LD emerged toward the exterior upon Cav159-178-NBD binding. The glass pipette was used for softly maintaining the DEV under the observatory region during injection Top right: external and internal surface areas corresponding to the pictures shown on the left. Bottom right: effect of Cav 159-178-NBD recruitment on artificial LD external Area. ("Aext var" represent the external area variation). Scale bar is 10 µm. F) Alpha-Synuclein did not bind to PC DEV (left), even after 30 min (right), and the artificial LD did not emerge. Arrows point to the GUV and droplet interface. The external area did not vary. Scale bars are 10 µm. G) A cytosolic extract from Huh7.5 cells and apolipoprotein B promoted artificial LD emergence from DEVs, while Plin1 93-192 or BSA did not. Corresponding external and internal surface areas variation are shown below each condition. For fluorescence confocal micrographs see Figure S1D. For each of these conditions, external area variation and coverage asymmetry is displayed over time in Figure S1D. Scale bars are 10 µm. H) The induced effects of the proteins on the aLD external surface area variation (top) and coverage asymmetry (bottom) are displayed. Error bars are measurement precision. I) The external monolayer tension of the artificial LD is measured by micro-aspiration technique (left images -fluorescent signal is rhodamine-PE (Rh-PE)), before and after addition of Apolipoprotein B and Plin1 93-192. A decrease of tension was measured when Apolipoprotein B was added, while tension remained constant with Plin1 93-192. Scale bar is 10 µm; error bars are measurement precision.

Figure 2 :

 2 Figure 2: Artificial LDs emerge toward the monolayer bearing excess phospholipids A) A Lyso-PC micellar solution containing NBD-PE (1% w/w to lyso-PC) was added to GUVs marked by Rhodamine-PE (Rh-PE) (0.5% w/w to PC). Lyso-PC molecules, reported by NBD, are recruited to the external GUV surface see arrows (and Figure S2B,C). Scale bar is 10 µm. B) Determination of the location of recruited lyso-PC molecules by following NBD-PE signal. Dithionite, an NBD quencher, is added to the external solution. The NBD signal disappears entirely, suggesting that lyso-PC enrichment occurred only on the external leaflet. Scale bar is 10 µm. C) Relative NBD signal on the bilayer displayed for different GUVs: before any treatment, after Lyso-PC micelle addition, and after dithionite treatment. Data are represented as mean ±SD. D) A Lyso-PC micellar solution containing NBD-PE was added to a DEV. When the NBD concentration around the DEV increased (See example in Figure S2B), the embedded droplet emerged towards the exterior. Scale bar is 10 µm. E) The droplet external area variation in (D) is displayed as function of the Lyso-PC/NBD-PE enrichment on the DEV. Error bars are measurement precision. F) Simulated situation with increasing the number of PC molecules in the external leaflet of a bilayer containing a 35 nm-sized triolein droplet. G) Examples of cross sections of multiple merged snapshots of the results for different PC imbalances (0%, 4.4%, 11% PC excess in upper leaflet). See Figure S2D for more examples. Scale bar is 7 nm. H) The aLD external area variation in (G) shows the externalization of the droplet with increased PC imbalance. Related to Figure S2D. Error bars are measurement precision.I) The external area fraction of aLDs in a DEV is displayed against the aLD radius over the GUV radius (Rld/Rguv), which takes into account the bilayer curvature. Red: values for experiments where phospholipids or proteins (that bound were added to the external medium. Black: no addition of materials. Error bars are measurement precision. J) Simulated relative aLD external area vs. Rld/Rguv with varying the coverage asymmetry (0, 0.2, 0.6, 0.98). K) On a flat and finite bilayer a large droplet will be in the middle of the bilayer, with external and internal areas being equal (top); curvature the bilayer will induces only partial emergence (middle); providing phospholipids to increase the coverage asymmetry alone (bottom) is sufficient to promote full emergence.

Figure 3 :

 3 Figure 3: Boosted LD formation is concomitant with remodeled ER morphology A) Representative images of wild-type (WT) Saccharomyces cerevisiae cells in stationary phase, imaged after 24 h, with or without treatment with oleic acid (OA). The ER marker protein, Sec63, was endogenously tagged with monomeric NeonGreen (Sec63-mNG), and LDs were labeled by monodansylpentane (MDH). Images shown represent z-projection of different confocal planes. Scale bar is 5 µm. B) Left: schematics and representative images of cells with regular or aberrant ER. An image is shown for each situation. Images shown represent z-projection of different confocal planes (shown in Figure S3F) Scale bar is 2 µm. Right: Quantification of the number of cells with aberrant ER, 0 h, 6 h and 24 h after OA treatment of cells in log phase. Control cells with no OA addition are shown at 24 h time point. N >200 cells/condition (Ntotal = 1276). Data are represented as mean ±SD. See Figure S3A-C for other tested conditions. C) Cells with the aberrant ER phenotype statistically contained more LDs than cells with a normal ER. t-test (p<0.0001), N = 140 cells, data are represented as mean ±SD. Similar results for pct1∆ are presented in Figure S3E. Scale bar is 5 µm. D) Left: illustrative image showing the measuring principle of total LD area, and cortical and nuclear ER surface. Scale bar is 2 µm. Right, quantification of the total LD over ER area at 24h in stationary phase with and without oleate. T-test show significant difference (p<0.0001); N=10 cells. Data are represented as mean ±SD. E) Left: Representative images of cells in stationary phase, imaged after 24 h, with or without with (OA). Right: the cells were additionally treated with 10µM inositol. Scale bars are 5 µm. F) Total amount of phospholipids/µg of proteins in the presence of OA, with and without inositol treatment. Quantification of thin layer chromatography (Figure S3H). Experiment was repeated three times. Data are represented as mean ±SD. G) Quantification of the number of cells with aberrant ER at 24 h for various inositol concentrations, with and without OA treatment (conditions displayed in E). N>200 cells/condition (Ntotal = 1979). Data are represented as mean ±SD. H) Cells presenting a normal ER statistically bear more LDs in inositol treatment. N = 30 cells (p<0.0001), data are represented as mean ±SD.

Figure 4 :

 4 Figure 4: LD proper emergence in Saccharomyces cerevisiae is facilitated by phospholipid production A) Saccharomyces cerevisiae cells in logarithmic growth phase treated with OA show LDs with large contact zones with the ER membrane (white arrows). Less LD-ER contact zones were observed after inositol treatment, as in non-oleated cells. Sec63-mNG marks ER and LD are MDH stained. Scale bar is 5 µm. B) Quantification of the number of cells presenting large ER-LD contact (with or without inositol (10 µM)): ER-LD contact is considered large if more than half of the LD surface is in contact with ER

Figure 5 :

 5 Figure 5: Drosophila cells with decreased CCT1 display increased ER-LD contacts A) Drosophila cells were depleted of CCT1 by dsRNA. LDs are induced by oleate and stained by BODIPY558/568; GFP-sec61β marks the ER. Top: representative confocal image of dsRNA CCT1 Cells. Insets show z-stacks of LDs in close contact with the ER membrane marker. Bottom left: representative WT control with z-stacks of example LDs. The ER-LD contact was less pronounced. Bottom right: dsRNA CCT1 cells present a higher number of LDs in close contact with ER (quantification made for N =150 in each condition). Scale bar are 2µm and 200nm (inset). See Figure 5SB for additional z-stacks. B) WT and CCT1 depleted Drosophila cells treated with OA were fixed by glutaraldehyde and observed by electro-microscopy following contrastation with uranylacetate and lead. In the section plane, three different LD-ER interactions are defined: top, LDs with no visible contact with ER; middle, LDs with multiple contact point with ER membrane; bottom: LDs in large contact, with a noticeable ribosome free zone delineated by the ER membrane. Frequency of these phenotypes in WT and dsRNA CCT1 cells is shown on the right with an average of fifteen LDs each. More Illustrative EM images are presented in Figure S5E. Scale bar are 2µm and 200nm for zoomed insets. See also supplementary Figure S5.

Figure 6 :

 6 Figure 6: Artificial LDs equally emerge on both sides of a same bilayer with fixed phospholipid number A) Example of a confocal image of DEV with small artificial LDs; the DEV contains other aLDs which were not visible in the shown confocal plane. Bodipy stained the droplet and Rhodamine-PE (Rh-PE) labeled the membranes. The inset is zoomed. The graph shows the quantification of the total artificial

Figure 7 :

 7 Figure 7: Proposed model for the impact of proteins and phospholipids on LD emergence A) Recruitment of both proteins and phospholipids to one monolayer promotes LD emergence toward this monolayer side. B) A balance of strongly inserted proteins will position a LD in the bilayer with various degree of contact to the surrounding aqueous phases. C) Continuous injection of phospholipid on the upper monolayer promotes LD emergence to the upper side. D) Lack of a monolayer phospholipid excess, or supply, induces LD equal emergence to both membrane sides, membrane stress and reorganization.

  Fig 5

TABLE REAGENT

 REAGENT Unless mentioned, in vitro experiments were performed in the following HKM buffer: 50 mM Hepes, 120 mM Kacetate, and 1 mM MgCl2 (in Milli-Q water) at pH 7.4 and 275±15 mOsm.

	Fatty acid free Bovine Serum Albumin (BSA)	Sigma	A8806
			Aldrich
	Oleic acid		Calbiochem	4954
	Critical Commercial Assays
	N/A	or RESOURCE	SOURCE	IDENTIFIER
	Antibodies Deposited Data
	N/A N/A
	Bacterial and Virus Strains Experimental Models: Cell Lines
	N/A S. cerevisiae: (BY4741) MATa	This paper	yPC10613
	Bacterial samples ura3∆0his3∆1leu2∆0met15∆0 SEC63-mNeonGreen-
	N/A HIS
	S. cerevisiae: (BY4741) MATa Chemicals, Peptides, and Recombinant Proteins Phospholipids (DOPC, Rhodamine-DOPE, LysoPC (18:1)) ura3∆0his3∆1leu2∆0met15∆0pct1::KAN SEC63-mNeonGreen-HIS Triolein Human hepatocytes Huh7.5 Bovine Serum Albumin (BSA) Dithionite Drosophila Kc167	This paper Avanti Polar Lipids Sigma Apath Missouri Aldrich Sigma Sigma Aldrich center screening Aldrich Harvard RNAi	yPC10672 https://avantilipids.com/ APC 49 T7140 71699-250G https://fgr.hms.harvard. A7906-100G edu
	Apolipoprotein B from human plasma Experimental Models: Organisms/Strains	Sigma Aldrich	SRP6302
	Plin1 108-137, Purity 96.83% (NBD-N/A	ProteoGenix,	https://www.proteogeni
	PPEKIASELKDTISTRLRSARNSISVPIAS) Oligonucleotides	France	x.science/fr/
	CAV fragment 159-178, Purity 96.83% (NBD-LFEAVGKIFSNVRINLQKEI) CCT1_RNAi_fwd: Alpha-Synuclein gtaatacgactcactataggATCGGAAGTTCAGGATCAG C CCT1_RNAi_rev: Plin1 fragment 93-192 gtaatacgactcactataggCCCGCTGAAGAATTGCTG CCT1_RNAi_fwd: gtaatacgactcactataggGATGGCCACCTCATCGATA C Fluorescently labeled Arf1 CCT1_RNAi_rev: gtaatacgactcactataggCTTTGTGAGATGGTGCGTGT Monodansyl pentane (MDH) rp49_fwd: ATCGGTTACGGATCGAACAA	Peptide 2.0, USA Harvard Bruno RNAi Screening Antonny's Center lab, Harvard IPMC CNRS Pr. David RNAi Screening Savage Center (Cambridge Harvard Metabolic RNAi Research Screening Laboratories, Center UK). This study, Harvard RNAi see methods Screening details Abgent Center Rajan et al.,	https://www.peptide2.co DRSC08157 m/ N/A DRSC08157 N/A DRSC27869 DRSC27869 N/A SM1000a N/A
	Oleic acid		Sigma Cell Metab.	O1008-1G
			Aldrich 2012; 151:
	Inositol		Sigma 123	I5125-50G
	Yeast nitrogen base with ammonium sulfate rp49_rev: GACAATCTCCTTGCGCTTCT	MP Biomedicals Cell Metab. Rajan et al.,	4027-522 N/A
	Yeast nitrogen base with ammonium sulfate, without	MP 2012; 151:	4027-412
	inositol		Biomedicals 123
	CSM-HIS-LEU-TRP-URA CCT1_qPCR_fwd: GGAAGCGGACCTACGAGATA	MP Copic et al.,	4550-022 N/A
			Biomedicals Nat
	Bacto yeast extract	BD Commun.	212750
	Bactopeptone Fluorescently labeled C12 fatty acid	BD 2018; 9: Thermo 1332	211677-500G D3835
			Fisher /
			Molecular
			Probes

Acknowledgments

We are thankful to Drs. David Savage and Bruno Antonny for the kindly gift of proteins, Alisa Gahlen and Elisabeth John for technical support and Astrid Schauss and Felix Babatz from the CECAD Imaging Facility for EM imaging. We thank all the group members for their valuable comments and critical discussions. This work was supported the ANR-NANODROP to ART and LM, ANR-LDEN, ANR-MOBIL, ANR-18-CE11-0012-01, and Paris Sciences et Lettres to ART.

Author contribution

AC and ART designed research, experiments and quantified all data. AC performed all in vitro experiments, with the help of KBM and MO, and DA performed lipid analysis. SW, JF, EJ and PC performed the yeast and the electron microscopy experiments. MB did the studies in Drosophila cells, and LM the molecular dynamics studies. RB purified proteins and helped on the elaboration of in vitro experiments. ART wrote the manuscript, and all the authors reviewed and edited it.

Declaration of interests

The authors declare no competing interests.

CONTACT FOR REAGENT AND RESOURCE SHARING

Further information and requests for reagents may be directed to, and will be fulfilled by the corresponding author Abdou Rachid Thiam (thiam@ens.fr).

EXPERIMENTAL MODEL AND SUBJECT DETAILS Yeast strain and culture media conditions

Saccharomyces cerevisiae cell strains used in the experiments are yPC10613 (MATa ura3∆0his3∆1leu2∆0met15∆0 SEC63-mNeonGreen-HIS) and yPC10672 (MATa ura3∆0his3∆1leu2∆0met15∆0pct1::KAN SEC63-mNeonGreen-HIS). Cells were grown at 30 °C in YPD liquid medium (1% yeast extract, 2% peptone, 2% glucose).

Huh7.5 culture and media conditions

Huh7.5 were maintained in Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% heat inactivated fetal bovine serum (Life Technologies), 0.1 gL-1 sodium pyruvate (Life Technologies) and 1% penicillin-streptomycin (Life Technologies). The cells were cultured at 37°C under a 5% CO2 atmosphere.

Drosophila Kc167 culture media conditions

Drosophila Kc167 cells were grown in Schneider's Drosophila medium with L-Glutamine including 10% heat inactivated fetal calf serum (medium and serum from PAN Biotech, Aidenbach, Germany) with Penicillin/Streptomycin.

METHOD DETAILS DEV preparation: GUVs + LDs