# Seipin Governs caveolin-1 trafficking through modulating sphingolipid-glycerolipid balance

Maxime Carpentier, Mohyeddine Omrane, Rola Shaaban, Jennica Träger, Naïma El Khallouki, Soazig Le Lay, Julie Patat, Marie Palard, Mehdi Zouiouich, Yoann Combot, et al.

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## To cite this version:

Maxime Carpentier, Mohyeddine Omrane, Rola Shaaban, Jennica Träger, Naïma El Khallouki, et al.. Seipin Governs caveolin-1 trafficking through modulating sphingolipid-glycerolipid balance. Cell Reports, 2025, 44 (10), pp.116320. 10.1016/j.celrep.2025.116320. hal-05369874

HAL Id: hal-05369874

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Submitted on 18 Nov 2025

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**Cell Reports**

# Seipin Governs caveolin-1 trafficking through modulating sphingolipid-glycerolipid balance

## Graphical abstract

<figure><img src="image_3.png" /></figure>

## Authors

Maxime Carpentier,
Mohyeddine Omrane, Rola Shaaban, ...,
Francesca Giordano, Xavier Prieur,
Abdou Rachid Thiam

## Correspondence

thiam@ens.fr

## In brief

Carpentier et al. suggest that seipin regulates palmitate flux between sphingolipids and glycerolipids. Seipin loss causes ceramide buildup, trans-Golgi disorganization, and misrouting of caveolin-1 to lipid droplets instead of the plasma membrane. These findings identify seipin as a key regulator of overall lipid metabolism, protein trafficking, and organelle membrane homeostasis.

## Highlights

* Seipin is essential for the correct trafficking of caveolin-1 and caveolae formation

* Loss of seipin alters cellular lipid fluxes and disrupts TGN membrane order

* Excess saturated lipids and ceramides misroute caveolin-1 to lipid droplets in WT cells

* Restoring lipid balance rescues caveolin-1 localization in seipin-deficient cells

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## Article

# Seipin Governs caveolin-1 trafficking through modulating sphingolipid-glycerolipid balance

Maxime Carpentier,<fnref n="1" /> Mohyeddine Omrane,<fnref n="1" /> Rola Shaaban,<fnref n="2" /> Jennica Träger,<fnref n="1" /> Naïma El Khallouki,<fnref n="3" />,<fnref n="4" /> Soazig Le Lay,<fnref n="5" />

Julie Patat,<fnref n="6" /> Marie Palard,<fnref n="2" /> Mehdi Zouiouich,<fnref n="1" /> Yoann Combot,<fnref n="2" /> Takeshi Harayama,<fnref n="8" /> Corinne Vigouroux,<fnref n="9" />,<fnref n="10" />

Kristine Schauer,<fnref n="6" />,<fnref n="7" /> Francesca Giordano,<fnref n="3" />,<fnref n="4" /> Xavier Prieur,<fnref n="2" /> and Abdou Rachid Thiam<fnref n="1" />,<fnref n="11" />,*

## SUMMARY

Dysregulation of caveolin-1 (CAV1), a core component of caveolae, causes pleiotropic disorders; yet, the mechanisms governing its trafficking remain poorly understood. Here, we show that the lipid droplet (LD) biogenesis factor seipin regulates CAV1 localization. Seipin deficiency in mice and HeLa cells resulted in the accumulation of saturated lipids and ceramides, thereby disrupting the membrane order of the trans-Golgi network (TGN). This impaired CAV1 trafficking to the plasma membrane, reduced caveolae formation, and redirected CAV1 to LDs—an effect also observed in seipin-deficient patient fibroblasts. We reproduced this phenotype in wild-type cells by supplementing them with palmitate or ceramide or by inhibiting stearoyl-CoA desaturase 1, indicating that saturated lipids’ accumulation is the root cause. Conversely, blocking fatty acid synthase in seipin knockout cells restored proper CAV1 localization. Our findings suggest that seipin regulates lipid fluxes between glycerolipids and sphingolipids, which is critical for TGN integrity and CAV1 sorting.

## INTRODUCTION

Cells adapt to various demands by adjusting their lipid metabolism, which is crucial for maintaining cell function. This regulation involves the uptake, secretion, and storage of lipids, requiring plasticity in the lipid composition and the mechanical properties of cellular membranes to meet these metabolic needs.<fnref n="1" />,<fnref n="2" /> Dysregulation in these mechanisms is implicated in various disorders.<fnref n="3" />⁻<fnref n="7" /> One of the major players in these processes is caveolin (CAV),<fnref n="3" />,<fnref n="8" />,<fnref n="9" /> which links membrane mechanics to metabolism.<fnref n="1" />

CAV1, CAV2, and CAV3 are three members of the caveolin family of proteins, which are crucial for caveolae formation in different tissues. Caveolae are small invaginations in the plasma membrane (PM) that participate in various cellular processes.<fnref n="10" /> CAV1 is the most widely expressed and abundant protein, found in multiple cell types, including adipocytes, endothelial cells, and fibroblasts.<fnref n="11" /> CAV2, which shares similarities with CAV1, is expressed in more restricted cell types, including endothelial cells

and specific smooth muscle cells. CAV3 is predominantly expressed in muscle cells, particularly in skeletal and cardiac muscle, where it plays a crucial role in muscle function.<fnref n="11" /> Despite their redundancies, CAVs exhibit structural specificities and a distinct impact on biological processes. CAV1, in particular, is essential for various biological processes, including mechanotransduction, lipid metabolism, and endocytosis.<fnref n="10" />,¹²⁻¹⁴ Deregulation of CAV1 dynamics is associated with several pathologies, such as cellular senescence, cancer, insulin resistance, and lipodystrophy.<fnref n="3" />⁻<fnref n="7" /> CAV1’s mechanosensing properties, enabled by caveolae formation, are critical for buffering PM tension fluctuations and modulating diverse signaling pathways. However, the mechanisms governing CAV1 trafficking and its delivery to target organelles remain poorly understood.

When translated into the endoplasmic reticulum (ER), CAV2 and CAV3 enter the secretory pathway but can also diffuse within the ER and target the surface of lipid droplets (LDs),<fnref n="5" />,¹⁵,¹⁶ which originate from and are connected to the ER. This localization

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<footnote n="1">¹Laboratoire de Physique de l'École normale supérieure, ENS, Université PSL, CNRS, Sorbonne Université, Université Paris Cité, Paris F-75005, France</footnote>

<footnote n="2">²Université de Nantes, CNRS, INSERM, l'institut du thorax, Nantes 44000, France</footnote>

<footnote n="3">³Institute for Integrative Biology of the Cell (I2BC), CEA, CNRS, Université Paris-Saclay, 91198 Gif-sur-Yvette Cedex, France</footnote>

<footnote n="4">⁴Inserm U1280, 91198 Gif-sur-Yvette Cedex, France</footnote>

<footnote n="5">⁵University Angers, SFR ICAT, Angers 49000, France</footnote>

<footnote n="6">⁶Cell Biology of Organelle Networks Team, Tumor Cell Dynamics Unit, Inserm U1279 Gustave Roussy Institute, Université Paris-Saclay, 94800 Villejuif, France</footnote>

<footnote n="7">⁷Centre National de la Recherche Scientifique (CNRS), Paris, France</footnote>

<footnote n="8">⁸Institut de Pharmacologie Moléculaire et Cellulaire, Université Côte d'Azur - CNRS UMR7275 - Inserm U1323, Valbonne 06560, France</footnote>

<footnote n="9">⁹Saint-Antoine Hospital, Reference Center for Rare Diseases of Insulin Secretion and Insulin Sensitivity (PRISIS), Department of Endocrinology, Assistance Publique-Hôpitaux de Paris (AP-HP), 184 rue du Faubourg Saint-Antoine, Paris 75012, France</footnote>

<footnote n="10">¹⁰Saint-Antoine Research Center, Institute of Cardio Metabolism and Nutrition (ICAN), Sorbonne University, Inserm UMR_S 938, Paris, France</footnote>

<footnote n="11">¹¹Lead contact</footnote>

<footnote>*Correspondence: thiam@ens.fr</footnote>

<footnote>https://doi.org/10.1016/j.celrep.2025.116320</footnote>
suggests a potential direct role for CAV proteins in lipid and LD
metabolism. In contrast, CAV1 enters the secretory pathway
rapidly¹⁷ and does not localize to LDs under similar conditions.
This difference is likely due to CAV1's rapid assembly into 8S
oligomeric complexes, which subsequently traffic to the Golgi,
particularly the trans-Golgi network (TGN), where they form
higher-order 70S complexes that function at the PM.¹⁰ CAV1
has been observed on LDs only when the Golgi apparatus is dis-
rupted by brefeldin A treatment,¹⁶ which causes the Golgi to
“collapse” into the ER. Although CAV1 does not typically target
LDs, it nonetheless plays a crucial role in lipid metabolism.¹⁸⁻²⁰

CAV1 regulates the transport of lipids and enzymes to various organelles, a process essential for their proper function.¹⁹,²¹,²² For example, it controls the PM translocation and levels of the glucose transporter type 4 (GLUT4),²³,²⁴ the lipid transporter CD36,²²,²⁵ and other membrane transporters,²⁶,²⁷ either directly by incorporating them into caveolae or indirectly through modulation of PM signaling domains. This regulation has significant effects on energy intake, storage, and utilization. Mutations in CAV1 are linked to the genetic disorder congenital generalized lipodystrophy (CGL), a condition marked by the failure to develop adipose tissue (AT), emphasizing a unique role for CAV1 in adipocyte differentiation and lipid homeostasis. Specifically, biallelic null variants in CAV1 cause congenital generalized lipodystrophy type 3 (CGL3),²⁸ characterized by a near-total absence of AT, defects in lipid storage, severe insulin resistance with metabolic complications, and esophageal neuropathy.⁶,²⁸ Similarly, loss-of-function mutations in BSCL2 (Berardinelli-Seip Congenital Lipodystrophy type 2), which encodes seipin, a key regulator of LD assembly and function at the ER,²⁹⁻³¹ lead to congenital generalized lipodystrophy type 2 (CGL2), presenting comparable insulin resistance-associated metabolic complications and cognitive impairment.³²

Seipin assembles into an undecameric complex,<sup>33,34</sup> and in ATs, it can also form dodecamers, creating a donut-like structure in which ADIG proteins are intercalated between seipin monomers.<sup>35</sup> The structural flexibility and functional differences between these oligomeric forms remain poorly understood but may be critical for the regulatory roles of the complex.<sup>36</sup> Seipin is known to regulate the flux of lipids synthesized in the ER into LDs<sup>37,38</sup> and to influence the diffusion or relocation of monotopic proteins from the ER to LDs.<sup>39,40</sup> Consequently, seipin could also modulate the dynamics of caveolins in relation to LDs, although this possibility has not yet been investigated.

Despite the roles of CAV1 and seipin in lipid metabolism and
LD biology, the relationship between these two proteins remains
unknown. Given that the dysfunction of either protein leads to
similar metabolic signatures and neurological involvement, we
hypothesized the existence of a functional axis linking seipin
and CAV1. We investigated such a potential link, which could
provide unprecedented insights into CAV1 dynamics and seipin
function in lipid metabolism.

**Results**

The removal of seipin redirects CAV1 to cytosolic vesicles and lipid droplets, resulting in fewer caveolae

CAV1 is one of the major proteins involved in lipid intake, partic-
ularly in intestinal cells for dietary fat absorption and in adipo-

cytes for lipid storage.¹⁹,⁴¹ Given the critical role of seipin in lipid
storage and adipogenesis,²⁹,³² as well as protein trafficking to
LDs, we investigated whether the dynamics of CAV1 are influ-
enced by seipin.

We first used wild-type (WT) and seipin knockout (SKO) HeLa cells expressing CAV1-GFP at comparable levels (Figure S1A) and observed them by confocal microscopy. In WT cells, overexpressed CAV1 localized primarily to the PM as well as to intracellular vesicles likely representing caveolae-derived endosomal structures (Figure 1A). In striking contrast, SKO cells exhibited a marked reduction in CAV1 signal at the PM. We then quantified the CAV1 signal along the PM contour and normalized it to the total cellular signal (Figure 1B), revealing a significant decrease in PM localization in SKO compared to WT cells. Notably, SKO cells exhibited a pronounced accumulation of CAV1 in numerous intracellular vesicles (Figure 1B). Furthermore, staining with LipidTox, which labels LDs, revealed that CAV1 was often found in proximity to pre-existing LDs, near CAV1 vesicular structures. Inhibition of DGATs after transfection did not rescue CAV1 PM localization, indicating that the observed phenotype is independent of triacylglycerol synthesis (Figure S1B). Collectively, these findings suggest a defect in CAV1 trafficking in the absence of seipin.

Seipin deletion is associated with the formation of supersized
LDs and an altered LD proteome.<sup>38,42,43</sup> To further investigate
the impact of seipin loss on CAV1 localization, we transfected
WT and SKO cells with CAV1-GFP and loaded them with
400 µM OA to induce more visible LDs (Figures 1C and 1D). As
expected, SKO cells displayed larger LDs than WT cells
(Figures 1C and 1D). In WT cells, LDs lacked a distinct CAV1-
GFP signal. In contrast, in SKO cells, LDs prominently recruited
CAV1-GFP to their surface (Figures 1C and 1D), consistent with
our above observation (Figure 1A). This recruitment pattern was
also observed for endogenous CAV1, revealed by immunofluo-
rescence (Figures 1E and 1F) or when the GFP tag was fused
to the N- instead of the C-terminal of the protein (Figures S1C
and S1D). To further characterize the redistribution of CAV1 in
SKO cells, we examined its localization at the PM. Cells were
exposed to a hypotonic medium to induce swelling and a more
spherical morphology, thereby enhancing visualization of the
PM signal. CAV1-GFP signal along the PM contour was quanti-
fied from an equatorial plane and normalized to the total cellular
signal (Figures S1E and S1F). CAV1 redistribution to LDs was
associated with a reduction of the PM signal in SKO cells
(Figures 1C, 1D, S1E, and S1F). Our findings reveal that the
behavior of CAV1 is unique within the caveolin protein family.
Indeed, unlike CAV1, both CAV2 and CAV3 localized to LDs inde-
pendently of seipin (Figures S1G–S1J). Additionally, endoge-
nous and overexpressed Cavin1, another critical component of
caveolae formation, did not accumulate around LDs as CAV1
did (Figures S1K and S1L). This suggests that the CAV1 signal
on LDs did not originate from “caveolae” assembling on LDs
or caveolae-LD contacts. Lastly, overexpression of seipin-
mScarlet in SKO cells restored the PM localization of CAV1 while
decreasing its association with LDs (Figures S2A–S2C). This
strongly indicates that seipin controls CAV1 behavior and its
PM localization.

To validate our findings in a more physiologically relevant
context, we repeated our study using skin fibroblasts from
<figure><img src="image_6.png" /></figure>

<figure><img src="image_7.png" /><figcaption>**Figure 1.** Seipin depletion disrupts CAV1 dynamics</figcaption></figure>

(A) HeLa WT and SKO cells overexpressing CAV1-GFP. LipidTox Deep Red labels endogenous LDs.

(B) Top: percentage of CAV1-GFP PM signal per cell in (A). Two-tailed Mann-Whitney analysis. *p* < 0.0001, *n* = 41 WT and 29 SKO cells; three replicates. Bottom: total CAV1-GFP vesicle area per cell in (A). Two-tailed Mann-Whitney analysis. *p* = 0.0041, *n* = 15 WT and SKO cells each; three replicates.

(C) HeLa WT and SKO cells overexpressing CAV1-GFP. LDs are induced after transfection by the supply of oleate. LipidTox Deep Red-stained LDs.

(D) Left and right: mean LD area and normalized CAV1-GFP signal around LDs per cell in (C). A two-tailed Mann-Whitney analysis was performed, yielding *p* < 0.0001 for both comparisons. For the LDs' area, *n* = 61 for WT cells and 39 for SKO cells; three replicates. For CAV1-GFP signal around LDs' quantification, *n* = 30 for WT and 29 for SKO cells; three experiments. Bottom: percentage of CAV1-GFP PM signal per swollen cell. Two-tailed Mann-Whitney analysis. *p* = 0.0004, *n* = 14 WT and 21 SKO cells; three replicates.

(E) Immunofluorescence against the N-terminal part of CAV1 in oleate-loaded HeLa WT and SKO cells.

(F) Top: percentage of endogenous CAV1 PM signal per cell in (E). Two-tailed Mann-Whitney analysis. *p* = 0.0338, *n* = 13 for WT and SKO; two replicates. Bottom: endogenous CAV1 recruitment score on LDs in (E). Two-tailed Mann-Whitney analysis. *p* < 0.0001, *n* = 16 WT and 23 SKO cells; two replicates.

(G) Fibroblasts from control and CGL2 patients overexpressing CAV1-GFP. LDs are induced after transfection by the supply of oleic acid.

(H) Normalized CAV1-GFP signal around LDs per cell for experiments in G. *P* = 0.0009, *n* = 15 cells for each cell line; three experiments. Scale bars, 5 µm.

healthy donors and patients with seipin deficiency. In fibroblasts from healthy donors, both transfected and endogenous CAV1 localized perfectly to the PM and showed minimal association with LDs (Figures 1G, 1H, and S2D). In contrast, fibroblasts from two different patients harboring distinct nonsense mutations in seipin displayed reduced CAV1 localization at the PM and pronounced recruitment of CAV1 to the LD surface (Figures 1G, 1H, and S2D). Altogether, our findings indicate that seipin regulates CAV1 trafficking and its localization to organelles.

We hypothesized that the decreased levels of CAV1 at the PM would impair caveolae formation. Since the trafficking of CAV1-containing 70S complexes from the Golgi to the PM to form caveolae depends on cholesterol,⁴⁴ we expressed the D4H cholesterol probe to assess potential cholesterol alterations. We observed significantly reduced recruitment of D4H-mCherry to the PM in SKO cells compared to WT (Figures S2E and S2F),

suggesting impaired cholesterol enrichment at the PM and a consequent reduction in caveolae formation. To further support this conclusion, we used total internal reflection fluorescence (TIRF) microscopy to image the basal PM of WT and SKO cells. SKO cells exhibited significantly fewer CAV1-GFP-positive puncta compared to WT cells (Figures 2A, 2B, and S2G), indicating a reduced number of caveolae and consistent with the overall decrease in CAV1 localization at the PM. Thus, WT cells contained more caveolae, whereas SKO cells showed increased accumulation of CAV1-containing vesicles in the cytosol (Figures 1A and 1B).

Considering the role of CAV1 in buffering PM surface tension to withstand mechanical stress,⁴⁵ we subjected both WT and SKO cells to the same hypotonic medium for 10 min. This treatment caused cell swelling and PM stretching (Figures 2C and S2I). We observed that WT cells were, on average, significantly larger than SKO cells, which can be attributed to several
<figure><img src="image_8.png" /><figcaption>**Figure 2.** Seipin depletion disrupts caveola formation</figcaption></figure>

(A) Left: schematic representation of TIRF microscopy applied to visualize basal PM caveolae in HeLa cells overexpressing CAV1-GFP. Right: TIRF images of WT and SKO cells overexpressing CAV1-GFP.

(B) Number of CAV1-GFP puncta per cell in (A) corresponding to caveolae. *p* = 0.0067, *n* = 14 WT and 11 SKO cells; one experiment.

(C) Left: HeLa WT and SKO cells are submitted to a hypotonic medium with Bodipy 493/503 added. A negative color mode is used. Right: quantification of the swollen WT and SKO cell area from a median plane image. *p* < 0.0001, *n* = 120 for WT and SKO cells; three experiments.

(D) Left: example of a swollen HeLa cell manipulated with a micropipette. Right: quantification of the apparent area expansion modulus and lysis tension of swollen WT and SKO cells. Two-tailed Mann-Whitney analysis. Left: *p* = 0.061, *n* = 13 for WT and SKO cells, Right: *p* = 0.0303, *n* = 7 WT cells and 5 SKO cells; three experiments.

(E) Transmission electron microscopy of cells from control and iATSKO mice. Yellow arrowheads depict caveolae vesicles. Scale bars, 100 nm.

(F and G) Quantification of mean caveolae density per cell and mean internalized caveolae density, respectively. For caveolae density quantifications, a two-tailed Mann-Whitney analysis is done with respective *p* values of 0.0002 and <0.0001; *n* = 16 for CTRL and iATSKO. Scale bars, 5 µm.

signaling factors but may also be correlated with the loss of caveolar membrane reservoirs upon seipin deletion (Figures 2C and S2H). Additionally, the PM of WT cells exhibited altered mechanical properties (Figures 2D and S2I), including a lower apparent area expansion modulus and a higher lysis tension. These results indicate that seipin dysfunction impairs caveola and PM membrane mechanics.

Lastly, we sought to determine whether our findings extend to a more physiological context and at the organismal level, specifically in adipocytes. To this end, we analyzed AT from mice with an inducible adipocyte-specific deletion of seipin following tamoxifen administration (iATSKO). Electron microscopy (EM)

analysis of adipocytes from these mice revealed a pronounced reduction in the number and density of caveolae, along with an increase in intracellular vesicles (Figures 2E–2G, and S2J). These observations align with our cellular data, which showed decreased CAV1 levels at the PM and its accumulation in internal vesicles. Together, these results reinforce our conclusion that seipin deficiency disrupts CAV1 trafficking and impairs caveolae formation.

## Seipin loss is associated with altered Golgi function

CAV1 exits the ER in preassembled 8S complexes, which then relocate to the Golgi apparatus. Once in the TGN, these
structures undergo further assembly into higher-order 70S oligomers, which are subsequently transported to the PM to fulfill their functional roles at the cell surface.¹⁷,⁴⁶ Disruption at any step of this pathway could potentially result in CAV1 mistargeting, leading to its mislocalization to LDs rather than the PM.

To test this, we employed the retention using selective hooks (RUSH) assay⁴⁷ with a glycosylphosphatidylinositol (GPI)-anchored protein to assess trafficking efficiency. We used the GPI protein because CAV1 is palmitoylated, and we hypothesized that it would follow similar trafficking pathways. Early transport of the GPI protein from the ER to the Golgi apparatus, triggered by biotin addition, was comparable between WT and SKO cells, indicating that initial secretory trafficking was unaffected (Figures 3A and 3B). However, after 1 h, in WT cells, the GPI protein was nearly undetectable in the Golgi, having reached the PM. In contrast, in SKO cells, it remained in the Golgi. It gradually reached the PM only after 24 h, achieving similar levels as in the WT condition (Figure S3A). These results indicate a block or delay in transport from the TGN to the PM, suggesting that SKO cells have a specific defect in the late secretory pathway. This defect likely accounts for the mistargeting of CAV1, implying that multiple cargo proteins may be rerouted from the TGN to alternative intracellular destinations.

To further investigate the potential causes of impaired trafficking, we assessed the structural integrity of the Golgi apparatus by staining for GM130, a cis-Golgi marker, or Golgin97, a trans-Golgi marker. In SKO cells, the Golgi appeared rounder compared with WT cells (Figures 3C, 3D, S3B, and S3C). Similar changes in Golgi morphology were observed in fibroblasts from patients compared to healthy donor cells, but only when they were supplied with OA (Figures S3D–S3F). These findings suggest that the Golgi organization is compromised in SKO cells and CGL2 patient fibroblasts, potentially affecting protein trafficking.

The TGN's lateral organization plays a critical role in cargo sorting, to the PM for example, or endosomes and lysosomes.⁴⁸,⁴⁹ The coordination of this process relies in part on lipid-ordered domains enriched in cholesterol and sphingomyelin (SM), essential components of specialized TGN subdomains that drive efficient cargo sorting.⁴⁴ Cholesterol in these regions is delivered by the oxysterol-binding protein (OSBP), which transfer it from the ER to the TGN in exchange for phosphatidylinositol 4-phosphate (PI4P).⁵⁰ This lipid trade occurs at ER-TGN contact sites. SM is synthesized in the TGN by sphingomyelin synthase 1, which uses ceramides delivered from the ER by the ceramide transfer protein (CERT).⁵¹ We analyzed the impact of seipin on these lipid transport proteins.

In SKO cells, overexpressed OSBP exhibited a “stuck” phenotype in the perinuclear Golgi region, contrasting with its soluble and dynamic localization in WT cells (Figures S3G and S3H). This phenotype mimicked the effect observed in WT cells treated with 25-hydroxycholesterol (Figures S3I and S3J), an OSBP inhibitor, suggesting a defect in OSBP-mediated lipid transport. Such a defect is often concomitant with impaired PI4P turnover at the TGN.⁵² Consistent with this, cells transfected with the PI4P reporter P4M showed stronger perinuclear recruitment in SKO compared with WT cells, indicative of PI4P accumulation at the TGN (Figures S3K and S3L). Similarly, overexpressed CERT dis-

played a focused perinuclear localization in WT cells, which was lost in SKO cells, where CERT exhibited a more dispersed cytoplasmic distribution (Figures S3M and S3N). In parallel, the SM-binding toxin lysenin also revealed a scattered SM signal in SKO cells, with reduced concentration in the perinuclear region compared to WT (Figures S3O and S3P).

The above findings suggest that the disrupted TGN organization observed in SKO cells is concomitant and may stem from impaired levels of cholesterol and SM in this compartment. To test this hypothesis, we employed the solvatochromic Prodan-based PK dye, a sensitive reporter of membrane order whose fluorescence emission shifts depending on the lipid environment.⁵³ Following dye supplementation, we analyzed the perinuclear region. We observed, through ratiometric imaging,⁵⁴ a marked decrease in membrane order in SKO cells compared to WT cells (Figures 3E and 3F), consistent with an altered lipid composition. To further validate this observation, we performed lipidomic analysis on Golgi-enriched fractions isolated by ultracentrifugation using Golgin-97 as a marker. This analysis revealed a reduction in both SM and cholesterol levels in SKO cells (Figures S3Q and S3R), consistent with the dispersed lysenin staining pattern observed previously (Figures S3O and S3P) and the altered D4H signal in SKO cells (Figures S2E and S2F).

Finally, to determine whether altered TGN membrane order alone could drive CAV1 mislocalization and recapitulate the phenotype observed in SKO cells, we pharmacologically inhibited the involved lipid transfer proteins in WT cells. OSW-1, a potent OSBP inhibitor, proved toxic to most cells; however, in the surviving population, CAV1 levels at the PM were reduced, but CAV1 did not localize to LDs. We next used 25-hydroxycholesterol (25-HC), a broader OSBP inhibitor, which induced a mild decrease in CAV1 PM signal (Figures 3G, 3H, and S4A). In these cells, CAV1 accumulated more prominently near LDs, yet without a clear association with the LD surface. In contrast, inhibition of CERT using HPA-12 in WT cells effectively phenocopied the SKO condition: CAV1 levels decreased at the PM, and the protein prominently localized to the LD surface (Figures 3I, 3J, and S4B). This impact of CERT inactivity was further confirmed by siRNA-mediated knockdown of CERT (Figures S4C and S4D), reinforcing the conclusion that reduced CERT activity is sufficient to fully mimic the SKO-associated mislocalization of CAV1.

## Excess ceramides cause CAV1 to shift its localization from the PM to LDs

Our data support a model in which proper lipid order at the TGN membrane is crucial for the correct trafficking of CAV1 to the PM. Disruptions in cholesterol and ceramide (or SM) levels impair this process, leading to CAV1 accumulation in cytosolic vesicles and its relocalization to LDs, a phenotype particularly sensitive to sphingolipid imbalance (Figures 3G–3J).

To investigate whether, and which, lipids are altered upon seipin loss, we analyzed the lipid composition of AT from control (CTRL) and iATSKO mice. This model enables a temporally controlled and adipocyte-specific depletion of seipin. This inducible system minimizes confounding effects from potential compensation or systemic adaptation in KO conditions, allowing a more direct assessment of seipin's role in lipid homeostasis.
<figure><img src="image_9.png" /><figcaption>Figure 3. Seipin depletion perturbs the lipid composition and function of the TGN</figcaption></figure>

(A) HeLa WT and SKO cells overexpressing eGFP-GPI-RUSH are imaged before and at 20 and 60 min after biotin addition.

(B) Top: the mean signal ratio between the Golgi and the ER at 20 min. *p* = 0.7278; three experiments (8 cells for each). Bottom: the mean signal ratio between the PM and the Golgi at 60 min. *p* < 0.0001, *n* = 24 cells for each; three replicates.

(C) Immunofluorescence of WT and SKO HeLa cells against Golgi resident protein GM130 and labeled with Hoechst.

(D) Golgi aspect ratio measuring the ratio between the minimum and maximum Feret diameters of the largest Golgi particle. *p* = 0.0136, *n* = 15 each; three experiments.

(E) Ratiometric images of PK dye incubated with HeLa WT and SKO cells.

(F) Relative intensity (%) associated with the corresponding ratio value from experiments in I; *n* = 24 for WT and 37 for SKO cells; three experiments.

(G) HeLa WT overexpressing CAV1-GFP and oleate-loaded. A supply of 2.5 µM 25-hydroxycholesterol (25-HC) is provided during the transfection and maintained afterward. Loading control is done with ethanol (EtOH).

(H) Left: mean normalized CAV1-GFP signal around LDs per cell. Two-tailed Mann-Whitney analysis is done, *p* = 0.1303, *n* = 10 cells for WT + EtOH from two replicates and 15 cells for WT + 25-HC from three replicates. Right: percentage of CAV1-GFP PM signal per swollen cell. *p* = 0.00161, *n* = 13 cells for WT EtOH and 18 cells for WT + 25-HC; two replicates.

(I) HeLa WT cells overexpressing CAV1-GFP and with LDs induced. A loading of 2.5 µM HPA-12 is done during the transfection and maintained afterward. Control is done with DMSO.

(J) Left: mean normalized CAV1-GFP signal around LDs per cell. Two-tailed Mann-Whitney analysis, *p* = 0.0001, *n* = 21 cells for each condition from three replicates. Right: percentage of CAV1-GFP PM signal per swollen cell. *p* = 0.00403, *n* = 17 WT + DMSO and 15 WT + HPA-12 cells; three experiments. Scale bars, 5 µm.
<figure><img src="image_10.png" /><figcaption>Figure 4. Excess ceramides shift CAV1 localization from PM to LDs</figcaption></figure>

(A) Whole tissue lipidomics of CTRL and iATSKO cells, 30 days after induction of the KO. PE, PC, DAG, SM, ceramide (Cer), and TAG have *p* values of 0.0201, 0.2592, <0.0001, 0.8836, 0.1088, and 0.0046, respectively; *n* = 7 for CTRL and 8 for iATSKO.

(B) Lipid fold change of iATSKO versus Control at 14 and 30 days after KO induction.

(C) Oleate-loaded HeLa WT cells overexpressing CAV1-GFP and supplemented with 50 µM of Cer).

(D) Top: mean normalized CAV1-GFP signal around LDs per cell. *p* < 0.0001, *n* = 30 for WT and 15 for WT + Cer. Cells from three experiments. Bottom: percentage of CAV1-GFP PM signal per swollen cell. *p* = 0.0036, *n* = 15 for WT cells from three replicates and 5 cells for WT + Cer.

(E) Immunofluorescence of HeLa WT cells against the N-terminal part of CAV1 after incubation with oleic acid for 24 h and 50 µM of ceramides.

(F) Oleate-loaded HeLa SKO cells overexpressing CAV1-GFP are incubated with 2.5 µM Myriocin (indicated by Myr in the figure) during the transfection and maintained afterward. A control is done with DMSO.

(G) Top: mean normalized CAV1-GFP signal around LDs per cell. *p* = 0.0019, *n* = 15 each from three replicates. Bottom: percentage of CAV1-GFP PM signal per swollen cell. Two-tailed Mann-Whitney analysis. *p* = 0.1804, *n* = 15 for SKO + DMSO and 11 for SKO + Myr cells from two replicates.

(H) Transcriptomics analysis of AdipoQ and C/EBP homologus protein (CHOP) expression of inguinal iATSKO and control cells in the presence or absence of myriocin. A two-tailed Mann-Whitney analysis is performed with respective *p* values of 0.9654, 0.4967, <0.0001, 0.4082, 0.7197, and 0.0006. *n* for AdipoQ experiments is 10 for CTRL VH, 9 for CTRL Myr, 9 for CRE VH, and 10 for CRE Myr cells; *n* for CHOP experiments is 10 for CTRL VH, 8 for CTRL Myr, 9 for CRE VH, and 10 for CRE Myr. Scale bars, 5 µm.
<figure><img src="image_11.png" /><figcaption>**Figure 5.** The glycerolipid-ceramide branchpoint regulates protein traffic</figcaption></figure>

(A) Schematic representation of palmitic acid metabolic pathways and the associated enzymes and metabolites.

(B) Top: comparison of total saturated lipid content from ER fraction lipidomics analysis between WT and SKO. *p* = 0.0382, *n* = 3 experiments. Bottom: normalized PC 16:0/16:0 count from ER fraction. *p* = 0.0462; three experiments.

(C) Oleate-loaded HeLa WT cells overexpressing CAV1-GFP. 1 µM of an SCD1 inhibitor (SCD1i) is added 5 h before transfection and maintained afterward. A control is done with DMSO.

(D) Top: mean normalized CAV1-GFP signal around LDs per cell. A two-tailed Mann-Whitney analysis is performed: *p* < 0.0001, *n* = 21 WT + DMSO and 15 WT + SCD1i cells from three replicates. Bottom: percentage of CAV1-GFP PM signal per swollen cell. *p* = 0.0002, *n* = 17 WT + DMSO and 12 WT + SCD1i cells from 3 to two replicates.

(E) Immunofluorescence against the N-terminal part of CAV1 in HeLa WT. 1 µM of SCD1i is added 5 h before inducing LDs.

(F) Oleate-loaded HeLa SKO cells overexpressing CAV1-GFP and SCD1.

(G) Top: mean normalized CAV1-GFP signal around LDs per cell, from experiments in F. Two-tailed Mann-Whitney analysis, *p* < 0.0001. *n* = 29 for SKO and 15 for SKO + SCD1 cells, from three replicates. Bottom: percentage of CAV1-GFP PM signal per swollen cell from experiments shown in (F). Two-tailed Mann-Whitney analysis. *p* = 0.0088. *n* = 21 for KO and 6 for KO + SCD1 from one experiment.

(legend continued on next page)
Western blot analysis revealed that the absence of seipin in SKO cells did not alter SCD1 levels compared with those in WT cells (Figure S6E), suggesting that seipin might locally coordinate SCD1 function.

To test our hypothesis that seipin deficiency alters local SCD1 activity, we reasoned that restoring SCD1 levels should rescue SKO phenotypes. To this end, we overexpressed the mouse version of SCD1 in SKO cells. Remarkably, SCD1 over-expression was sufficient to restore CAV1 localization to the PM and significantly reduced its mislocalization to LDs (Figures 5F, 5G, and S6F). Under these conditions, subsequent lipidomic analyses revealed a significant decrease in ceramide levels (Figure S6G). SM levels increased, although the changes were not statistically significant (Figure S6G). These findings strongly suggest that in SKO cells, impaired routing of palmitate toward glycerolipid synthesis leads to its accumulation as ceramides, which in turn drives the observed phenotypes. Put differently, elevated palmitate levels promote ceramide overproduction and disrupt CAV1 dynamics.

To further investigate the mechanism, we supplemented WT cells with palmitic acid and OA, or OA alone. Co-supplementation with palmitate and OA was sufficient to mislocalize CAV1 from the PM to LDs (Figures 5H, 5I, and S6H), whereas OA alone had no such effect. Importantly, blocking ceramide synthesis using myriocin in palmitate-treated WT cells prevented CAV1 accumulation on LDs, but did not restore its localization to the PM. Instead, CAV1 accumulated in vesicular structures adjacent to LDs, a phenotype also observed in myriocin-treated SKO cells (Figures 5J, 5K, and S6H; Figure 4F). These results indicate that the conversion of palmitate into ceramides was responsible for the altered trafficking of CAV1. Finally, because SKO cells displayed elevated palmitate levels, we pharmacologically inhibited fatty acid synthase (FASN), the key enzyme in de novo lipogenesis (Figure 5A). This treatment restored CAV1 localization at the PM and abolished its LD association (Figures 5L, 5M, S6L, and S6J). Altogether, these findings demonstrate that routing palmitate toward glycerolipid synthesis rather than ceramide production in the ER is essential for proper CAV1 trafficking and cellular homeostasis, and this process is under the control of seipin.

DISCUSSION

Seipin plays a critical role in LD nucleation and growth, yet evidence suggests its functions extend beyond LD dynamics to

broader aspects of lipid metabolism.⁶²⁻⁶⁴ While LDs can form independently of protein involvement,⁶⁵ seipin ensures the coordinated formation and proper functionality of these organelles. In the absence of seipin, adipogenesis is impaired, and the removal of seipin results in the loss of adipose tissue despite the potential presence of LDs. This contrasts with mice lacking the diacylglycerol O-acyltransferase (DGAT) triglyceride synthesis enzymes and LDs; yet they are still capable of developing adipose tissue.⁶⁶ These findings decouple the roles of seipin in the physical control of LD formation, lipid metabolic pathways, and adipogenesis, emphasizing the complexity and multifaceted nature of its functions.

We found that seipin deficiency leads to the accumulation of palmitate, resulting in elevated ceramide levels. This observation aligns with a recent study in yeast, which demonstrates that seipin negatively regulates sphingolipid metabolism.⁶³ Our findings reveal that ceramide accumulation impairs the TGN membrane order, likely impairing its ability to efficiently sort and transport cargo such as CAV1 to the PM, thereby reducing caveolae formation. Inactivating serine SPT to block ceramide synthesis prevented CAV1 localization to LDs. Still, it did not restore its proper relocalization to the PM, indicating that well-tuned ceramide levels, neither in excess nor in shortage, are critical for appropriate TGN function and cargo trafficking. Conversely, overexpression of SCD1 or inhibition of FASN rescued both the LD and PM localization defects of CAV1, highlighting the critical role of saturated fatty acid accumulation in these phenotypes, as supported by our lipidomics data. Furthermore, supplementation of WT cells with palmitate or ceramide reproduced the phenotypes associated with seipin deficiency, confirming their causative roles in the observed defects. Overall, our results underscore the essential role of seipin in regulating protein trafficking, particularly that of CAV1. Seipin operates at the intersection of ceramide and glycerolipid biosynthesis, modulating the allocation of synthesized palmitate between these lipid pathways. This regulatory function is crucial for maintaining cellular lipid homeostasis and ensuring proper protein trafficking (Figure 5A).

Seipin assembles into an oligomeric, donut-shaped structure comprising 11–12 monomers,³³⁻³⁵ providing multiple potential interaction sites to coordinate local metabolic reactions. Notably, seipin interacts with several key enzymes involved in the synthesis of ceramides and glycerolipids, including SPT, SCD1, glycerol-3-phosphate acyltransferases, DGAT2, 1-acylglycerol-3-phosphate O-acyltransferase 2 (AGPAT2), and Lipin1.⁶¹⁻⁶³,⁶⁷,⁶⁸ These

(H) HeLa WT cells overexpressing CAV1-GFP. After transfection, a palmitate-oleate mixture is added for 24 h

(I) Top: mean normalized CAV1-GFP signal around LDs per cell. Quantification when oleic acid was incubated alone or mixed with palmitate. *p* < 0.0001, *n* = 29 for WT and 9 for WT + Palm from 3 to 2 replicates. Bottom: percentage of CAV1-GFP PM signal per swollen cell. Quantification when oleic acid was incubated alone or mixed with palmitate. Two-tailed Mann-Whitney analysis: *p* = 0.00228, *n* = 14 for WT and 9 for WT + Palm cells from one experiment.

(J) HeLa WT cells overexpressing CAV1-GFP were incubated with 2.5 µM Myr. After transfection, a palmitate-oleate mixture is added for 24 h

(K) Quantification of WT cells loaded with a palmitate-oleate mix with or without Myr. Left: mean normalized CAV1-GFP signal around LDs per cell, *p* = 0.0303, *n* = 9 for WT – Myr and 5 for WT + Myr cells. Right: percentage of CAV1-GFP PM signal per swollen cell. *p* = 0.8693, *n* = 9 for WT – Myr and 8 for WT + Myr cells from one experiment.

(L) Oleate-loaded HeLa WT cells overexpressing CAV1-GFP. 1 µM of an FASN inhibitor (FASNi) is added 5 h before transfection and maintained afterward. Loading control is with DMSO.

(M) Top: mean normalized CAV1-GFP signal around LDs per cell. A two-tailed Mann-Whitney analysis is done, *p* = 0.0010, *n* = 15 cells each from three experiments. Bottom: percentage of CAV1-GFP PM signal per swollen cell. Two-tailed Mann-Whitney *p* = 0.0012, *n* = 15 cells each from three experiments. Scale bars, 5 µm.
interactions suggest that seipin may play a regulatory role in the activity of these enzymes in response to metabolic signals, coordinating lipid-related biochemical reactions in specific subcellular ER regions.⁶⁹

For example, we recently found that seipin is recruited to the mitochondria-associated membrane (MAM) during lipogenesis, a site where LDs either form or receive lipid.⁷⁰,⁷¹ Interestingly, SCD1, SPT subcomplexes, and DGAT2 are enriched in MAMs,⁶⁷,⁷² highlighting the need to concentrate and regulate their activity there, potentially via a seipin-dependent mechanism.⁶¹ In the absence of seipin, overexpression of SCD1 may compensate by achieving the necessary local concentration and activity of the enzyme at MAMs, thereby restoring normal lipid fluxes and rescuing CAV1 trafficking in SKO cells.

Based on our findings and analysis, we propose that specific seipin complexes localize at MAMs and create a microenvironment that balances the allocation of palmitate between glycerolipid and ceramide pathways. This may be achieved by efficiently channeling lipid substrates and products to enzymes localized near the seipin complex. Such an organization would ensure coordinated and efficient lipid metabolic processes, thereby maintaining proper cellular lipid and membrane homeostasis, critical for adequate protein trafficking.

The influence of seipin on the ceramide/glycerolipid balance could be determined by the rate of palmitate synthesis or uptake, which is governed by the cell's metabolic demands and seipin levels. When ceramide levels reach a critical threshold, cargo transport from the TGN to the PM is altered, particularly for those that rely on membrane order. This disruption can cause the mislocalization of cargos like CAV1 to LDs, potentially serving as a cellular strategy to alleviate stress caused by lipid homeostasis disturbances. This pathway might be activated in specific cell types under particular physiological or pathophysiological conditions and could be constitutively active in the absence of seipin. This pathway may be triggered in specific instances to deliver lipids to LDs.⁴⁶,⁷³

Our findings demonstrate that seipin deficiency results in the mislocalization of CAV1 to the PM and a decrease in caveolae. Similarly, CAV1 and Cavin1 deficiency are responsible for CGL3 and CGL4, respectively, resulting in a reduction of caveolae. Lastly, in newborn mice deficient for AGPAT2, the gene whose mutation causes CGL1, the adipocyte displays an early loss of caveolae that precedes lipodystrophy⁷⁴. Therefore, all CGLs share the loss of caveolae, which may be the key element in the dysfunction of primary adipocytes that characterizes CGLs. Notably, several pieces of evidence produced in CAV1-deficient adipocytes support the crucial role of caveolae in lipid handling and insulin signaling in adipocytes¹⁸.

CAV1 plays a crucial role in maintaining lipid raft domains, which are essential for the proper localization and stabilization of the insulin receptor at the PM in adipocytes. In addition to supporting insulin receptor function, CAV1 regulates downstream signaling pathways, including the secretion of adipokines and free fatty acids, vital for normal adipocyte function and lipid storage. When CAV1 is dysfunctional, it accelerates the degradation of both the insulin receptor and the glucose transporter,²³,²⁴ leading to impaired insulin signaling and systemic insulin resistance. This dysfunction is strongly linked to metabolic disorders,

such as type 2 diabetes and obesity. Given seipin's role in CAV1 PM trafficking, our study uncovers a cell-autonomous mechanism where seipin deficiency causes insulin resistance through the accumulation of ceramides.

Finally, our data suggest that the accumulation of ceramide, palmitate, and stearate, all known predictors of incident type 2 diabetes,⁷⁵ could also be associated with CAV1 mislocalization and, thus, the impaired trafficking of insulin receptor and other transporters. This opens a new avenue for pathophysiological studies of human insulin resistance, possibly leading to innovative pharmacological interventions.

## Limitations of the study

A current limitation of our study is that the precise mechanism by which CAV1 is delivered to LDs remains unclear. To date, the only known trafficking pathway for monotopic membrane proteins to LDs occurs via the ER. One plausible hypothesis is that, in the absence of seipin, CAV1 becomes retained in the ER and subsequently diffuses laterally from the ER to LDs. However, we could not visualize such a phenomenon happening. The delivery mechanism of CAV1 to LDs requires further investigation.

## RESOURCE AVAILABILITY

### Lead contact

Further information and requests for resources and reagents should be directed to and will be fulfilled by the lead contact, Abdou Rachid Thiam (thiam@ens.fr).

### Materials availability

This study did not generate new, unique reagents.

### Data and code availability

* Data: The metabolomics data have been deposited in MetaboLights⁷⁶ repository with the study identifier MTBLS12759.

* Code: This study did not generate any new code.

* Other: Additional information is available from the **lead contact** upon request.

## ACKNOWLEDGMENTS

We thank the Thiam team for the helpful discussions and Dalila Ajjaji for assisting in some experiments. We thank Gilliane Chadeuf for the fractionation experiments. The French National Research Agency, ANR, via ANR-18-CE11-0012-01-MOBIL, ANR-21-CE11-0032-02-LIPRODYN, ANR-22-CE11-0024-01/MADE, and ANR-22-CE11-MAMA supported this work. This work has also benefited from the Imagerie-Gif core facility, supported by ANR-10-INBS-04/FranceBioImaging; ANR-11-IDEX-0003-02/Saclay Plant Sciences. C.V. is a member of the European Reference Network on Rare Endocrine Conditions—Project ID no. 739527. The ANR supported T.H. through the “Investments for the Future” programs LABEX SIGNALIFE ANR-11-LABX-0028, IDEX UCAJedi ANR-15-IDEX-01, and the ATIP-Avenir program (CNRS/Inserm). The authors acknowledge the Université Paris-Saclay, INSERM US23 / CNRS UAR 3655, Imaging Core Facility, Gustave Roussy, 94805, Villejuif, France, for the expertise and advice in using instruments and methodological developments.

## AUTHOR CONTRIBUTIONS

A.R.T. and M.C. designed research and experiments. M.C. conducted all cell experiments and analyses with the assistance of M.O. and M.Z. J.T. performed biophysical measurements. N.E.K. and F.G. assisted with image analysis. K.S. and J.P. helped with TIRF experiments. R.S., M.P., Y.C., and X.P. performed the mouse work and the EM of tissues, with assistance from S.L.L. T.H. helped
with the myriocin and ceramide experiments. C.V. assisted in the fibroblast experiments. A.R.T. wrote the manuscript and reviewed it with the co-authors.

### DECLARATION OF INTERESTS

A.R.T. is an advisor and cofounder of Oria Bioscience.

### STAR★METHODS

Detailed methods are provided in the online version of this paper and include the following:

* **KEY RESOURCES TABLE**

* **EXPERIMENTAL MODEL AND STUDY PARTICIPANT DETAILS**

* Cell culture

* Animal studies

* **METHOD DETAILS**

* Transfection and lipid droplet formation

* Hypotonic medium

* Caveola proteins immunofluorescence staining

* Micromanipulation

* Cells' area quantification

* LDs' area quantification

* Protein-targeting to LD quantification

* CAV1-GFP PM signal quantification

* CAV1 immunofluorescence quantifications

* Blocking metabolism and lipid transport

* Ceramides loading into HeLa cells

* Trans-Golgi-network domains quantification

* Lipidomics, sample preparation, and analysis

* RUSH experiment

* Golgi aspect ratio

* Quantification of OSBP, P4M, Lysenin & CERT

* ER purification from mice extracts

* Myriocin mice treatment

* Lipidomics of mice extracts

* Electron microscopy of caveola

* SCD1 and CAV1-GFP expression analysis

* SCD1 overexpressing cell lipidomics

* CERT knockdown experiments

* TIRF imaging of CAV1-GFP

* Confocal imaging, data representation

* **QUANTIFICATION AND STATISTICAL ANALYSIS**

### SUPPLEMENTAL INFORMATION

Supplemental information can be found online at https://doi.org/10.1016/j.celrep.2025.116320.

Received: December 5, 2024

Revised: June 19, 2025

Accepted: August 29, 2025

Published: September 22, 2025

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40. Song, J., Mizrak, A., Lee, C.W., Cicconet, M., Lai, Z.W., Tang, W.C., Lu, C. H., Mohr, S.E., Farese, R.V., Jr., and Walther, T.C. (2022). Identification of two pathways mediating protein targeting from ER to lipid droplets. *Nat. Cell Biol.* **24**, 1364-1377.

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42. Fei, W., Du, X., and Yang, H. (2011). Seipin, adipogenesis and lipid droplets. *Trends Endocrinol. Metab.* **22**, 204-210.

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46. Matthaeus, C., and Taraska, J.W. (2020). Energy and dynamics of caveolae trafficking. *Front. Cell Dev. Biol.* **8**, 614472.

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52. Mesmin, B., Bigay, J., Polidori, J., Jamecna, D., Lacas-Gervais, S., and Antonny, B. (2017). Sterol transfer, PI 4P consumption, and control of membrane lipid order by endogenous OSBP. *EMBO J.* **36**, 3156-3174.

53. Klymchenko, A.S. (2023). Fluorescent Probes for Lipid Membranes: From the Cell Surface to Organelles. *Acc. Chem. Res.* **56**, 1-12.

54. Danylchuk, D.I., Jouard, P.-H., and Klymchenko, A.S. (2021). Targeted solvatochromic fluorescent probes for imaging lipid order in organelles under oxidative and mechanical stress. *J. Am. Chem. Soc.* **143**, 912-924.

55. Dumesnil, C., Vanharanta, L., Prasanna, X., Omrane, M., Carpentier, M., Bhapkar, A., Enkavi, G., Salo, V.T., Vattulainen, I., Ikonen, E., and Thiam, A.R. (2023). Cholesterol esters form supercooled lipid droplets whose nucleation is facilitated by triacylglycerols. *Nat. Commun.* **14**, 915.

56. Lee, Y.-S., Choi, K.M., Choi, M.H., Ji, S.Y., Lee, S., Sin, D.M., Oh, K.W., Lee, Y.M., Hong, J.T., Yun, Y.P., and Yoo, H.S. (2011). Serine palmitoyltransferase inhibitor myriocin induces growth inhibition of B16F10 melanoma cells through G2/M phase arrest: Myriocin inhibits B16F10 melanoma cell growth. *Cell Prolif.* **44**, 320-329.

57. Rabionet, M., Gorgas, K., and Sandhoff, R. (2014). Ceramide synthesis in the epidermis. *Biochim. Biophys. Acta* **1841**, 422-434.

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59. Chen, L., Ren, J., Yang, L., Li, Y., Fu, J., Li, Y., Tian, Y., Qiu, F., Liu, Z., and Qiu, Y. (2016). Stearoyl-CoA desaturase-1 mediated cell apoptosis in colorectal cancer by promoting ceramide synthesis. *Sci. Rep.* **6**, 19665.

60. Boutet, E., El Mourabit, H., Prot, M., Nemani, M., Khallouf, E., Colard, O., Maurice, M., Durand-Schneider, A.M., Chrétien, Y., Grès, S., et al. (2009).
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61. Lounis, M.A., Lalonde, S., Rial, S.A., Bergeron, K.F., Ralston, J.C., Mutch, D.M., and Mounier, C. (2017). Hepatic BSCL2 (Seipin) Deficiency Disrupts Lipid Droplet Homeostasis and Increases Lipid Metabolism via SCD1 Activity. Lipids 52, 129-150.

62. Pagac, M., Cooper, D.E., Qi, Y., Lukmantara, I.E., Mak, H.Y., Wu, Z., Tian, Y., Liu, Z., Lei, M., Du, X., et al. (2016). SEIPIN regulates lipid droplet expansion and adipocyte development by modulating the activity of glycerol-3-phosphate acyltransferase. Cell Rep. 17, 1546-1559.

63. Su, W.-C., Lin, Y.-H., Pagac, M., and Wang, C.-W. (2019). Seipin negatively regulates sphingolipid production at the ER-LD contact site. J. Cell Biol. 218, 3663-3680.

64. Bi, J., Wang, W., Liu, Z., Huang, X., Jiang, Q., Liu, G., Wang, Y., and Huang, X. (2014). Seipin promotes adipose tissue fat storage through the ER Ca2+-ATPase SERCA. Cell Metab. 19, 861-871.

65. Thiam, A.R., and Ikonen, E. (2020). Lipid Droplet Nucleation. Trends Cell Biol. 31, 108-118.

66. Chitraju, C., Fischer, A.W., Ambaw, Y.A., Wang, K., Yuan, B., Hui, S., Walther, T.C., and Farese, R.V., Jr. (2023). Mice lacking triglyceride synthesis enzymes in adipose tissue are resistant to diet-induced obesity. eLife 12, RP88049.

67. Man, W.C., Miyazaki, M., Chu, K., and Ntambi, J. (2006). Colocalization of SCD1 and DGAT2: implying preference for endogenous monounsaturated fatty acids in triglyceride synthesis. J. Lipid Res. 47, 1928-1939.

68. Talukder, M.M.U., Sim, M.F.M., O'Rahilly, S., Edwardson, J.M., and Rochford, J.J. (2015). Seipin oligomers can interact directly with AGPAT2 and lipin 1, physically scaffolding critical regulators of adipogenesis. Mol. Metab. 4, 199-209.

69. Choudhary, V., El Atab, O., Mizzon, G., Prinz, W.A., and Schneiter, R. (2020). Seipin and Nem1 establish discrete ER subdomains to initiate yeast lipid droplet biogenesis. J. Cell Biol. 219, e201910177.

70. Guyard, V., Monteiro-Cardoso, V.F., Omrane, M., Sauvanet, C., Houcine, A., Boulogne, C., Ben Mbarek, K., Vitale, N., Faklaris, O., El Khallouki, N., et al. (2022). ORP5 and ORP8 orchestrate lipid droplet biogenesis and maintenance at ER-mitochondria contact sites. J. Cell Biol. 221, e202112107.

71. Freyre, C.A.C., Rauher, P.C., Ejsing, C.S., and Klemm, R.W. (2019). MIGA2 links mitochondria, the ER, and lipid droplets and promotes de novo lipo-genesis in adipocytes. Mol. Cell 76, 811-825.e14.

72. Aaltonen, M.J., Alecu, I., König, T., Bennett, S.A., and Shoubridge, E.A. (2022). Serine palmitoyltransferase assembles at ER-mitochondria contact sites. Life Sci. Alliance 5, e202101278.

73. Matthaeus, C., Lahmann, I., Kunz, S., Jonas, W., Melo, A.A., Lehmann, M., Larsson, E., Lundmark, R., Kern, M., Blüher, M., et al. (2020). EHD2-mediated restriction of caveolar dynamics regulates cellular fatty acid uptake. Proc. Natl. Acad. Sci. USA 117, 7471-7481.

74. Cautivo, K.M., Lizama, C.O., Tapia, P.J., Agarwal, A.K., Garg, A., Horton, J.D., and Cortés, V.A. (2016). AGPAT2 is essential for postnatal development and maintenance of white and brown adipose tissue. Mol. Metab. 5, 491-505.

75. Hilvo, M., Salonurmi, T., Havulinnä, A.S., Kauhanen, D., Pedersen, E.R., Tell, G.S., Meyer, K., Teeriniemi, A.M., Laatikainen, T., Jousilahti, P., et al. (2018). Ceramide stearic to palmitic acid ratio predicts incident diabetes. Diabetologia 61, 1424-1434.

76. Yurekten, O., Payne, T., Tejera, N., Amaladoss, F.X., Martin, C., Williams, M., and O'Donovan, C. (2024). MetaboLights: open data repository for metabolomics. Nucleic Acids Res. 52, D640-D646.

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## STAR★METHODS

### KEY RESOURCES TABLE

<table><thead><tr><th>REAGENT or RESOURCE</th><th>SOURCE</th><th>IDENTIFIER</th></tr></thead><tbody><tr><td colspan="3"><strong>Antibodies</strong></td></tr><tr><td>Purified Mouse Anti-GM130</td><td>Bd Biosciences</td><td>Cat# 610822; RRID: AB_398142</td></tr><tr><td>Anti-Golgin-97 Antibody, clone 4K16, ZooMAb® Rabbit Monoclonal</td><td>Sigma-Aldrich</td><td>Cat# ZRB1346</td></tr><tr><td>caveolin-1 Antibody (7C8)</td><td>Santa Cruz Biotechnology</td><td>Cat# sc-53564; RRID:AB_628859</td></tr><tr><td>PTRF Polyclonal Antibody</td><td>Proteintech</td><td>Cat# 18892-1-AP; RRID:AB_10596795</td></tr><tr><td>GAPDH monoclonal antibody</td><td>Invitrogen</td><td>Cat# AM4300; RRID: AB8437392</td></tr><tr><td>Donkey anti-Mouse IgG (H + L) Highly Cross-Adsorbed Secondary Antibody, Alexa Fluor™ 488</td><td>ThermoFisher</td><td>Cat# A-21202; RRID: AB_141607</td></tr><tr><td>Donkey anti-Rabbit IgG (H + L) Cross-Adsorbed Secondary Antibody, HRP</td><td>Invitrogen</td><td>Cat# SA1-200; RRID: AB_325994</td></tr><tr><td>Anti-Calreticulin antibody [FMC 75]</td><td>Abcam</td><td>Cat# ab22683;RRID:AB_447253</td></tr><tr><td>GFP Monoclonal Antibody (GF28R)</td><td>Invitrogen</td><td>Cat# MA5-15256; RRID:AB_10979281</td></tr><tr><td>SCD Polyclonal antibody</td><td>Invitrogen</td><td>Cat# PA5-95762; RRID:AB_2807564</td></tr><tr><td>Beta Actin loading control monoclonal antibody (BA3R)</td><td>Invitrogen</td><td>Cat# MA5-15739; RRID:AB_10979409</td></tr><tr><td>Donkey anti-Mouse IgG (H + L) Cross-Adsorbed Secondary Antibody, HRP</td><td>Invitrogen</td><td>Cat# SA1-100; RRID: AB_228307</td></tr><tr><td>Polyclonal Goat Anti-Mouse immunoglobulins/ Hrp conjugated antibody</td><td>Agilent Technologies</td><td>Cat# P0447; RRID:AB_2617137</td></tr><tr><td>Donkey anti-Rabbit IgG (H + L) Highly Cross-Adsorbed Secondary Antibody, Alexa Fluor™ 488</td><td>Invitrogen</td><td>A-21206; RRID:AB_2535792</td></tr><tr><td colspan="3"><strong>Chemicals, Peptides and Recombinant Proteins</strong></td></tr><tr><td>DMEM, High glucose, GlutaMAX™ Supplement, pyruvate</td><td>Dutscher</td><td>Cat# 31966-021</td></tr><tr><td>DMEM, Low glucose, GlutaMAX™ Supplement, pyruvate</td><td>Gibco</td><td>Cat# 21885-025</td></tr><tr><td>Fetal bovin serum from South America</td><td>Dutscher</td><td>Cat# 500105</td></tr><tr><td>Penicillin-Streptomycin Solution 100X</td><td>Gibco</td><td>Cat# P4333</td></tr><tr><td>Trypsin-EDTA (0.5%), no phenol red</td><td>Gibco</td><td>Cat# 15400-054</td></tr><tr><td>JetPEI transfection reagent</td><td>PolyPlus</td><td>Cat# #101-10 N</td></tr><tr><td>Bovine serum albumin</td><td>Sigma-Aldrich</td><td>Cat# A1595</td></tr><tr><td>Oleic acid</td><td>Sigma Aldrich</td><td>Cat# O1008-1G</td></tr><tr><td>Palmitic acid</td><td>Sigma Aldrich</td><td>Cat# P0500</td></tr><tr><td>Methyl-β-cyclodextrin</td><td>Sigma-Aldrich</td><td>Cat# C4555</td></tr><tr><td>Dimethyl sulfoxide</td><td>Sigma Aldrich</td><td>Cat# 472301</td></tr><tr><td>Cholesterol</td><td>Sigma-Aldrich</td><td>Cat# C8667</td></tr><tr><td>HCS LipidTox™ Deep Red Neutral Lipid Stain</td><td>Thermofisher</td><td>Cat# H34477</td></tr><tr><td>Formaldehyde</td><td>Sigma-Aldrich</td><td>Cat# P-6148</td></tr><tr><td>Saponin</td><td>Fisher Scientific</td><td>Cat# 10294440</td></tr><tr><td>Cold fish gelatin</td><td>Sigma-Aldrich</td><td>Cat# G7041</td></tr><tr><td>ProLong™ Gold Antifade Mountant</td><td>Invitrogen</td><td>Cat# P36934</td></tr><tr><td>ProLong™ Diamond Antifade Mountant</td><td>Invitrogen</td><td>Cat# P36961</td></tr><tr><td>Bodipy 493/503</td><td>Invitrogen</td><td>Cat# D3922</td></tr></tbody></table>

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Continued

<table><thead><tr><th>REAGENT or RESOURCE</th><th>SOURCE</th><th>IDENTIFIER</th></tr></thead><tbody><tr><td>mPEG5K-silane</td><td>Merck</td><td>Cat# JKA3037</td></tr><tr><td>PF-04620110</td><td>Sigma-Aldrich</td><td>Cat# PZ0207</td></tr><tr><td>PF-06424439</td><td>Sigma-Aldrich</td><td>Cat# P20233</td></tr><tr><td>Myriocin</td><td>Sigma-Aldrich</td><td>Cat# M1177</td></tr><tr><td>HPA-12</td><td>Sigma-Aldrich</td><td>Cat# SML2894</td></tr><tr><td>25-Hydroxycholesterol</td><td>Sigma-Aldrich</td><td>Cat# 700019P</td></tr><tr><td>MF-438</td><td>Sigma-Aldrich</td><td>569406</td></tr><tr><td>GSK837149A</td><td>Sigma-Aldrich</td><td>G5547</td></tr><tr><td>C18 Ceramide (d18:1/18:0)</td><td>Sigma-Aldrich</td><td>860518P</td></tr><tr><td>C12 NBD Ceramide (d18:1/12:0)</td><td>Cayman Chemical</td><td>10007958</td></tr><tr><td>PK-Dye</td><td>Valanciunaite et al.<sup>77</sup></td><td>N/A</td></tr><tr><td>Hoechst 33342</td><td>ThermoFisher</td><td>Cat# 62249</td></tr><tr><td>Opti-MEM<sup>TM</sup> I Reduced Serum Medium</td><td>Gibco</td><td>Cat# 31985-062</td></tr><tr><td>Lipofectamine<sup>TM</sup> RNAiMAX Transfection Reagent</td><td>Invitrogen</td><td>Cat #13778150</td></tr><tr><td>DPBS</td><td>Gibco</td><td>Cat# 14200-059</td></tr><tr><td>Complete<sup>TM</sup> protease inhibitors</td><td>Roche</td><td>Cat# 04693116001</td></tr><tr><td>Nupage<sup>TM</sup> LDS Sample Buffer</td><td>Invitrogen</td><td>Cat# NP0007</td></tr><tr><td>Nupage<sup>TM</sup> 4-12% Bis-Tris gels</td><td>Invitrogen</td><td>Cat# NP0321BOX</td></tr><tr><td>NuPAGE<sup>TM</sup> MES SDS Running Buffer (20X)</td><td>Invitrogen</td><td>Cat# H34477</td></tr><tr><td>NuPAGE<sup>TM</sup> MOPS SDS Running Buffer (20X)</td><td>Invitrogen</td><td>Cat# NP0001</td></tr><tr><td>NuPAGE<sup>TM</sup> Transfer Buffer (20X)</td><td>Invitrogen</td><td>Cat# NP00061</td></tr><tr><td>D-Biotin</td><td>Invitrogen</td><td>Cat# B1585</td></tr><tr><td>Tween 20</td><td>Calbiochem</td><td>Cat# 9005-64-5</td></tr></tbody></table>

Critical commercial assays

<table><tr><td>35 mm Dish | No. 1.5 Coverslip | 14 mm Glass Diameter | Uncoated</td><td>MatTek Corp., Ashland, MA</td><td>Cat# P35G-1.5-14-C</td></tr><tr><td>Pierce Western electrochemiluminescence kit</td><td>Thermo scientific</td><td>Cat# 32106</td></tr><tr><td>Pierce BCA Protein Assay Kit</td><td>Thermo Scientific</td><td>Cat# 23225</td></tr><tr><td>Whaman PROTRAN Nitrocellulose Transfer Membrane BA85</td><td>Whatman</td><td>Cat# 10401196</td></tr><tr><td>Glass cover slips</td><td>Epredia</td><td>Cat# CB00180RA120MNZ0</td></tr><tr><td>VWR Microscope Slides</td><td>Avantor Science Central</td><td>Cat# 631-1550</td></tr><tr><td>Glass capillaries</td><td>Harvard Apparatus</td><td>W3 30-0017</td></tr></table>

Cell lines

<table><tr><td>HeLa WT and SKO cells</td><td>Gift from Robert Yang's lab</td><td>N/A</td></tr><tr><td>Fibroblasts: CTRL, BSCL2 p.Arg138* and p.Tyr289Leufs*64 homozygous mutations</td><td>Magré et al.<sup>32</sup></td><td>N/A</td></tr></table>

Experimental models: Organisms/strains

<table><tr><td>C57BL6J Bscl2lox/lox</td><td>European Mouse Mutant Cell Repository</td><td>http://www.phenomin.fr</td></tr><tr><td>C57BL6J ERT2-Adipoq-CRE</td><td>Pr S Offermanns lab (Max Plank institute) Sassmann et al.<sup>78</sup></td><td>N/A</td></tr></table>

Oligonucleotides

<table><tr><td>siLUC</td><td>Gift from Kristine schauer's lab</td><td>N/A</td></tr><tr><td>CGUACGCGGAAUACUUCGA(dT)(dT)</td><td></td><td></td></tr><tr><td>UCGAAGUAUUCCGCGUACG(dT)(dT)</td><td></td><td></td></tr></table>

(Continued on next page)
Continued

<table><thead><tr><th>REAGENT or RESOURCE</th><th>SOURCE</th><th>IDENTIFIER</th></tr></thead><tbody><tr><td>ON-TARGETplus SMARTPool siRNA COL4A3BP</td><td>Dharmacon</td><td>Cat# L-012101-00-0005</td></tr><tr><td>GAACAGAGGAAGCAUAUAA</td><td></td><td></td></tr><tr><td>GAUGGUGACUUCUUGCAUA</td><td></td><td></td></tr><tr><td>GUGGAAACAUUAGCUGAUA</td><td></td><td></td></tr><tr><td>GAACAGCACAAGACUGAAU</td><td></td><td></td></tr></tbody></table>

Recombinant DNA

<table><tbody><tr><td>PCMV3-CAV1-GFP-spark</td><td>Sinobiological</td><td>(#HG11440-ACG)</td></tr><tr><td>CAV1-mCherry</td><td>Hayer et al.<sup>79</sup></td><td>Addgene plasmid # 27705</td></tr><tr><td>EGFP-caveolin-3</td><td>Luetterforst et al.<sup>80</sup></td><td>Addgene plasmid # 68396</td></tr><tr><td>cavin-1-mEGFP</td><td>Hayer et al.<sup>79</sup></td><td>Addgene plasmid # 27709</td></tr><tr><td>pEGFPN1-Cav2h</td><td>Gift from Albert Pol's lab</td><td>N/A</td></tr><tr><td>GFP-CAV1</td><td>Gift from Albert Pol's lab, Spain</td><td>N/A</td></tr><tr><td>pSH-EFIRES-P-Seipin-miniAA7-mScarlet-I</td><td>Li et al.<sup>81</sup></td><td>Addgene plasmid # 129721</td></tr><tr><td>Str-KDEL_SBP-EGFP-GPI</td><td>Boncompain et al.<sup>47</sup></td><td>Addgene plasmid # 65294</td></tr><tr><td>OSBP-mCherry</td><td>Gift from Bruno Mesmin's lab</td><td>N/A</td></tr></tbody></table>

Software and algorithms

<table><tbody><tr><td>ImageJ</td><td>Schneider et al.<sup>7</sup></td><td>https://imagej.nih.gov/ij/</td></tr><tr><td>GraphPad Prism</td><td>GraphPad Software, Inc.</td><td>www.graphpad.com/</td></tr><tr><td>Adobe Illustrator</td><td>Adobe Creative Cloud</td><td>https://www.adobe.com/creativecloud.html</td></tr><tr><td>Affinity Designer 2.0</td><td>Serif Europe</td><td>https://affinity.serif.com</td></tr><tr><td>Stardist</td><td>Von Chamier et al. 2021</td><td>https://github.com/stardist/stardist</td></tr><tr><td>CellProfiler 4.0</td><td>Stirling et al.<sup>82</sup>; McQuin et al.<sup>83</sup>; Kamentsky et al.<sup>84</sup>; Carpenter et al.<sup>85</sup></td><td>https://cellprofiler.org/</td></tr><tr><td>Python 3.7.3</td><td>Python Software Foundation</td><td>https://www.python.org/</td></tr><tr><td>MetaboLights</td><td>EMBL-EBI</td><td>https://www.ebi.ac.uk/metabolights/Study identifier: MTBLS12759</td></tr></tbody></table>

EXPERIMENTAL MODEL AND STUDY PARTICIPANT DETAILS

Cell culture

Fibroblasts used in this study, obtained from the Inserm U938 cell repository (declared collection # DC-2008-529), originated from patients harboring *BSCL2* p.Arg138* or p. Tyr289Leufs*64 homozygous mutations (Magré J et al., Nat Genet, 2001).32 HeLa WT and SKO cells were obtained from Robert Yang. Cells were maintained DMEM, High glucose, (4.5 g. L-1), pyruvate supplemented with sodium pyruvate, fetal bovine serum (FBS) (10%), and penicillin-streptomycin (1%). Fibroblast cells were cultivated in DMEM, Low glucose (1.0 g. L-1) with FBS (10%) and penicillin-streptomycin (1%). All cells were incubated at 37°C in a humidified atmosphere containing 5% CO₂. A mycoplasma detection test was realized every month.

Animal studies

Bsc12lox/lox was established at the Institut Clinique de la Souris (http://www.phenomin.fr). Three clones were ordered from the European Mouse Mutant Cell Repository (https://www.eummcr.org/). All three clones were validated by Southern blot (internal Neo probe) and PCR confirmation of the presence of the 3' *LoxP* site. Clone HEPD0757_2_A10 was karyotyped by chromosome spreading and Giemsa staining and microinjected in BALB/cN blastocysts. Resulting male chimeras were obtained and germ line transmission was achieved for the tm1a allele (knock-out first with conditional potential). The KO allele (tm1b) was obtained after breeding tm1a animals with a Cre deleter line (Birling et al., Genesis, 2012)86 and phenotyped by PHENOMIN-ICS. All data are freely available through the IMPC web site (https://wwwjacm.org/data/genes/MGI:1298392). The cKO Bsc12 allele (tm1c; Lox/Lox) was obtained by breeding the tm1a allele with a Flp deleter line (Birling et al. Genesis, 2012).86 Bsc12lox/lox mice were crossed with ERT2-Adipoq-CRE (Sassmann et al., Genesis, 2010)78 mice to generate Bsc12lox/lox mice X ERT2-Adipoq-CRE mice. Mice were housed at 21°C with a 12:12 h light-dark cycle with free access to food and water. CRE activation was performed by 5 days intraperitoneal injection of tamoxifen resuspend in sunflower oil: ethanol (9:1) to 8 weeks-old males. The ethics committee of the French national veterinary agency approved all animal protocols used in this study.
## METHOD DETAILS

### Transfection and lipid droplet formation

Cells were seeded in MatTek 3.5 mm glass-bottom dishes (MatTek Corp., Ashland, MA) at least 16 h before transfection. Cells were transfected with 1–2 µg of plasmid DNA using JetPEI transfection reagent following the manufacturer guidelines for the DNA-transfection reagent ratios. Twenty-four hours post-transfection, the medium was replaced, and lipid incubation was performed when required. To induce triacylglycerol (TAG)-enriched lipid droplet (LD) formation, cells were incubated for 24 h in DMEM supplemented with fatty acids conjugated to bovine serum albumin (BSA) (1% v/v). In most experiments, 400 µM oleic acid was used to induce LDs. For experiments shown in Figures 5H, 5J, and S6M, cells were incubated with 200 µM palmitic acid and 200 µM oleic acid. For cholesterol-enriched LD formation in Figure S4H, cells were treated for 24 h with cholesterol complexed with methyl-β-cyclodextrin (molar ratio 1:20) to achieve a final cholesterol concentration of 200 µM. Lipid Droplets were labeled before visualization by incubating cells with HCS LipidTox Deep Red Neutral Lipid Stain (0.05% v/v) for 10 min in a 37°C humidified atmosphere containing 5% CO₂.

### Hypotonic medium

Cells were swollen by replacing the culture media with a hypotonic medium, (DMEM, in Milli-Q water' 5:95% v/v) for 10 min at 37°C in a humidified atmosphere with 5% CO₂ before visualization. This strategy was used to have spherical cells and quantify cell size or PM signals.

### Caveola proteins immunofluorescence staining

HeLa cells or fibroblasts were grown on coverslips, fixed with 4% paraformaldehyde for 20 min, and permeabilized for 20 min at 37°C using permeabilization buffer (PFS): DPBS with saponin (0.025% w/v) and gelatin from cold water fish skin (0.7% w/v). Cells were incubated with an anti-CAV1 antibody targeting the N-terminal region or an anti-Cavin1 antibody (Dilution 1:100 in PFS) for 2 h at 37°C. After washing three times for 5 min each with PFS, cells were incubated with an anti-mouse or rabbit Alexa 488-conjugated secondary antibody (Dilution 1:100 in PFS) for 60 min at 37°C. HCS LipidTox Deep Red Neutral Lipid Stain (0.1% v/v) was added during secondary antibody incubation in order to label LDs. The coverslips were washed twice with PFS and once with DPBS for 5 min each before being mounted on slides with ProLong Gold or Diamond.

### Micromanipulation

HeLa WT and SKO cells were prepared as described above. The micromanipulation measurements were arranged as previously described by Santinho et al. (Nature Communications, 2024).⁸⁷ Cells were swollen with diluted DMEM (5:95% v/v) at 37°C to create a hypotonic shock. The swollen cells were then placed on a BSA pre-treated coverslip and Bodipy 493/503 was added to visualize the membranes. The glass pipettes (pulled from borosilicate capillaries: 1.0OD, 0.58ID, 30-0017 GC100-15b; Harvard Apparatus, Hollisten, MA) were cleaned utilizing a plasma cleaner, covered with mPEG5K-silane and filled with diluted DMEM. The pipettes were placed in the homemade setup for the micromanipulation measurements and aligned with the microscope settings. Swollen cells were carefully captured with the micropipettes. The aspiration pressure was measured with a manometer (PCE P15) attached to the pipette and controlled manually by a syringe. The membrane tension © can be calculated using Laplace's law as follows:

$$ \gamma = \frac{\Delta P * R_p}{2 \left( 1 - \frac{R_p}{R_{cell}} \right)} \qquad (\text{Equation 1}) $$

With ⊗P being the suction pressure, Rₚ the inner pipette radius, and Rcell the radius of the swollen cell.

Lysis tension: the plasma membrane the initial aspiration pressure □ (at minimal aspiration to obtain a tongue in the pipette) and the final aspiration pressure □end (rupture of the membrane) was measured. The final aspiration pressure was achieved by increasing the aspiration at a constant rate (approx. 10 mbar/min) until the membrane ruptured by a pore opening. The lysis tension was then calculated via Equation 1. With ⊗P being the difference of suction pressure ⊗P = □end-□i.

Apparent area expansion modulus: Measurements were taken in tensed membrane regimes (above 0.75 mN/m). After obtaining the initial membrane tension ©0 the aspiration tension was slowly increased to different membrane tensions © with approx. 20 s waiting period in between each value to reach equilibrium. The membrane tongue length Lₚ,© and cell radius Rcell,© were measured for each applied aspiration tension. With the collected values the surface area A© and its variation of area ⊗A© can be calculated:

$$ A_{\gamma} = 2\pi \left( R_{cell,\gamma}^2 \left[ 1 + \sqrt{1 - \left(\frac{R_p}{R_{cell,\gamma}}\right)^2} \right] + R_p * L_{p,\gamma} \right) \qquad (\text{Equation 2}) $$

$$ \Delta A_{\gamma} = 2\pi \left( R_{cell,\gamma}^2 \left[ 1 + \sqrt{1 - \left(\frac{R_p}{R_{cell,\gamma}}\right)^2} \right] - R_{cell,\gamma0}^2 \left[ 1 + \sqrt{1 - \left(\frac{R_p}{R_{cell,\gamma0}}\right)^2} \right] + R_p (L_{p,\gamma} - L_{p,\gamma0}) \right) \qquad (\text{Equation 3}) $$
The applied membrane tension © was then plotted against the relative surface area variations of the plasma membrane $\frac{\Delta A_{\gamma}}{A_{\gamma,0}}$. By the apparent area expansion modulus $\langle_{\text{expansion}}$ the membrane tension can be linked to the surface area variation:

$$ \alpha_{\text{expansion}} = \frac{\gamma}{\frac{\Delta A_{\gamma}}{A_{\gamma,0}}} \qquad (\text{Equation 4}) $$

By extracting the slope (linear regression) of the membrane tension against the surface area variation, the $\langle_{\text{expansion}}$ can be obtained.

## Cells' area quantification

HeLa WT and SKO cells were seeded in MatTek 3.5 mm glass-bottom dishes and swollen with a hypotonic shock. To identify and visualize cells by confocal microscopy, Bodipy 493/503 (0.05% v/v) was added to the hypotonic media to label the inner and plasma membranes. To ensure a proper number of cells, z-stacks images were acquired with a wide field of view by confocal microscopy. Cell area was then quantified using Fiji Software by measuring the area of a circular selection from the median plane of swollen cells.

## LDs' area quantification

LDs' area was quantified using a Stardist home-trained model (trained on 106 cropped images via ZeroCostDL4Mic Google Colab Notebook, von Chamier et al., Nature Communications, 2021).88 The code for predictions was adapted from https://github.com/stardist/stardist. Cell masks were generated using CellPose (Stringer et al., Nature Methods, 2020).89 LD areas were measured using a CellProfiler 4.0 (Stirling et al., BMC Bioinformatics, 2021; McQuin et al., PLoS Biol., 2018; Kamentsky et al., Bioinformatics. 2011; Carpenter et al., Genome Biology. 2006)82–85 pipeline with the MeasureObjectSizeShape module. For each cell, the LD areas were averaged, and pixel values were converted to µm² using custom Python scripts and the corresponding file metadata.

## Protein-targeting to LD quantification

LD's protein-targeting quantification was performed by measuring the mean signal value from three selected points around each LD outline (Multipoint tool in Fiji software; Dots XXXL size). The values from these points were averaged. A basal cell signal was obtained by measuring mean values (Profile tool: 5-pixel width) from cytoplasmic areas without LDs or vesicles. For each cell, five points were taken and then averaged. For each LD, the mean protein signal was adjusted by subtracting the basal cell signal and then dividing by the basal cell signal. Negative values were set to zero. The normalized signals for all LDs in each cell were then averaged.

## CAV1-GFP PM signal quantification

CAV1-GFP plasma membrane localization was quantified in adherent HeLa WT and SKO cells (Figures 1A and 1B). Cell boundaries were segmented using Cellpose (Stringer et al., Nature Methods, 2020),89 and masks were processed in CellProfiler 4.0 (Stirling et al., BMC Bioinformatics, 2021; McQuin et al., PLoS Biol., 2018; Kamentsky et al., Bioinformatics. 2011; Carpenter et al., Genome Biology. 2006).82–85 A second mask was generated by shrinking the original by 1 pixel to exclude the plasma membrane region. Integrated intensities from both masks were used to calculate the membrane-associated signal by subtraction, which was then normalized to total cellular fluorescence to determine the percentage of CAV1-GFP at the plasma membrane.

To improve quantification consistency, cells were briefly incubated in hypotonic medium to enhance membrane uniformity. Z-stacks were acquired, and the median plane was used for analysis. In Fiji, the cell perimeter was manually traced to define a region of interest (ROI), then duplicated and shrunk by 0.35 µm to exclude the membrane signal. The membrane-associated intensity was obtained by subtracting the inner ROI signal from the full ROI, and normalized to total cell fluorescence.

## CAV1 immunofluorescence quantifications

For image analysis, cell segmentation masks were generated using Cellpose (Stringer et al., Nature Methods, 2020)89 to identify individual cells, while CAV1 puncta and lipid droplets (LDs) were segmented independently using StarDist (von Chamier et al., Nature Communications, 2021).88 The resulting masks were imported into a CellProfiler pipeline for quantitative analysis (Stirling et al., BMC Bioinformatics, 2021; McQuin et al., PLoS Biol., 2018; Kamentsky et al., Bioinformatics. 2011; Carpenter et al., Genome Biology. 2006).82–85 The pipeline began with a manual filtering step to exclude cells located at the image borders. Next, LD masks were expanded by 2 pixels, and a new mask of CAV1 puncta was generated by retaining only those puncta that overlapped with the expanded LD masks, thereby identifying CAV1 puncta associated with LDs. For each cell, we quantified: the number of CAV1 puncta associated with LDs, the total number of CAV1 puncta, and the total number of LDs. A CAV1 recruitment score was calculated for each cell as follows: Recruitment score = (CAV1 puncta on LDs)/(total puncta × total LDs).

## Blocking metabolism and lipid transport

To modulate metabolic pathways and lipid transport, several inhibitors were used in this study. PF-04620110, PF-06424439 (inhibitors of DGAT1 and 2 respectively), Myriocin, HPA-12 and 25-Hydroxycholesterol were added during transfection at a final concentration of 2.5 µM. To increase or decrease palmitic acid concentration, MF-438, an SCD1 inhibitor, or GSK837149A, a FASN inhibitor, were added at a final concentration of 1 µM, either 5 h before transfection for live-cell experiments or at least 16 h before lipid
loading for fixed-cell experiments. All inhibitors were maintained in the medium until visualization. Solvent controls were performed using DMSO (for Myriocin, HPA-12, MF-438, and GSK837149A) or ethanol (for 25-Hydroxycholesterol).

## Ceramides loading into HeLa cells

To load ceramides into HeLa cells, C18 Ceramide (d18:1/18:0), dissolved in ethanol, was mixed with DMEM and filtered to prepare a ceramide-enriched medium at a final concentration of 62.5 µM. This medium was added to MatTek dishes containing seeded and previously transfected cells during the 24-h lipid droplet induction step. During this step, oleic acid was also added at a final concentration of 400 µM, resulting in a final ceramide concentration of 50 µM. For C12 NBD Ceramide (d18:1/12:0) loading, an initial supplementary step involved evaporating the chloroform in which the lipids were dissolved, resulting in a pellet of ceramide-NBD. This pellet was subsequently dissolved in ethanol. The NBD ceramide solution was then mixed with an ethanol solution containing non-labeled ceramides at a molar ratio of labeled to non-labeled ceramides of 1:5. The final ethanol solution of ceramides was then combined with DMEM as described above.

## Trans-Golgi-network domains quantification

HeLa cells were seeded on 3.5 mm glass-bottom MatTek dishes. PK-Dye (final concentration: 0.01%) was a gift from Andrey S. Klymchenko (Valanciunaite et al., Analytical Chemistry, 2020).<sup>77</sup> It was added to the culture medium 10 min prior to imaging at 37°C and 5% CO₂ to allow cellular uptake. PK-Dye is a ratiometric fluorescent probe that exhibits a shift in emission from green to red depending on the local molecular environment. This shift is influenced by membrane order and polarity; in ordered membrane domains, such as those enriched in cholesterol and SM (e.g., within the trans-Golgi network), a higher green fluorescence signal is typically observed. To capture this environment-dependent spectral shift, cells were imaged using a confocal microscope with 488 nm laser excitation and two spectrally separated emission channels.

Quantification and analysis were conducted using the Ratio J 3.8 plugin in Fiji software. For each pixel, the ratio between the red signal (Channel 2) and the green signal (Channel 1) was calculated. In the presence of domains, the intensity of the green signal predominates, resulting in a lower ratio value. The plugin generated a plot of cumulative intensities corresponding to each ratio value. This study focused on the TGN domains, selecting ratio values between 0.85 and 1.5 as representative of this compartment. To normalize the plots created for wild-type (WT) and knockout (SKO) cells, cumulative intensities were transformed into relative intensity values (%). Normalization was performed by summing all intensities within the selected ratio value range of 0.85-1.5. Each intensity value was then expressed as a percentage of the total intensity within the range.

## Lipidomics, sample preparation, and analysis

HeLa cells were seeded on T-175 Petri dishes and grown to confluence. Once confluent, cells were placed on ice and washed twice with ice-cold PBS (pH 7.4). A 10 mM Tris/HCl buffer (pH 7.4) was then added to the cells for 1 min. The cells were scraped into a homogenization buffer containing 10 mM Tris/HCl, 1 mM EGTA, 0.5 mM EDTA, and 0.25 M sucrose at pH 7.4, supplemented with Complete protease inhibitors. The cell suspension was homogenized using ten strokes of a loose-fitting Dounce homogenizer. All subsequent steps were performed at 4°C. Post-nuclear supernatants (PNS) were obtained by centrifugation at 1000g for 10 min. A 2 mL aliquot of PNS was layered onto a sucrose gradient consisting of successive layers of 1 mL of 40% (w/v) sucrose, 1 mL of 30%, 2 mL of 25%, 2 mL of 20%, 2 mL of 15%, and 2 mL of 10% sucrose. The gradients were centrifuged for at least 16 h at 175,000 rpm. Following centrifugation, 10 mL fractions were collected starting from the top of the gradient. The pellet was resuspended in 1 mL of 0.25 M sucrose, as described by Omrane et al. (2019).<sup>90</sup>

SDS-PAGE, followed by western blotting, was performed to determine each fraction's organelle identity. Samples were boiled and denatured in Nupage LDS Sample Buffer at 95°C for five minutes. They were then loaded onto Nupage 4-12% Bis-Tris gels with a thickness of 1.5 mm and 15 wells. MES buffer or MOPS buffer were used as the running buffer. A semi-dry transfer was carried out at a constant current of 130 mA for 1 h. Blocking was performed with 0.1% PBS-Tween 20 and 5% milk for 1 h at room temperature. Membranes were incubated overnight at 4°C with the anti-Golgin-97 primary antibody solution. (Dilution 1:1000 in 0.1% PBS-Tween 20; 1% Milk). The next day, membranes were washed three times for 10 min each and then incubated with the HRP-conjugated anti-Rabbit secondary antibody (Dilution 1:1000 in PBS-Tween 20 0.1%; Milk 1%) for an hour at room temperature. ER enriched fractions were determined by incubating the membranes with an anti-Calreticulin antibody (1:1000 dilution in PBS-Tween20 0.1%; Milk 1%) for 2 h at room temperature; followed by an hour room temperature incubation with an anti-mouse-HRP conjugated antibody (1:1000-2000 dilution in PBS-Tween20 0.1%; Milk 1%), (Cat: SA1-100, Invitrogen). Detection was performed using a Pierce Western electrochemiluminescence kit on an ImageQuant LAS 4000 mini. Fractions 3 and 6 were used as Golgi and ER fractions for the lipidomics analysis.

To purify lipids from whole lysate, ER and Golgi supernatants were extracted using a modified Folch extraction method (Folch et al., J Biol Chem, 1957).<sup>91</sup> Supernatants were homogenized in chloroform/methanol (2:1) to a final volume of 5 mL. The mixture was agitated for 15-20 min on an orbital shaker at room temperature. After homogenization, the mixture was centrifuged to collect the liquid phase, which was then washed with 1 mL of water. Following brief vortexing, the mixture was centrifuged at 2000 rpm to separate the two phases, and the lower chloroform phase containing lipids was carefully recovered. The lipid-containing lower phase was evaporated under a nitrogen stream, and the lipid extract was resuspended in 1 mL of chloroform/methanol (2:1). The solution was transferred to a 1.5 mL Eppendorf tube and further dried under a nitrogen stream. The final lipid extract was resuspended in 100 µL of
chloroform/methanol (2:1) for subsequent analysis. Samples were analyzed using the Institut Curie lipidomics platform's mass spectrometry (MS). Data were normalized to standardize the total lipid content across all samples, mitigating differences in total material quantity between samples. A Z score was then calculated for each replicate of the knockout (SKO) samples, from either the ER or Golgi, using the following formula: (Observed mean - Theoretical mean)/Standard deviation, where the mean of the wild-type (WT) samples (Golgi or ER) was used as the theoretical mean. Saturated, monounsaturated, and polyunsaturated lipids were then categorized, and the total amount of saturated lipids was summed.

## RUSH experiment

HeLa WT and SKO cells were plated on MatTek 3.5 mm glass-bottom dishes and transfected with GPI-RUSH (Boncompain et al., Nature Methods. 2012).<sup>47</sup> In one condition, no biotin was added, while in two conditions, fresh media containing 40 µM D-biotin (Invitrogen, Cat# B1585) was added. Cells were incubated with biotin for either 20 or 60 min at 37°C in a humidified atmosphere with 5% CO<sub>2</sub>. After incubation, cells were fixed with 4% paraformaldehyde for 20 min, stored in DPBS at 4°C, and visualized the following day.

For images taken at 20 min, the ER and Golgi signal were quantified by measuring the mean GPI signal in each organelle five times (Profile tool: 5-pixel width) and averaging the values. The mean Golgi signal was then divided by the mean ER signal (Golgi/ER).

At 60 min, the plasma membrane signal was measured by drawing profiles across the membrane (5-pixel width, maximum value). Five points were measured per cell and averaged. The Golgi signal was also measured by drawing profiles and averaging five points. The plasma membrane-to-Golgi ratio (PM/Golgi) was then calculated for each cell.

## Golgi aspect ratio

HeLa cells and fibroblasts seeded on coverslips were stained with immunofluorescence for GM130 (Dilution 1:100; Secondary antibody: Anti-ms Alexa 488, Dilution 1:100) or Golgin-97 (Dilution 1:100; Secondary antibody: Anti-rb Alexa 488, Dilution 1:100). Nucleus were labeled with Hoechst 33342 Solution (0.1% v/v) and LDs with HCS LipidTox Deep Red Neutral Lipid Stain. For one fibroblast experiment, cells were treated with 400 µM of oleic acid for 24 h before fixation. Z-projections were taken using the maximum value and Golgi masks were created with Otsu thresholding (Otsu, 1979).<sup>92</sup> Particles were analyzed (minimum size threshold of 0.05 µm<sup>2</sup>) by measuring area and Feret diameter, with both minimum and maximum Feret values recorded for each particle. The particle with the largest area was used to represent the overall shape, and the aspect ratio was calculated by dividing the minimum Feret by the maximum Feret.

## Quantification of OSBP, P4M, Lysenin & CERT

HeLa WT and SKO cells were plated on MatTek 3.5 mm glass-bottom dishes, transfected (OSBP-mCherry, P4M-GFP, GFP-CERT or LyseninSM) and visualized by confocal microscopy at 37°C using a heating plate. Z-stacks were taken and Z-projection averaging was performed. Masks were created using Otsu thresholding from the perinuclear signal. Particles were analyzed (minimum size threshold of 0.05 µm<sup>2</sup>) by measuring area, mean signal, and integrated intensity, and the sum of the integrated intensity was calculated. The cytoplasmic signal was measured by drawing five profiles (5-pixel width). The sum of integrated intensity was then divided by the mean cytoplasmic signal.

## ER purification from mice extracts

Fresh WAT (1g) was immediately weighed and transferred in PBS. The tissue was minced and submitted to collagenase type II digestion (25g/Kg) in DMEM medium supplemented with 10% SVF, 60 min at 37°C under stirring (90rpm). The enzyme was neutralized, and the solution was centrifuged for 10 min at 700g. Digestion released floating adipocytes. The adipocyte ring was resuspended in cold STM buffer (250 mM sucrose, 50 mM Tris, HCl (pH 7.4), 5 mM MgCl<sub>2</sub>) and the cells were disrupted successively with glass potter. The homogenate was centrifuged at 700g for 10 min. The supernatant was centrifuged for 15 min at 10500g. The supernatant was saved and centrifuged at 100.000g for 1h, resulting in the isolation of the ER fraction as the pellet). ER fractions were then analyzed by mass spectrometry for lipidomics and proteomics analysis. The latter was realized by the ProteiSeine Plateform of Institut Jacques Monod UMR 7592.

## Myriocin mice treatment

Bscl2 lox/lox mice were crossed with ERT2-Adipoq-CRE (Sassmann et al., Genesis, 2010)<sup>86</sup> mice to generate Bscl2lox/lox mice X ERT2-Adipoq-CRE mice. Mice were housed at 21°C with a 12:12 h light-dark cycle with free access to food and water. To induce Cre-mediated recombination and deletion of the *Bscl2* gene, iATSKO mice were administered tamoxifen via intraperitoneal (i.p.) injection at a dose of 100 mg/kg body weight once daily for five consecutive days. Following a recovery period, mice were treated with myriocin to inhibit *de novo* sphingolipid synthesis. Myriocin was administered by i.p. injection at a concentration of 0.3 mg/kg every other day for a total duration of 4 weeks.

## Lipidomics of mice extracts

Total adipose tissue or endoplasmic reticulum (ER) fractions were first resuspended in 400 µL of deionized water. Each sample was mixed with 1600 µL of a cyclohexane-isopropanol solution in a 3:2 ratio for lipid extraction. The mixtures were then agitated for 1 h at
room temperature. Following agitation, the samples were centrifuged at 10,000 g for 5 min. The upper organic phase was carefully collected and transferred to a glass tube, evaporating at 60°C under nitrogen gas to remove the solvent. The lipid phase was subsequently resuspended in a solvent mixture of isopropanol, acetonitrile, and water (65:30:5) and subjected to analysis using a liquid chromatography-mass spectrometry (LC-MS) system.

## Electron microscopy of caveola

Adipose tissue samples (1 mm³) were initially fixed in 2.5% glutaraldehyde (LFG Distribution, Lyon, France) in 0.1 M Sorensen buffer (pH 7.4) at 4°C for 16 h. Following fixation, the samples were rinsed with 0.2 M cacodylate buffer (pH 7.4) and subsequently post-fixed in a solution of 2% osmium tetroxide and 1.5% potassium ferrocyanide in 0.2 M cacodylate buffer (pH 7.4) for 45 min at room temperature.

After post-fixation, the samples were washed with milliQ water and dehydrated through a graded ethanol series. The dehydrated samples were then embedded in EMbed 812 epoxy resin (LFG Distribution), polymerizing at 60°C for 48 h.

Semithin sections (1 µm thickness) were prepared using an EM-UC7 ultramicrotome (Leica, Nanterre, France) and stained with 1% methylene blue solution and 1% Azur II. These sections were examined using an Olympus AX-60 microscope (Olympus, Rungis, France) equipped with a QImaging QIClick digital CCD camera (QImaging, Surrey, Canada).

For transmission electron microscopy, thin sections (60 nm thickness) were cut and analyzed with a Jeol JEM 1400 microscope (Jeol, Croissy sur Seine, France) operated at an accelerating voltage of 120 kV.

## SCD1 and CAV1-GFP expression analysis

HeLa WT and SKO cells were seeded onto six-well plates and transfected with CAV1-GFP. Cells were harvested by scraping in 1 mL of RIPA buffer 48 h post-transfection, Protein concentrations were determined using a BCA assay. Proteins denaturation was performed with NuPAGE LDS Sample Buffer by heating at 95°C for 5 min in presence of 0.45 µM of DTT. SDS page was realized by loading 10 µg of protein for each sample using with MES as running buffer at 150V. Proteins were subsequently transferred onto nitrocellulose membranes at a constant current of 130 mA for 1 h. Membrane blocking was performed for an hour at room temperature with PBS-Tween 20 0.1% containing 5% of Milk. Primary antibody incubation against GFP was performed overnight at 4°C, followed the next day by three washes of 10 min each with PBS-Tween 0.1%. Secondary antibody incubation was conducted at room temperature for 1 h and 30 min using an HRP-conjugated mouse antibody (dilution 1:2000 in PBS-Tween 0.1%; 1% milk). To verify equal protein loading across conditions, membranes were incubated with a mouse anti-GAPDH antibody, using the same procedure as described above. Detection of protein bands was carried out using the Pierce Western ECL substrate and images were acquired with an ImageQuant LAS 4000 mini system.

The same protocol was used for SCD1 expression analysis, excluding the transfection step. A rabbit anti-SCD1 primary antibody (1:21,000 dilution in PBS-Tween 0.1% with 1% milk) was used overnight at 4°C, followed by a 1-h incubation with an anti-rabbit HRP-conjugated secondary antibody (1:1000). Beta-actin was used as a loading control and detected with a mouse anti-β-actin antibody (1:1000, 2 h at RT), followed by an HRP-conjugated anti-mouse secondary antibody (1:1000, 1 h at RT), all in PBS-Tween 0.1% with 1% milk.

## SCD1 overexpressing cell lipidomics

A total of 100,000 SKO cells were seeded into six-well plates and either left untransfected or transfected with SCD1. Lipid droplet induction was performed 24 h later by incubating cells with 400 µM oleic acid for 24 h. Following incubation, the medium was removed, and cells were washed with 1 mL of ammonium bicarbonate. Cells were then scraped and collected into 1.5 mL of ammonium bicarbonate. Samples were centrifuged at 13,000 rpm for 5 min at 4°C. The resulting supernatant was discarded, and the pellet was resuspended in 200 µL of ammonium bicarbonate. Samples were snap-frozen in liquid nitrogen and stored at -80°C until further analysis. Lipidomic profiling was conducted using mass spectrometry at the Institut Curie lipidomics platform. Data were normalized to standardize total lipid content across all samples. A Z score was calculated for each replicate of SKO cells overexpressing SCD1, using untransfected SKO samples as the reference for mean and standard deviation.

## CERT knockdown experiments

HeLa WT were seeded in MatTek 3.5 mm glass-bottom dishes. The day after siRNA knockdown of CERT was realized with OptiMem and Lipofectamine RNAiMAX Transfection Reagent following the manufacturer guidelines. A control was realized by doing a siRNA mediated knockdown of Luciferase. Media was changed the following day and cells were transfected with CAV1-GFP for 24 h. Subsequently, lipid LDs induction was carried out the day after by incubating cells for 24 h with 400 µM oleic acid.

## TIRF imaging of CAV1-GFP

HeLa WT and SKO cells were seeded in 3.5 mm glass-bottom dishes (MatTek). Cells were transfected with CAV1-GFP for 24 h. The following day, lipid droplets were induced by incubating cells with 400 µM oleic acid for an additional 24 h. TIRF microscopy was performed at the Imaging Core Facility of Institut Gustave Roussy (Université Paris-Saclay, INSERM US23/CNRS UAR 3655). Quantification was conducted using Fiji software by applying Otsu thresholding (Otsu, 1979)⁹² to segment CAV1-positive puncta, followed by analysis of particle number and area.
Confocal imaging, data representation

All images were acquired using a Carl ZEISS LSM 800 microscope. Green fluorescent proteins or dyes were excited at 488 nm, and emission was detected between 510 and 550 nm. Red-tagged proteins and dyes were excited at 540 nm, with emission detected between 580 and 650 nm. Deep-red fluorescence was excited at 640 nm, and emission was detected above 650 nm.

Image analysis and quantification were performed using ImageJ. OPF LUTs (Orange, Purple, Fresh green) were obtained from the ChristLUTs GitHub repository (https://github.com/cleterrier/ChrisLUTs/tree/master). All images were visualized and analyzed in Im- ageJ. Figures were created using Affinity Designer 2. In some instances, images were modified in Affinity Designer by applying effects to the corresponding layers (e.g., Figures 2A, 2C, S2G, and S2H, shading: negative and then dividing). Data plots were generated using GraphPad Prism 8.0 and further modified in Affinity Designer 2. The plots are represented as scatterplots, where each dot cor- responds to an individual cell. Large dots (supermeans) represent the mean value of each replicate. The middle line represents the mean of the supermeans, and the extremities indicate the standard deviation of the supermeans.

QUANTIFICATION AND STATISTICAL ANALYSIS

Statistical analyses were performed using GraphPad Prism 8.0. Normality of data distribution was assessed using the D'Agostino-Pearson test at a significance level of *p* = 0.05. For smaller sample sizes, the Shapiro-Wilk test was applied using the same threshold. Parametric unpaired t-tests were used for data that followed a normal distribution. If no statistical test is explicitly mentioned but *p*-values are indicated, unpaired t-tests were used by default. For non-normally distributed data, two-tailed Mann-Whitney tests were applied, as noted in the figure legends where applicable. Asterisks in the figures indicate significance levels as follows:* *p* < 0.05; ** *p* < 0.01; *** *p* < 0.001; **** *p* < 0.0001. Graphical results are shown as Supermeans—the mean of each biological replicate—when possible. Error bars represent the Mean ± SD of the Supermeans. When Supermeans are not used, graphs display the Mean ± SD of the dataset.