# The α/β hydrolase domain-containing protein 1 (ABHD1) acts as a lysolipid lipase and is involved in lipid droplet formation

Ismael Torres-Romero, Bertrand Légeret, Marie Bertrand, Damien Sorigue, Alicia Damm, Stéphan Cuiné, Florian Veillet, Carla Blot, Sabine Brugière, Yohann Couté, et al.

<figure><img src="image_1.png" /></figure>

## To cite this version:

Ismael Torres-Romero, Bertrand Légeret, Marie Bertrand, Damien Sorigue, Alicia Damm, et al. The α/β hydrolase domain-containing protein 1 (ABHD1) acts as a lysolipid lipase and is involved in lipid droplet formation. National Science Review, 2024, 11 (12), pp.nwae398. 10.1093/nsr/nwae398. hal-04807931

HAL Id: hal-04807931

https://hal.science/hal-04807931v1

Submitted on 4 Apr 2025

HAL is a multi-disciplinary open access archive for the deposit and dissemination of scientific research documents, whether they are published or not. The documents may come from teaching and research institutions in France or abroad, or from public or private research centers.

L'archive ouverte pluridisciplinaire HAL, est destinée au dépôt et à la diffusion de documents scientifiques de niveau recherche, publiés ou non, émanant des établissements d'enseignement et de recherche français ou étrangers, des laboratoires publics ou privés.

<figure><img src="image_2.png" /></figure>

Distributed under a Creative Commons Attribution 4.0 International License
¹Aix Marseille Univ.
CEA, CNRS, Institute of Bioscience and Biotechnology of Aix Marseille, BIAM, Saint-Paul-Lez-Durance 13108, France;
²Laboratoire de Physique de l'École Normale Supérieure, ENS, Université PSL, CNRS, Sorbonne Université, Université de Paris Cité, Paris 75005, France;
³Univ. Grenoble Alpes, INSERM, CEA, UMR BioSanté U1292, CNRS, CEA, FR2048, Grenoble 38000, France;
⁴Institute of Biological Chemistry, Washington State University, Pullman 99164, USA and
⁵Single-Cell Center, CAS Key Laboratory of Biofuels and Shandong Key Laboratory of Energy Genetics, Qingdao Institute of Bioenergy and Bioprocess Technology, Chinese Academy of Sciences, Qingdao 266101, China

α/β hydrolase domain-containing protein 1 acts as a lysolipid lipase and is involved in lipid droplet formation

Ismael Torres-Romero¹,†, Bertrand Légeret¹,†, Marie Bertrand¹, Damien Sorigue¹, Alicia Damm², Stéphan Cuiné¹, Florian Veillet¹, Carla Blot¹, Sabine Brugière³, Yohann Couté ID ³, Matthew G. Garneau ID ⁴, Hari K. Kotapati⁴, Yi Xin⁵,‡, Jian Xu ID ⁵, Philip D. Bates⁴, Abdou R. Thiam², Fred Beisson ID ¹ and Yonghua Li-Beisson ID ¹,*

ABSTRACT

Lipid droplets (LDs) are the major sites of lipid and energy homeostasis. However, few LD biogenesis proteins have been identified. Using model microalga *Chlamydomonas*, we show that ABHD1, an α/β-hydrolase domain-containing protein, is localized to the LD surface and stimulates LD formation through two actions: one enzymatic and one structural. The knockout mutants contained similar amounts of triacylglycerols (TAG) but their LDs showed a higher content of lyso-derivatives of betaine lipid diacylglyceryl-*N*,*N*,*N*-trimethylhomoserine (DGTS). Over-expression of *ABHD1* increased LD abundance and boosted TAG content. Purified recombinant ABHD1 hydrolyzed lyso-DGTS, producing a free fatty acid and a glyceryltrimethylhomoserine. *In vitro* droplet-embedded vesicles showed that ABHD1 promoted LD emergence. Taken together, these results identify ABHD1 as a new player in LD formation by its lipase activity on lyso-DGTS and by its distinct biophysical property. This study further suggests that lipases targeted to LDs and able to act on their polar lipid coat may be interesting tools to promote LD assembly in eukaryotic cells.

**Keywords:** algae, betaine lipid, α/β hydrolase, lipid droplet, lysolipid

INTRODUCTION

Lipid droplets (LDs) are the site of lipid storage accumulation in eukaryotic cells. LDs are made of a neutral lipid core, consisting of mostly triacylglycerols (TAGs), and are surrounded by a monolayer of polar lipids (hemi-membrane) embedded with proteins [1]. LDs evolved to sequester the non-membrane-forming neutral lipids, i.e. TAG, away from membranes. In addition to their major role as carbon storage, LDs contribute to maintain energy homeostasis, prevent lipotoxicity, act as a temporary depot for acyl chains and participate in lipid signaling [2–5]. Thermodynamically speaking, the oil-in-water configuration of LDs sets them apart from all other subcellular organelles. Their formation therefore requires tight coordination between the biochemical properties of their neutral lipid core and am-

Philipatic polar lipids, and those of their protein coat. Accumulating evidence points to the importance of both proteins and lipids in maintaining a thermodynamically stable LD configuration and population [6–8].

LD biogenesis occurs in response to environmental cues (such as light, nutrition and temperature) or is developmentally regulated (as in seeds or in aging cells). Considering the paramount importance of TAG metabolism for human health and for biotechnology, LD formation has been studied intensively in various organisms ranging from yeast to mammals, land plants and microalgae [3,9]. Current understanding is that the LD, at least most of it, emerges from specific subdomains of the endoplasmic reticulum (ER) membrane defined by the enrichment of various proteins including the protein SEIPIN found

<footnote>*Corresponding author. E-mail: yonghua.li@cea.fr</footnote>

<footnote>†Equally contributed to this work.</footnote>

<footnote>‡Present affiliation: State Key Laboratory of Marine Resource Utilization in the South China Sea, Hainan University, Haikou 570228, China.</footnote>

<footnote>Received 28 January 2024; Revised 14 September 2024;</footnote>

<footnote>Accepted 20 September 2024</footnote>
in mammals, plants and fungi [10]. Initial membrane
lipid remodeling and TAG synthesis results in the
formation of an ‘oil lens’ between the leaflets of the
ER bilayer, which is then followed by its growth, and
finally it pinches off the membrane toward the cy-
tosol, giving rise to an LD [6]. It is worth pointing
out that new evidence suggests that LDs may not
necessarily ever pinch off from the ER, but rather
remain attached to the ER [11]. Thanks to the de-
velopment of affordable proteomics tools, LD pro-
teomes of many species along the evolutionary tree
are now published, pointing to the divergent nature
of the proteins that are associated with LDs [12,13].
Up to now, however, only a handful of proteins have
been demonstrated to play a role in LD biogenesis in
plants, animals or yeast. These include oleosin, cale-
osin, PERILIPIN1, FSP27 (fat-specific protein 27),
FIT2 (fat storage-inducing transmembrane protein
2), the FATP1-DGAT2 (fatty acid transport protein
1-diacylglycerol acyltransferase 2) complex, the LD-
associated protein (LDAP), and finally SEIPIN with
its interactors, i.e. LD-assembly factor 1 (LDAF), the
LDAP-interacting protein (LDIP) or the membrane-
tethering protein VAP27-1 (vesicle-associated pro-
tein 27-1) [1,9,14–23].

In the past 10 years, *Chlamydomonas reinhardtii*
has been used as a model green microalga to in-
terrogate LD biogenesis and turnover [2]. LD for-
mation and degradation in *Chlamydomonas* can be
manipulated easily, for example, by simply changing
the nitrogen (N) level in the media [24]. N star-
vation is one of the most potent triggers in induc-
ing TAG accumulation, which results in a drastic in-
crease in both LD number and size in algal cells [24].
In addition to research on environmental conditions
or the genetic basis of TAG metabolism and reg-
ulation [14,24–35], the protein and lipid composi-
tion of the LD coat has also been reported [32,36–
38]. The monolayer lipid-coat of an LD found in
*Chlamydomonas* is made predominantly of the be-
taine lipid diacylglyceryl-N,N,N-trimethylhomo-Ser
(DGTS), followed by sulfoquinovosyl diacylglyc-
erol (SQDG) and phosphatidylethanolamine (PE)
and small amounts of galactolipids [32]. Over 200
proteins were found in an LD-enriched fraction in
*Chlamydomonas* [37], but only a few have been
characterized. This includes: the major lipid droplet
protein (MLDP), considered a structural protein
of *Chlamydomonas* LDs [36,39], the betaine lipid
synthase BTA1 [40], the long-chain acyl activat-
ing enzyme LCS2 [41,42] and various lipid traf-
ficking proteins [43]. No protein involved in LD
budding and expansion has been identified yet
in algae.

To investigate LD biogenesis in *Chlamydomonas*,
we have focused on a putative α/β hydrolase

domain-containing protein (ABHD1), which is
present in the proteomics of LDs isolated from
N-starved and high light-exposed cells [32,37].
Using cell biological, biophysical, genetic (gain-
and loss-of-function) and biochemical studies in
*Chlamydomonas*, we demonstrate here that ABHD1
is a lipase mostly acting on lyso-DGTS that can pro-
mote LD formation *in vivo* and *in vitro*. A tentative
model describing the dual role—enzymatic and
non-enzymatic—of ABHD1 in the mechanism of
LD formation is also proposed.

RESULTS

Gene expression and structural and
evolutionary features of ABHD1 protein

Searching public databases on mRNA expression
levels for ABHD1, one can observe that its gene ex-
pression is increased 2-fold upon N starvation [44].
Furthermore, our immunoblot using anti-ABHD1
antibodies showed that ABHD1 significantly in-
creased in *Chlamydomonas* cells during N depri-
vation, a condition promoting TAG synthesis and
LD formation (Fig. 1A, Fig. S1). Taken together,
these data strongly suggest that ABHD1 may play
a role in LD formation in *Chlamydomonas*. In ad-
dition to ABHD1, the *Chlamydomonas* genome en-
codes another protein (Cre01.g010550) showing
51% sequence identity to ABHD1, which we named
ABHD2. ABHD2 is absent in the three previously
published LD proteomics for *Chlamydomonas* either
isolated from N-starved cells or after high light expo-
sure [32,45,46], and moreover its expression is not
regulated by N deprivation [37,44], therefore it is
very unlikely that ABHD2 plays a complementary
role in terms of LD biology.

To gain insight into ABHD1 protein functions,
we examined its primary amino acid sequence, its
hydrophobicity [47] and its predicted structural fea-
tures using alpha-fold [48]. ABHD1 is composed of
three major parts: an N-terminal transmembrane do-
main, one central α/β-hydrolase fold-containing do-
main, and an intrinsically disordered domain at its C-
terminus (Fig. S2A–C). In addition, the occurrence
of a hydrophobic helix (alanine-rich domain) at its
C-terminus (AA 396–490) (Fig. S2B and C) could
possibly serve as an anchor to LDs. Similar structural
motifs at the C-terminus of the phosphatidylserine-
specific phospholipase A1 have been shown to bind
lipids [49]. In terms of hydrophobicity (Fig. S2E),
ABHD1 shares similar overall hydrophobic features
to MLDP but is a bit less hydrophobic (Gravy in-
dex of 0.05 compared to 0.11 for MLDP) and does
not contain the major hairpin hydrophobic domain
present in oleosins [36].
<figure><img src="image_3.png" /><figcaption>**Figure 1.** ABHD1 protein is located on the surface of LDs and its quantity increases with prolonged N starvation. (A) ABHD1 protein level as detected by immunoblot analysis using anti-ABHD1 antibodies. Total protein extracts (from 0.4 million cells) were loaded onto each lane. Ponceau's stained proteins transferred onto nitrocellulose membrane were shown as the control for protein load (image height of whole lane was compressed). Abbreviations: α, antibody; d, day; N, nitrogen; TAP, tris-acetate-phosphate media. (B) Confocal imaging. Cells were cultured in TAP-N (1 d) and stained with BODIPY to observe LDs. Pseudo-colors were used: BODIPY-stained LDs in yellow, chlorophyll autofluorescence in blue, mCherry and mCherry-tagged ABHD1 in red. DIC: differential interference contrast. Bar = 10 µm.</figcaption></figure>

A phylogenetic tree based on an amino acid sequence was constructed to explore the evolutionary history of ABHD1, which included 92 representative proteins from different clades of life (Table S1). The closest homologs to ABHD1 were grouped in a clade distinct from bacterial and cyanobacterial homologues and this clade only included algae of the Chlorophytina taxon, which is composed of the classes Ulvophyceae, Trebouxiophyceae and Chlorophyceae (Fig. S3). ABHD1 homologs are absent in plant or mammalian cells, therefore ABHD1 is specific to Chlorophyta. The lack of homology in these different lineages seems to have been a rule rather than an exception for LD-associated proteins such as, for example, the protein Delayed in TAG Hydrolysis 1 (DTH1) [35]. Increasing evidence seems to point out that organisms from distinct lineages equip their LDs with unique sets of proteins that do not share primary sequence homology but follow the

overarching principles in LD structure/architecture and function.

## ABHD1 coats the entire LD and its over-expression induces LD formation

To investigate the functional role of ABHD1, we first sought to confirm that the protein was actually associated with LDs *in vivo* and determine whether it is its major or only location. We therefore fused the fluorescent protein mCherry at the N- or C-terminus of the ABHD1 protein (Fig. S4A, Table S2) and transformed the nuclear genome of *Chlamydomonas* with a construct expressing this chimeric protein under the strong constitutive photosystem I protein D (PSAD) promoter. To facilitate LD identification under the confocal microscope, N-deprived cells were examined and stained with the lipid dye BODIPY. More than 500 antibiotic-resistant *Chlamydomonas* transformants (both N- and C-terminal fusions) were screened based on mCherry signal. The mCherry signal was detected all around LDs, regardless of the N- or C-terminal position of the tag (Fig. 1B, Figs S5 and S6, Movie S1). Nevertheless, a higher number of positive lines were obtained when mCherry was fused to the N-terminus of ABHD1 protein. The fusion of mCherry to the C-terminus of ABHD1 seems to be less stable, leading to a higher rate of truncated or misfolded chimeric protein. If the disordered C-terminus region is important for proper folding and 3D structure or targeting to LDs, this could explain the lower amounts of over-expressors (OEs) identified (Fig. S2B). We have therefore focused on the lines where mCherry is fused to the N-terminus of the ABHD1 protein to minimize misfolded proteins, such as aggregates or mis-targeted protein, which could occasionally be observed via mCherry signal in the cytosol. A more likely reason for mCherry signal in the cytosol is the use of the strong PSAD promoter, which resulted in potential ABHD1 over-expression. The robust signal emitted by mCherry fluorescence around the LD surface strongly indicates that ABHD1 is essentially a LD-associated protein and coats the entire LD (Movie S1).

While performing the subcellular localization experiment, we noticed LDs seem to be more abundant in *ABHD1* overexpressing lines during optimal growth than in the control parental line i.e. UVM4 cells (Fig. 2A). Lipid analysis revealed that, while there was no difference in TAG content during N starvation between the *ABHD1* OEs and the parental strain UVM4 (Fig. S4B), OEs contained more TAGs during N replete growth (Fig. 2B) without significant difference in TAG composition (Fig. S4C). Interestingly, the number of LDs per
<figure><img src="image_4.png" /><figcaption>**Figure 2.** *ABHD1* over-expression results in LD formation and a higher TAG content. (A) LD imaging in cells over-expressing *ABHD1* in nitrogen-replete mixotrophic growth, i.e. TAP medium (three independent lines are shown, bar = 5 µm). (B) TAG amount in WT versus *ABHD1* over-expressing lines during nitrogen replete mixotrophic growth. Four independent experiments, for six independent lines (three technical replicates for each line), are shown. Error bars show mean and standard deviation, Mann-Whitney test: \*\*\*P = 0.0002. (C) The distribution of LD numbers per cell during nitrogen replete mixotrophic growth (>200 cells examined per group). Histograms represent mean and standard deviation of three biological replicates for WT and five independent lines for OE (*ABHD1* over-expressor). (D) TAG quantification in the H1246 yeast strain expressing the *Chlamydomonas ABHD1* gene. Yeast transformants expressing the respective gene were harvested and analyzed for total TAG amount using gas chromatography-mass spectrometry (GC-MS). Data are means of three biological replicates with standard deviation bars shown. A significance of *q* = 0.13 for empty vector (EV) vs. +CrABHD1, *q* = 0.02 for EV vs. +ScDGA1 and *q* = 0.13 for +ScDGA1 vs. +CrABHD1 was determined using the Kruskal-Wallis test followed by a two-stage step-up Benjamini, Krieger and Yekutieli false discovery rate procedure for multiple comparison correction.</figcaption></figure>

cell was also altered, with a shift toward a higher number of LDs per cell in the OEs compared to the parental line (Fig. 2C). These results thus indicate that in *Chlamydomonas* ABHD1 is not a limiting factor for LD accumulation under N-depletion but can induce the formation of LDs, or block LDs' degradation under N-replete conditions when over-expressed. In order to see whether ABHD1 could induce LD formation in cells other than a microalga, we also expressed *ABHD1* in the quadruple yeast mutant H1246 strain where TAG synthesis

is greatly reduced. The yeast H1246 strain is knocked out in genes encoding a diacylglycerol acyltransferase1 (DGA1), lecithin cholesterol acyltransferase (LRO1) and acyl-coenzyme A: cholesterol acyl transferase-related enzyme 1 and 2 (ARE1 and ARE2). It has greatly reduced but not nulled TAG synthesizing activity [50]. *Chlamydomonas ABHD1 (CrABHD1)* expression alone in the *Saccharomyces cerevisiae* H1246 mutant resulted in a significant increase in TAG compared to the vector control line (Fig. 2D). This increase in TAG was not as dramatic as with the native *S. cerevisiae* TAG-synthesizing enzyme diacylglycerol acyltransferase 1 (ScDGA1), but it was nevertheless not negligible. This result suggests that a similar role of ABHD1 either in LD induction or in preventing LD turnover/TAG degradation exists in yeast. It also rules out that the observed TAG increase in *Chlamydomonas* ABHD1 OEs is due to the presence of the mCherry tag.

## Absence of ABHD1 affects the composition of the LD coat

To further investigate the function of ABHD1 in LD biogenesis, we isolated two insertional mutants from the *Chlamydomonas* mutant library (CLIP) [41]. The two independent lines *abhd1-1* and *abhd1-2* harbor an insertion at the 3rd and 10th exon, respectively (Fig. 3A). Using gene specific primers, reverse transcription-polymerase chain reaction (RT-PCR) analyses showed that there was no *ABHD1* expression in *abhd1-1* and *abhd1-2* mutant alleles (Fig. 3B). Immunoblot analyses of total proteins extracted from isolated LDs from respective WT and the two mutant strains showed that no ABHD1 protein could be detected in either mutant (Fig. 3C). Interestingly, the immunoblot from WT cells indicated that ABHD1 could be present as a dimer. Taken together, both gene expression and immunoblot analyses firmly confirmed both alleles as true knockout mutants. We then compared the TAG amount in the two mutants (*abhd1-1* and *abhd1-2*) to their parental strain (i.e. CC4533). No difference in TAG content was observed under N-replete or N-deplete conditions (Fig. S7A and C). Whole-cell lipidomics showed no significant change in TAG composition nor membrane lipids either (Fig. S7B, D, E). LD number and size are also similar between CC4533 and the two knockout mutants (Fig. S7F and G).

To investigate further the role that ABHD1 may have in LD formation *in vivo*, we performed comparative lipidomics and proteomics of isolated LDs from N-starved WT and the two knockout mutants. DGTS was identified as a major component of LD lipid-coat, followed by SQDG and PE. The mutants' LD coat showed a remodeled lipid
<figure><img src="image_5.png" /><figcaption>**Figure 3.** Isolation and characterization of lipids for the two *abhd1-1* and *abhd1-2* mutants. (A) The site of the *AphVIII* cassette insertion in the *ABHD1* gene for *abhd1-1* and *abhd1-2* mutants. (B) RT-PCR analysis of *ABHD1* transcript. (C) Immunoblot analysis of ABHD1 protein on isolated LDs from the three strains. LDs were extracted from cells N-starved for 2 d. Proteins were extracted then loaded into each lane, based on a fixed number of fatty acid methyl esters (FAMEs) (i.e. ca. 55 µg FAME equivalent of LDs loaded). Silver-stained SDS-PAGE of LD-isolated proteins acts as a loading control. (D) Membrane coat composition of isolated LDs from nitrogen starved cultures. Levels of lyso-DGTS species detected in the polar lipid fraction of isolated LDs. Signal from ultra-high performance liquid chromatography with quadrupole time-of-flight mass spectrometry (LC-MS) was normalized by FAME quantities in the polar lipid fraction. (E) Changes in LD lyso-DGTS species in the mutants in comparison to WT during nitrogen starvation. For both (D) and (E): data are means of five biological replicates from three independent LD isolations. Bars indicate standard deviation. *P* values were determined, after passing Bartlett's test for homoscedasticity and Shapiro-Wilk's test for normality, by a two-way ANOVA with Bonferroni correction: \* *P* < 0.05; \*\* *P* < 0.01. Abbreviations: CBLP: *Chlamydomonas* beta subunit-like polypeptide; FAME, fatty acid methyl ester; MGDG, monogalactosyldiacylglycerol; DGDG, digalactosyldiacylglycerol; PG, phosphatidylglycerol; SQDG, sulfoquinovosyldiacylglycerol; DGTS, diacylglyceryl-*N*,*N*,*N*-trimethylhomoserine; PE, phosphatidylethanolamine; PI, phosphatidylinositol.</figcaption></figure>

composition (Fig. 3D), with a lower proportion of the major lipid class (DGTS) but a higher proportion of SQDG, a class of negatively charged lipids. Among the lipid species with the most significant differences were some lyso-DGTS species, in particular lyso-DGTS 16:0, which were present in significantly higher amounts in the mutants' LD coat compared to WT (Fig. 3E).

To determine whether the mutants affected the flux of nascent fatty acids through membrane lipids into TAG during N-starvation, we performed a [¹⁴C]acetate pulse-chase analysis (Fig. S8). There were limited differences in the precursor-product relationships of membrane lipids and TAG between WT, *abhd1* mutants and the over-expressing lines,

indicating that the action of ABHD1 does not significantly affect the major flux of acyl chains into TAG during N-starvation.

We explored potential proteomics changes in the isolated LDs (Fig. S9, Data Set S1). Among the 598 proteins repeatedly detected by mass spectrometry (MS)-based proteomics, only four showed a significant change to the relative amount between the WT and the two mutants (Data Set S1). These four proteins were reduced in the knockout mutant LDs: ABHD1 (as expected), the long-chain acyl-CoA synthetase (LCS2), the photosystem I reaction center subunit H (PSAH), and a putative glycosyl hydrolase (GHL1). The reduction in the LCS2 protein amount in the two knockout mutants supports the idea that in the mutant there is indeed a reduced need for acyl-activation, corroborating the enzymology data below showing that ABHD1 is a lipase (next section). GHL1 has been reported to potentially function as a galactolipid galactosyl hydrolase and provide DAG for TAG synthesis [51], but it remains to be tested whether GHL1 participates in glyceryl-*N*,*N*,*N*-trimethylhomoserine (GTS) hydrolysis, allowing, therefore, glycerol recycling downstream of ABHD1. Taken together, these data suggest that ABHD1 may act in the modification of the monolayer of the LD coat, as its absence results in changes in both LD membrane lipid and protein composition.

## ABHD1 has lyso-DGTS lipase activity

To investigate the potential enzymatic activity of ABHD1, we expressed in *Escherichia coli* a truncated version of ABHD1 lacking the N-terminal hydrophobic portion (with amino acids 1 to 46 removed) resulting in recombinant ABHD1 (rABHD1). Purification of the soluble native rABHD1 was only partial but, in the presence of urea, a highly pure rABHD1 protein was obtained from inclusion bodies and refolded (Fig. 4A, Table S3). To identify its potential substrate(s), the refolded purified rABHD1 was first incubated with total lipid extracts of *Chlamydomonas abhd1-1* mutant. Interestingly, we observed a reduction in lyso-DGTS species and an increase in free fatty acids, but no change in the other major lipid molecular species (Fig. 4B, Fig. S10A). We could also detect changes in some other very minor lysolipid species (such as lyso-MGDG, lyso-SQDG, lyso-PG and lyso-PE) but not lyso-DGDG (Fig. S10B). These other lysolipids were not further tested because they are >100 times less abundant than lyso-DGTS. ABHD1 activity was then tested using pure lyso-DGTS as substrate, prepared from the partial spontaneous hydrolysis of commercial
<figure><img src="image_6.png" /><figcaption>**Figure 4.** Enzyme activity analysis of the refolded rABHD1 protein. (A) SDS-PAGE of rABHD1 purified from *E. coli* inclusions bodies and refolded. (B) Activity test of rABHD1 on *abhd1-1* total lipid extracts identified lyso-DGTS as potential ABHD1 substrate. LC-MS detected over 10 000 m/z peaks for both the reactions with and without rABHD1. The full spectrum from LC-MS is provided in Fig. S10. (C) The lipase activity of rABHD1 toward purified lyso-DGTS. (D) The lipase activity of rABHD1 toward commercial lyso-PC. (E) Activity assay on lyso-PE, lyso-PG, MAG, PC and DGTS. But, no activity could be detected on PC or on DGTS for rABHD1. Data are the mean of three independent experiments and error bars refer to standard deviation. Student's t-test is applied after passing Bartlett's test for homoscedasticity and Shapiro-Wilk's test for normality: \*\*P < 0.01, \*\*\*P < 0.001. Free fatty acid (FFA) formation was quantified using 19:0 fatty acid as an internal standard. Reaction conditions for all activity tests: Teorell Stenhagen universal buffer pH 7.5, with 100 mM NaCl and Triton-X100 at 1 CMC. During the enzymatic reaction (incubation of 2 h), during the preparation of lysolipid substrate or during lipid extraction, there is always some autolysis of the substrate, which is responsible for the presence of a certain amount of free fatty acids in the negative control.</figcaption></figure>

DGTS. Results showed that ABHD1 indeed acted as a lipase on pure lyso-DGTS, as the two expected co-products of the reaction (Fig. 4C), palmitic acid and GTS, could be identified by MS (Fig. S11).

Similar activity could be obtained using lyso-phosphatidylcholine (lyso-PC), a commercially available lipid structurally similar to lyso-DGTS (Fig. 4D). Due to the difficulty in obtaining purified lyso-DGTS, further characterization of the putative enzymatic activity of rABHD1 was carried out on lyso-PC. Activity assays conducted under various conditions of buffer, pH, detergent or cations (Fig. S12) allowed us to select a reaction medium for further studies. Release of free

fatty acid from lyso-PC was found to increase with time and with rABHD1 amount, which showed that it corresponded to genuine enzymatic activity (Fig. S13).

ABHD1 did not show activity toward lipids with two acyl-groups (Fig. 4B, Table S4), therefore we tested the activity of rABHD1 toward other lipids containing one fatty acid, e.g. lyso-phosphatidylethanolamine (lyso-PE), lyso-phosphatidylglycerol (lyso-PG) and monoacylglycerol (MAG). Results showed that rABHD1 protein released fatty acids from lyso-PE, MAG and lyso-PG but showed relatively low activity toward the latter (Fig. 4E). This was observed not only when incubated with total lipid extracts, but also purified commercial substrates as well (Fig. 4D and E). These results thus indicated that rABHD1 has some preference for lysolipids over diacyl-lipid analogs (no activity toward lipids with two acyl-groups as shown in Fig. 4B), and it did not show any acyltransferase or transacylase activity (Table S4).

ABHD1 favors LD budding in vitro

We further investigated the effect of ABHD1 association and lysolipid hydrolysis on the geometry of LDs. To do so, we employed the droplet-embedded vesicle (DEV) system [52,53]. Model LDs are incorporated into pre-formed giant unilamellar vesicles (GUVs) producing DEVs (Fig. 5A). We then deposited DEVs on a glass coverslip which caused them to rupture (Fig. 5B), resulting in a 2D bilayer with the droplets incorporated [54]. Resulting LDs were immobilized and their geometry was followed throughout. The phospholipid composition was as follows: dioleoyl phosphatidylcholine (DOPC)/lyso-PC/rhodamine-PE (79.5/20/0.5% w/w), and TAG to form neutral LDs. Fluorescent lipid rhodamine-PE acted as a membrane reporter and nitrobenzoxadiazole (NBD)-labeled TAG allowed us to visualize the LD. The model LDs' initial shape is a spherical cap, and addition of the purified rABHD1 resulted in the transition of the model LD from a flattened to a budded shape. This transition is shown by the fact that in the lipid bilayer the projected droplet surface area decreased with time by 40% as compared to the control, i.e. without any protein added (Fig. 5C and D). Moreover, we obtained similar results with pure DOPC DEVs, i.e. in the absence of lyso-PC, the major substrates of ABHD1 (Fig. 5E and F). Furthermore, when a non-refolded urea-purified form of rABHD1, which was enzymatically completely inactive (see Material and Methods), was tested in the DEV system, budding was still observed despite having slightly weaker capacity in comparison to the wild-type version of
<figure><img src="image_7.png" /><figcaption>**Figure 5.** Addition of the rABHD1 protein to the GUV boosts curvature formation. (A) Schematic view of the experimental protocol. GUVs are put in contact with a thin emulsion of NBD-labeled triacylglycerol (TAG) that forms droplets at the lipid bilayer. When placed on a glass coverslip, membrane rupture results in a 2D lipid bilayer at the surface with droplets of TAG incorporated in. Right: typical image of a DEV after rupture is shown with Rho-PE in red and TAG NBD in green. Scale bar is 10 µm. (B) The droplet initially forms a spherical cap and is budding from the membrane 45 min after rABHD1 addition in the observation buffer (100 µg mL⁻¹). (C) Time lapse of a TAG droplet at the membrane plane (NBD TAG signal in upper panel and bright field in lower panel), showing the decrease of the surface after addition of rABHD1. Scale bars are 2 µm. (D) Control time lapse of a TAG droplet at the membrane plane without addition of protein. Scale bars are 2 µm. (E, F) Measurement of the droplet surface at the membrane plane according to time in two different lipid compositions: (E) DOPC/Lyso-PC/RhoPE and (F) DOPC/RhoPE. The measured area of each droplet is normalized by the initial surface occupied at t = -5 min. Dashed lines represent the time traces of individual droplets and thick lines are the average trend. Light yellow lines correspond to the control in the absence of ABHD1 [(E) N = 3 droplets analyzed, (F) N = 4], and dark red lines to the addition of rABHD1 at 100 µg mL⁻¹ [(E) N = 16, (F) N = 13].</figcaption></figure>

the protein (Fig. S14A). The fact that the denatured rABHD1 had weaker budding than the refolded rABHD1 may be due to the harsh denaturation conditions used, which may have impacted negatively (directly or indirectly) the rABHD1 domain involved in the budding. Addition of bovine serum albumin (BSA) instead of ABHD1 protein did not result in LD budding, providing further support that the above observation is specific to ABHD1 (Fig. S14B). Taken together, the above two lines of evidence clearly show that, independently of its lyso-DGTS lipase function, the binding of ABHD1 to the model LDs' surface participates in droplet budding [S2].

## DISCUSSION

Although the proteome of *Chlamydomonas* LDs was published more than 10 years ago [36,37], only a few LD-associated proteins have been studied in any detail, and none have a demonstrated role in LD biogenesis. Here, using a combination of bio-physical, cell-biological, genetic and lipidomic approaches, we show that the α/β hydrolase domain-containing protein ABHD1 acts as a lysolipid lipase and participates in LD biogenesis. Thus far, some lysolipid acyltransferases have been studied in algae, but no lysolipid lipase has yet been identified. We demonstrate here that ABHD1 catalyzes the hydrol-ysis of an acyl group from a lyso-DGTS molecule, and it is therefore a characterization of a new enzyme acting as a lyso-DGTS lipase, and the first lysolipid lipase identified among algae. Our current work-ing model is that ABHD1 promotes LD emergence likely through two actions at the LD surface: a lyso-DGTS lipase activity, and a distinct non-enzymatic property that promotes LD emergence, probably al-tering the metabolic fate of DGTS. We provide a model of molecular events involving ABHD1 that may lead to LD emergence (Fig. 6) and we then discuss the possible implications of our findings for DGTS and TAG metabolism.

## ABHD1 is a lyso-DGTS lipase acting on the LD surface

ABHD1 belongs to the α/β-hydrolase fold domain-containing superfamily, one of the largest protein superfamilies, present in virtually all sequenced genomes [S5]. Despite their large number and wide occurrence, the physiological substrates and prod-ucts for members of the ABHD family remain largely uncharacterized [S5]. Here, through *in vivo* and *in vitro* experiments, we provide firm evidence that ABHD1 is located at the LD surface (Fig. 1) and is a novel type of lipase, namely a lyso-DGTS li-pase (Fig. 3). ABHD1 is indeed shown to be active *in vitro* on lyso-glycerolipids, including lyso-DGTS, but not on diacyl-glycerolipids such as DGTS. Similar to some other members of its superfam-ily, ABHD1 contains the lipase catalytic triad, i.e. Asp179, Asp328 and His356 (Fig. S2C). Further-more, the orientation of the protein on the LD sur-face could be favored by the C-terminal hydropho-bic helix (Fig. S2B), bringing the putative active site of the enzyme closer to the LD surface.

## Lysolipids and biophysical properties of ABHD1 contribute to LD budding

Independent of its enzymatic function, ABHD1 is also found to play a structural role in favoring LD
<figure><img src="image_8.png" /><figcaption>**Figure 6.** A model explaining the multifaceted actions of lyso-DGTS and ABHD1 protein in promoting LD budding, growth and TAG synthesis. ABHD1 catalyzes the hydrolysis of lyso-DGTS to produce a free fatty acid and a GTS moiety. ABHD1 is present at basal levels during optimal growth, and it increases in transcription and protein amount as N starvation is initiated, paralleling the increases in both LD number and size [67]. During LD budding, the formation of lyso-DGTS and the synthesis of ABHD1 protein both promote positive membrane curvature and therefore LD budding. Later on, ABHD1 activity may facilitate LD growth as less curvature is needed by degrading lyso-DGTS. The resulting fatty acid (FA) can be activated to acyl-CoA by LCS2 and contribute to TAG synthesis. Additionally, by consuming lyso-DGTS, ABHD1 is pulling the flow of acyls from either a putative PDAT or a PLA2, whose activity drives either TAG or FA production, respectively. Indeed, in two knockout mutants, LCS2 protein is present in a reduced amount, likely because the flow of free FA is reduced without ABHD1. Taken together, the major role of ABHD1 lies in LD coat remodeling and maintaining LD stability. Abbreviations: BTA1, betaine lipid synthase 1; DAG, diacylglycerol; DGAT, diacylglycerol acyltransferase; DGTS, diacylglyceryl-3-O-4'-(N,N,N-trimethyl)-homoserine; FAS, fatty acid synthase; G3P, glycerol-3-phosphate; GPAT, glycerol-3-phosphate acyltransferase; MLDP, major lipid droplet protein; LCS2, long chain acyl-CoA synthetase; Lyso-PA, lysophosphatidic acid; LPLAT, lysophospholipid acyltransferase; PA, phosphatidic acid; PDAT, phospholipid: diacylglycerol acyltransferase; PLA2, phospholipase A2; TAG, triacylglycerol.</figcaption></figure>

budding and growth based on both *in vivo* and *in vitro* experiments (Figs 2 and 5). Lipidomics of isolated LDs from the two knock-out mutants show an increase in lyso-DGTS. Lysolipids contain a big polar head and only one acyl chain, and in general are present as hexagonal tubes I ($H_1$) and form positively curved monolayers [56,57]. This property could be important for the budding process because positive curvature forms sharp angles. Indeed, a higher amount of lyso-DGTS was present in LDs of the two mutants. We therefore propose that at the initiation of LD formation, DGTS is converted to a lyso-DGTS, which is important for the facilitation of membrane budding. In addition, a new study has recently reported that lyso-DGTS enhances the activity of PON1 (Paraoxonase 1), and by so doing prevents oxidation of low-density lipoproteins (LDL) [58]. This study therefore points out an additional role for lyso-DGTS, i.e. boosting enzyme activity by perhaps changing LD biophysics.

Nevertheless, the accumulation of lysolipids is deleterious to the ER membrane integrity [59]. Therefore, while the generation of lyso-DGTS from DGTS may facilitate LD nucleation and budding,

it will be crucial to prevent the accumulation of lysolipids. This could be done by balancing the actions of DGTS-to-lyso-DGTS conversion and lyso-DGTS degradation (Fig. 6). ABHD1 may mediate the specific degradation of lyso-DGTS (and possibly other lysolipids of the LD) by releasing free fatty acids from lysolipids. These fatty acids are then possibly used to make TAGs. A question remains as to why this reaction occurs on LDs: very likely because the LD surface offers more accessibility to lipids, as it has more packing defects [8,60], which might grant better access to lyso-DGTS and their hydrolysis/detoxification. Knowing that the ER and LDs are in contiguity, lysolipids can diffuse between these organelles, and their degradation at LDs would behave as a thermodynamic pump, driving lyso-DGTS from the ER-to-LDs for their continuous degradation.

## DGTS metabolism and TAG synthesis at the LD surface

*Chlamydomonas* lacks PC and contains instead the betaine lipid DGTS as its major extra-chloroplast membrane lipid covering the LD surface [37].
Compared to the large variety of proteins known
to be involved in acylglycerol lipid metabolism,
only one protein (i.e. BTA1) of betaine lipid
metabolism has so far been studied, first in *Rhodobac-
ter sphaeroides* [61], then in *Chlamydomonas* [40],
and recently in *Nannochloropsis oceanica* [62]. BTA1
catalyzes the addition of S-adenosylmethionine to
diacylglycerol (DAG) forming DGTS [40]. Re-
cently, it has been shown that DGTS remodeling
contributes to TAG synthesis during ER stress and
this process is regulated by the transcription factor
bZIP1 in *Chlamydomonas* [25]. Pulse-chase labeling
experiments revealed that DGTS turnover could
provide acyl groups for TAG biosynthesis, however
at a much lower level than MGDG or DGDG,
similar to other previous results in *Chlamydomonas*
[63,64]. This explains the lack of flux of ¹⁴C-acyl
groups from DGTS to TAG in the *abhd1* mutants,
consistent with the finding that its role is not in the
major TAG biosynthetic pathway, but rather in the
DGTS remodeling at the LD surface. Interestingly,
BTA1 together with ABHD1 is found to be among
the top 10 most abundant proteins in LDs from WT
cells (Data Set S1), indicating that the LD surface
provides a platform for active DGTS synthesis and
turnover that may be separated from the roles of
DGTS in the ER membrane. The role of ABHD1 in
LD coat remodeling and TAG synthesis is further
supported by the changes in protein compositions
observed in the two knock-out mutants (Fig. 6).

Taken together, the synthesis and degradation of
lyso-DGTS at the LD surface indicates that ABHD1
plays a dual role to initiate LD formation and en-
sure LD growth, important in adapting membrane
biophysical properties as well as in supplying acyl-
chains for TAG synthesis. The fact that the knock-
out mutants (abhd1-1 or abhd1-2) did not show dif-
ferences in their TAG content suggests that ABHD1
does not constitute a major flux in supplying acyl-
chains for TAG synthesis, but rather it is important
in membrane lipid coat remodeling and in prepara-
tion of biophysical membrane properties essential
for LD emergence and assembly. Over-expression
of the ABHD1 protein during N-replete optimal
growth boosted LD formation and oil content in
*Chlamydomonas* but also in yeast (Fig. 2), suggesting
that ABHD1 could be used to increase TAG content
of microalgae, and possibly other organisms, without
compromising/halting cell growth.

MATERIALS AND METHODS

Strains and culture conditions

All *Chlamydomonas* strains were maintained in agar
plates containing Tris-Acetate-Phosphate (TAP)

media [65] under constant light at 25°C. In the case
of mutants, TAP agar plates were supplemented
with 15 µg mL⁻¹ hygromycin or 10 µg mL⁻¹ paro-
momycin as appropriate. Cell cultures were grown
in TAP liquid media in conical glass flasks kept in in-
cubators (Multitron, Infors HT) shaking at 120 rpm,
25°C, under continuous light (80–100 µmol pho-
tons m⁻² s⁻¹). Cell concentration, size and volume
were quantified with a Multisizer 4 (Beckman
Coulter). To start N deprivation, log-phase cultures
were washed with fresh media without N (NH₄Cl
replaced by an equal molar amount of NaCl) at 450 g
for 3 min.

DATA AVAILABILITY

All data are available in the main text or the supplementary
materials. The MS proteomics data have been deposited in the
ProteomeXchange Consortium via the Proteomics Identifica-
tion Database (PRIDE) [66] partner repository, with the data
set identifier PXD036778. All other methods are described in
*Supplementary Materials and Methods*.

SUPPLEMENTARY DATA

Supplementary data are available at *NSR* online.

ACKNOWLEDGEMENTS

We thank Stéphanie Blangy and Pascaline Auroy in providing
guidance on protein expression and molecular biology work re-
spectively. We also thank Ousmane Dao in helping with culture
preparations and Pascale David for verifying some constructions.
We thank Audam Chhun for helping with graphic design. We
thank the ZoOM Microscopy facility (BIAM, CEA Cadarache)
and the Chlamydomonas Resource Center at the University of
Minnesota for providing the indexed *C. reinhardtii* insertional mu-
tants.

FUNDING

This work was supported by the European Union Regional De-
veloping Fund (ERDF), the Région Provence Alpes Côte d'Azur,
the French Ministry of Research and the French Alternative En-
ergies and Atomic Energy Commission (CEA) by way of fund-
ing the HelioBiotec platform. This work was also partly supported
by Fundamental Research Division (DRF) Impulsion LD power.
I.T.R. is the recipient of an international PhD studentship from
CEA (program Irtelis). A.R.T. is supported by ANR-21-CE11-
0032-02-LIPRODYN. M.G.G., H.K.K. and P.D.B. are supported
by the National Science Foundation (PGRP-IOS-1829365). The
proteomic experiments were partially supported by Agence Na-
tionale de la Recherche under projects ProFI (Proteomics French
Infrastructure, ANR-10-INBS-08) and GRAL (Grenoble Alliance
for Integrated Structural & Cell Biology), a program from the
Chemistry Biology Health (CBH) Graduate School of University
Grenoble Alpes (ANR-17-EURE-0003). We are grateful for the
financial support of a collaborative project between the Chinese
Academy of Sciences and CEA.
## AUTHOR CONTRIBUTIONS

I.T.R., B.L., F.B. and Y.L.B. planned and designed the research. I.T.R. and M.H. performed the subcellular localization and studied the insertional knockout mutants. I.T.R., B.L., F.V., C.B. and D.S. performed in vitro protein production and activity analysis. I.T.R. and S.C. purified LDs. I.T.R., M.H. and B.L. performed lipid analysis. S.B. and Y.C. carried out proteomics. Y.X. and J.X. performed yeast expression analysis. M.G.G., H.K.K., P.B. performed pulse-chase labeling work. A.D. and A.R.T. performed in vitro model droplet assays. Y.L.B., F.B., I.T.R. and A.R.T. wrote the manuscript.

*Conflict of interest statement.* None declared.

## REFERENCES

1. Huang AHC. Oil bodies and oleosins in seeds. *Annu Rev Plant Physiol Plant Mol Biol* 1992; **43**: 177–200.

2. Goold H, Beisson F, Peltier G *et al*. Microalgal lipid droplets: composition, diversity, biogenesis and functions. *Plant Cell Rep* 2015; **34**: 545–55.

3. Chapman KD, Dyer JM, Mullen RT. Biogenesis and functions of lipid droplets in plants: thematic review series: lipid droplet synthesis and metabolism: from yeast to man. *J Lipid Res* 2012; **53**: 215–26.

4. Nguyen TB and Olzmann JA. Lipid droplets and lipotoxicity during autophagy. *Autophagy* 2017; **13**: 2002–3.

5. Xu C and Fan J. Links between autophagy and lipid droplet dynamics. *J Exp Bot* 2022; **73**: 2848–58.

6. Thiam AR, Farese RV, Walther TC. The biophysics and cell biology of lipid droplets. *Nat Rev Mol Cell Biol* 2013; **14**: 775–86.

7. Chorlay A, Monticelli L, Veríssimo Ferreira J *et al*. Membrane asymmetry imposes directionality on lipid droplet emergence from the ER. *Dev Cell* 2019; **50**: 25–42.

8. Chorlay A and Thiam AR. Neutral lipids regulate amphipathic helix affinity for model lipid droplets. *J Cell Biol* 2020; **219**: e201907099.

9. Farese RV Jr and Walther TC. Lipid droplets finally get a little R-E-S-P-E-C-T. *Cell* 2009; **139**: 855–60.

10. Szymanski KM, Binns D, Bartz R *et al*. The lipodystrophy protein seipin is found at endoplasmic reticulum lipid droplet junctions and is important for droplet morphology. *Proc Natl Acad Sci USA* 2007; **104**: 20890–5.

11. Valm AM, Cohen S, Legant WR *et al*. Applying systems-level spectral imaging and analysis to reveal the organelle interac-tome. *Nature* 2017; **546**: 162–7.

12. de Vries J and Ischebeck T. Ties between stress and lipid droplets pre-date seeds. *Trends Plant Sci* 2020; **25**: 1203–14.

13. Dadras A, Fürst-Jansen JMR, Darienko T *et al*. Environmental gradients reveal stress hubs pre-dating plant terrestrialization. *Nat Plants* 2023; **9**: 1419–38.

14. Xu N, Zhang SO, Cole RA *et al*. The FATP1-DGAT2 complex facilitates lipid droplet expansion at the ER-lipid droplet interface. *J Cell Biol* 2012; **198**: 895–911.

15. Cai Y, Goodman JM, Pyc M *et al*. Arabidopsis SEIPIN proteins modulate triacylglycerol accumulation and influence lipid droplet proliferation. *Plant Cell* 2015; **27**: 2616–36.

16. Adeyo O, Horn PJ, Lee SK *et al*. The yeast lipin orthologue Pah1p is important for biogenesis of lipid droplets. *J Cell Biol* 2011; **192**: 1043–55.

17. Horn PJ, James CN, Gidda SK *et al*. Identification of a new class of lipid droplet-associated proteins in plants. *Plant Physiol* 2013; **162**: 1926–36.

18. Yang HY, Galea A, Sytnyk V *et al*. Controlling the size of lipid droplets: lipid and protein factors. *Curr Opin Cell Biol* 2012; **24**: 509–16.

19. Pyc M, Cai Y, Gidda SK *et al*. Arabidopsis lipid droplet-associated protein (LDAP)—interacting protein (LDIP) influences lipid droplet size and neutral lipid homeostasis in both leaves and seeds. *Plant J* 2017; **92**: 1182–201.

20. Pyc M, Gidda SK, Seay D *et al*. LDIP cooperates with SEIPIN and LDAP to facilitate lipid droplet biogenesis in Arabidopsis. *Plant Cell* 2021; **33**: 3076–103.

21. Chung J, Wu X, Lambert TJ *et al*. LDAF1 and Seipin form a lipid droplet assembly complex. *Dev Cell* 2019; **51**: 551–63.

22. Greer MS, Cai Y, Gidda SK *et al*. SEIPIN isoforms interact with the membrane-tethering protein VAP27-1 for lipid droplet formation. *Plant Cell* 2020; **32**: 2932–50.

23. Miklaszewska M, Zienkiewicz K, Klugier-Borowska E *et al*. CA-LEOSIN 1 interaction with AUTOPHAGY-RELATED PROTEIN 8 facilitates lipid droplet microautophagy in seedlings. *Plant Physiol* 2023; **193**: 2361–80.

24. Siaut M, Cuiné S, Cagnon C *et al*. Oil accumulation in the model green alga Chlamydomonas reinhardtii: characterization, variability between common laboratory strains and relationship with starch reserves. *BMC Biotechnol* 2011; **11**: 7.

25. Yamaoka Y, Shin S, Choi BY *et al*. The bZIP1 transcription factor regulates lipid remodeling and contributes to ER stress management in Chlamydomonas reinhardtii. *Plant Cell* 2019; **31**: 1127–40.

26. Yoon K, Han D, Li Y *et al*. Phospholipid:diacylglycerol acyl-transferase is a multifunctional enzyme involved in membrane lipid turnover and degradation while synthesizing triacylglycerol in the unicellular green microalga Chlamydomonas reinhardtii. *Plant Cell* 2012; **24**: 3708–24.

27. Bai F, Yu L, Shi J *et al*. Long-chain acyl-CoA synthetases activate fatty acids for lipid synthesis, remodeling and energy production in Chlamydomonas. *New Phytol* 2022; **233**: 823–37.

28. Tsai C-H, Warakanont J, Takeuchi T *et al*. The protein Compro-mised Hydrolysis of Triacylglycerols 7 (CHT7) acts as a repressor of cellular quiescence in Chlamydomonas. *Proc Natl Acad Sci USA* 2014; **111**: 15833–8.

29. Jang S, Kong F, Lee J *et al*. CrABCA2 facilitates triacylglyc-erol accumulation in Chlamydomonas reinhardtii under nitrogen starvation. *Mol Cells* 2020; **43**: 48–57.

30. Kong F, Liang Y, Legeret B *et al*. Chlamydomonas carries out fatty acid beta-oxidation in ancestral peroxisomes using a bona fide acyl-CoA oxidase. *Plant J* 2017; **90**: 358–71.

31. Kong F, Burlacot A, Liang Y *et al*. Interorganelle communica-tion: peroxisomal MALATE DEHYDROGENASE 2 connects lipid catabolism to photosynthesis through redox coupling in Chlamy-domonas. *Plant Cell* 2018; **30**: 1824–47.
32. Goold HD, Cuine S, Legeret B *et al.* Saturating light induces sustained accumulation of oil in plastidal lipid droplets in Chlamydomonas reinhardtii. *Plant Physiol* 2016; **171**: 2406–17.

33. Kajikawa M, Sawaragi Y, Shinkawa H *et al.* Algal dual-specificity tyrosine phosphorylation-regulated kinase, triacylglycerol accumulation regulator1, regulates accumulation of triacylglycerol in nitrogen or sulfur deficiency. *Plant Physiol* 2015; **168**: 752–64.

34. Merchant SS, Kropat J, Liu B *et al.* TAG, you’re it! Chlamydomonas as a reference organism for understanding algal triacylglycerol accumulation. *Curr Opin Biotechnol* 2012; **23**: 352–63.

35. Lee J, Yamaoka Y, Kong F *et al.* The phosphatidylethanolamine-binding protein DTH1 mediates degradation of lipid droplets in Chlamydomonas reinhardtii. *Proc Natl Acad Sci USA* 2020; **117**: 23131–9.

36. Moellering ER and Benning C. RNA interference silencing of a major lipid droplet protein affects lipid droplet size in Chlamydomonas reinhardtii. *Euk Cell* 2010; **9**: 97–106.

37. Nguyen HM, Baudet M, Cuiné S *et al.* Proteomic profiling of oil bodies isolated from the unicellular green microalga Chlamydomonas reinhardtii: with focus on proteins involved in lipid metabolism. *Proteomics* 2011; **11**: 4266–73.

38. Tsai C-H, Zienkiewicz K, Amstutz CL *et al.* Dynamics of protein and polar lipid recruitment during lipid droplet assembly in Chlamydomonas reinhardtii. *Plant J* 2015; **83**: 650–60.

39. Wang X, Wei H, Mao X *et al.* Proteomics analysis of lipid droplets from the oleaginous alga Chromochloris zofingiensis reveals novel proteins for lipid metabolism. *Genom Proteomics Bioinformatics* 2019; **17**: 260–72.

40. Riekhof WR, Sears BB, Benning C. Annotation of genes involved in glycerolipid biosynthesis in Chlamydomonas reinhardtii: discovery of the betaine lipid synthase BTA1(Cr). *Euk Cell* 2005; **4**: 242–52.

41. Li X, Zhang R, Patena W *et al.* An indexed, mapped mutant library enables reverse genetics studies of biological processes in Chlamydomonas reinhardtii. *Plant Cell* 2016; **28**: 367–87.

42. Bai F, Yu L, Shi J *et al.* Long-chain acyl-CoA synthetases activate fatty acids for lipid synthesis, remodeling and energy production in Chlamydomonas. *New Phytol* 2022; **233**: 823–37.

43. Warakanont J, Tsai C-H, Michel EJS *et al.* Chloroplast lipid transfer processes in Chlamydomonas reinhardtii involving a TRIGALACTOSYLDIACYLGLYCEROL 2 (TGD2) orthologue. *Plant J* 2015; **84**: 1005–20.

44. Zheng H-Q, Chiang-Hsieh Y-F, Chien C-H *et al.* AlgaePath: comprehensive analysis of metabolic pathways using transcript abundance data from next-generation sequencing in green algae. *BMC Genomics* 2014; **15**: 196.

45. Moellering ER and Benning C. RNA interference silencing of a major lipid droplet protein affects lipid droplet size in Chlamydomonas reinhardtii. *Euk Cell* 2010; **9**: 97–106.

46. Nguyen HM, Baudet M, Cuiné S *et al.* Proteomic profiling of oil bodies isolated from the unicellular green microalga Chlamydomonas reinhardtii: with focus on proteins involved in lipid metabolism. *Proteomics* 2011; **11**: 4266–73.

47. Kyte J and Doolittle RF. A simple method for displaying the hydropathic character of a protein. *J Mol Biol* 1982; **157**: 105–32.

48. Jumper J, Evans R, Pritzel A *et al.* Highly accurate protein structure prediction with AlphaFold. *Nature* 2021; **596**: 583–9.

49. Nagai Y, Aoki J, Sato T *et al.* An alternative splicing form of phosphatidylserine-specific phospholipase A1 that exhibits lysophosphatidylserine-specific lysophospholipase activity in humans. *J Biol Chem* 1999; **274**: 11053–9.

50. Sandager L, Gustavsson MH, Ståhl U *et al.* Storage lipid synthesis is non-essential in yeast. *J Biol Chem* 2002; **277**: 6478–82.

51. Gu X, Cao L, Wu X *et al.* A lipid bodies-associated galactosyl hydrolase is involved in triacylglycerol biosynthesis and galactolipid turnover in the unicellular green alga Chlamydomonas reinhardtii. *Plants (Basel)* 2021; **10**: 675.

52. Chorlay A, Monticelli L, Ferreira JV *et al.* Membrane asymmetry imposes directionality on lipid droplet emergence from the ER. *Dev Cell* 2019; **50**: 25–42.

53. Chorlay A, Santinho A, Thiam AR. Making droplet-embedded vesicles to model cellular lipid droplets. *STAR Protoc* 2020; **1**: 100116.

54. Salo VT, Li S, Vihinen H *et al.* Seipin facilitates triglyceride flow to lipid droplet and counteracts droplet ripening via endoplasmic reticulum contact. *Dev Cell* 2019; **50**: 478–93.

55. Lord CC, Thomas G, Brown JM. Mammalian alpha beta hydrolase domain (ABHD) proteins: lipid metabolizing enzymes at the interface of cell signaling and energy metabolism. *Biochim Biophys Acta Mol Cell Biol Lipids* 2013; **1831**: 792–802.

56. Fuller N and Rand RP. The influence of lysolipids on the spontaneous curvature and bending elasticity of phospholipid membranes. *Biophys J* 2001; **81**: 243–54.

57. Jouhet J. Importance of the hexagonal lipid phase in biological membrane organization. *Front Plant Sci* 2013; **4**: 494.

58. Khattib A, Musa S, Halabi M *et al.* Lyso-DGTS lipid derivatives enhance PON1 activities and prevent oxidation of LDL: a structure–activity relationship study. *Antioxidants* 2022; **11**: 2058.

59. Ben M’barek K, Ajjaji D, Chorlay A *et al.* ER membrane phospholipids and surface tension control cellular lipid droplet formation. *Dev Cell* 2017; **41**: 591–604.

60. Chorlay A, Forêt L, Thiam AR. Origin of gradients in lipid density and surface tension between connected lipid droplet and bilayer. *Biophys J* 2021; **120**: 5491–503.

61. Klug RM and Benning C. Two enzymes of diacylglyceryl-O-4’-(N,N,N’-trimethyl)homoserine biosynthesis are encoded by btaA and btaB in the purple bacterium Rhodobacter sphaeroides. *Proc Natl Acad Sci USA* 2001; **98**: 5910–5.

62. Murakami H, Nobusawa T, Hori K *et al.* Betaine lipid is crucial for adapting to low temperature and phosphate deficiency in nannochloropsis. *Plant Physiol* 2018; **177**: 181–93.

63. Li X, Moellering ER, Liu B *et al.* A galactoglycerolipid lipase is required for triacylglycerol accumulation and survival following nitrogen deprivation in Chlamydomonas reinhardtii. *Plant Cell* 2012; **24**: 4670–86.

64. Young DY and Shachar-Hill Y. Large fluxes of fatty acids from membranes to triacylglycerol and back during N-deprivation and recovery in Chlamydomonas. *Plant Physiol* 2021; **185**: 796–814.

65. Harris EH. *The Chlamydomonas Sourcebook: Introduction to Chlamydomonas and Its Laboratory Use: Volume 1*. Amsterdam: Elsevier Science, 2009.

66. Perez-Riverol Y, Csordas A, Bai J *et al.* The PRIDE database and related tools and resources in 2019: improving support for quantification data. *Nucleic Acids Res* 2019; **47**: D442–50.

67. Miller R, Wu G, Deshpande RR *et al.* Changes in transcript abundance in Chlamydomonas reinhardtii following nitrogen deprivation predict diversion of metabolism. *Plant Physiol* 2010; **154**: 1737–52.