# Betaine lipids overproduced in seed plants are not imported into plastid membranes and promote endomembrane expansion

Sarah Salomon, Marion Schilling, Catherine Albrieux, Grégory Si Larbi, Pierre-Henri Jouneau, Sylvaine Roy, Denis Falconet, Morgane Michaud, Juliette Jouhet

► To cite this version:

Sarah Salomon, Marion Schilling, Catherine Albrieux, Grégory Si Larbi, Pierre-Henri Jouneau, et al.. Betaine lipids overproduced in seed plants are not imported into plastid membranes and promote endomembrane expansion. Journal of Experimental Botany, 2025, 76 (4), pp.980-996. 10.1093/jxb/erae458. hal-04868681

HAL Id: hal-04868681

https://hal.science/hal-04868681v1

Submitted on 6 Jan 2025

HAL is a multi-disciplinary open access
archive for the deposit and dissemination of sci-
entific research documents, whether they are pub-
lished 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_1.png" /></figure>

Distributed under a Creative Commons Attribution 4.0 International License
RESEARCH PAPER

# Betaine lipids overproduced in seed plants are not imported into plastid membranes and promote endomembrane expansion

Sarah Salomon<fnref n="1" />, Marion Schilling<fnref n="1" />, ID, Catherine Albrieux<fnref n="1" />, Grégory Si Larbi<fnref n="1" />, ID, Pierre-Henri Jouneau<fnref n="2" />, Sylvaine Roy<fnref n="1" />, ID, Denis Falconet<fnref n="1" />, ID, Morgane Michaud<fnref n="1" />,* ID, and Juliette Jouhet<fnref n="1" />,* ID

Received 1 October 2024; Editorial decision 31 October 2024; Accepted 10 November 2024

Editor: Oswaldo Valdés-López, Universidad Nacional Autonoma de Mexico, Mexico

## Abstract

Plants and algae have to adapt to environmental changes and face various stresses that negatively affect their growth and development. One common stress is phosphate (Pi) deficiency, which is often present in the environment at limiting levels. In response to Pi deficiency, these organisms increase Pi uptake and remobilize intracellular Pi. Phospholipids are degraded to provide Pi and are replaced by non-phosphorus lipids, such as glycolipids or betaine lipids. During evolution, seed plants lost the ability to synthesize betaine lipids. By expressing *Bta1* genes, which are involved in the synthesis of diacylglyceryl-*N*,*N*,*N*-trimethyl-homoserine (DGTS), from different species, we showed that DGTS can be produced in seed plants. In Arabidopsis, expression of BTA1 under a phosphate starvation-inducible promoter resulted in limited DGTS production without having any impact on plant growth or lipid remodelling. In transient expression systems in *Nicotiana benthamiana*, leaves were able to accumulate DGTS to up to 30% of their glycerolipid content at a slight expense to galactolipid and phospholipid production. At the subcellular level, we showed that DGTS is absent from plastids and seems to be enriched in endomembranes, inducing endoplasmic reticulum membrane proliferation. Finally, the DGTS synthesis pathway seems to compete with phosphatidylcholine (PC) synthesis via the Kennedy pathway but does not appear to be derived from the PC diacylglycerol backbone and therefore does not interfere with the eukaryotic pathway involved in galactolipid synthesis.

**Keywords:** Betaine lipid, chloroplast, endomembrane, phosphate starvation, phospholipid, seed plants.

## Introduction

Plants and algae are organisms that must constantly adapt to changes in their environment and, consequently, face numerous stresses such as nutrient deficiency, water scarcity, temperature fluctuations, or attacks from various pathogens. These stresses can negatively affect the growth and development of these organisms, thereby significantly impacting raw material

production yields. Phosphate (Pi) deficiency is one of the most common nutrient stresses, leading to yield reductions ranging from 25% to 60% in various crops (Nelson and Janke, 2007; Smit et al., 2009; Vitousek et al., 2009). Pi is an essential nutrient for the survival of all living organisms. Plants and algae acquire the Pi they need for growth and development from the

<footnote n="1">¹ Laboratoire Physiologie Cellulaire et Végétale, Université Grenoble Alpes, CNRS, CEA, INRAE, IRIG, Grenoble, France</footnote>

<footnote n="2">² Laboratoire Modélisation et Exploration des Matériaux, CEA, Université Grenoble Alpes, IRIG, Grenoble, France</footnote>

<footnote>* Correspondence: morgane.michaud@cea.fr or juliette.jouhet@cea.fr</footnote>
soil or aquatic environment. However, despite its abundance, this nutrient is not evenly distributed and is often present in forms that cannot be easily assimilated by plants (Tiessen, 2008). This low Pi availability is often compensated by the use of phosphorus-enriched fertilizers, which contribute to significant soil and water pollution. To reduce the dependence on these fertilizers while enhancing crop yields, it is essential to understand the mechanisms by which photosynthetic organisms adapt to Pi deficiency (Cuyas et al., 2023).

In response to Pi deficiency, photosynthetic organisms activate adaptation mechanisms to increase Pi uptake from the environment and remobilize intracellular Pi reserves. In cells, Pi is primarily stored in the vacuole but is also found in significant quantities in various biomolecules such as nucleic acids and certain lipids (Poirier and Bucher, 2002). Approximately one third of intracellular Pi is stored in plant phospholipids, the main constituents of extraplastidial cell membranes (Poirier et al., 1991). During Pi deficiency, a portion of these membrane phospholipids is degraded to release Pi, which is then replaced by non-phosphorous lipids to maintain cell membrane integrity (Jouhet et al., 2004; Van Mooy et al., 2009; Hölzl and Dörmann, 2019). When Pi is limiting in plants, glycolipids can substitute for phospholipids in cell membranes: sulfoquinovosyldiacylglycerol (SQDG) replaces phosphatidylglycerol (PG) in photosynthetic membranes, and digalactosyldiacylglycerol (DGDG) substitutes for phospholipids in extraplastidial membranes (Jouhet et al., 2004; Andersson et al., 2005; Bolik et al., 2022). However, even after prolonged Pi deficiency, only a fraction of the phospholipids is degraded, limiting intracellular Pi remobilization and halting cell growth, possibly due to the differing physicochemical properties of phospholipids and glycolipids (Kanduč et al., 2017).

Algae, along with some fungi, bacteria, or bryophytes, are able to synthesize another class of non-phosphorous glycerolipids called betaine lipids. Currently, three types of betaine lipids have been identified in photosynthetic organisms: diacylglyceryl-*N*,*N*,*N*-trimethyl-homoserine (DGTS), diacylglycerylhydroxymethyl-*N*,*N*,*N*-trimethyl-β-alanine (DGTA), and diacylglycerylcarboxy-*N*-hydroxymethylcholine (DGCC) (Sato, 1992; Dembitsky, 1996; Kato et al., 1996). In algae, it is often suggested in the literature that these two lipid classes (i.e. phospholipids and betaine lipids) are interchangeable. Indeed, betaine lipids are synthesized in the endoplasmic reticulum (ER) (Riekhof et al., 2005b; Zhang et al., 2016) and localize in extraplastidial membranes (Eichenberger et al., 1993; Künzler et al., 1997) like phosphatidylcholine (PC), the most abundant phospholipid in extraplastidial membranes. Betaine lipids also share a trimethyl-nitrogen fragment with PC (Eichenberger et al., 1993; Kato et al., 1996). Furthermore, *Chlamydomonas reinhardtii* lacks PC but contains only DGTS (Yang et al., 2004), and in the yeast *Saccharomyces cerevisiae*, PC can be replaced by DGTS without a major impact on cell growth (Riekhof et al., 2014). However, across a wide range of organisms, no correlation has been found between

DGTS and PC content (Künzler and Eichenberger, 1997). The complementary association between betaine lipids and phospholipids has long been considered an adaptive strategy for coping with periodic Pi deficiency in the natural environment. However, the degree of change in cellular betaine lipid and PC contents in response to fluctuations in external Pi availability varies significantly among species (Cañavate et al., 2017) and the physicochemical properties of betaine lipids differ from those of PC, with DGTS bilayers being thicker, more rigid, and more repulsive than PC bilayers (Bolik et al., 2023).

These species-specific differences suggest metabolic diversity among algae, as well as versatility in betaine lipid function and requirement under Pi-starved conditions (Cañavate et al., 2016). Yet we lack a comprehensive understanding of betaine lipid functions. Despite the need for further research on betaine lipids in microalgae, only a few studies have been conducted since the initial works compiled by Dembitsky (1996). The presence and characterization of betaine lipids in cells are currently based on biochemical analyses, which cover only a limited range of organisms (Dembitsky, 1996; Künzler and Eichenberger, 1997; Makewicz et al., 1997; Vaskovsky et al., 1998; Cañavate et al., 2016). In a simplified view, betaine lipids are present in lower plants, algae, and some fungi but even these groups show several exceptions and betaine lipids are absent from seed plants, i.e. gymnosperms and angiosperms (Rozentsvet, 2004). All these studies were conducted prior to updates in the classification and the use of genetic tools to improve phylogenetic analysis. Since the genes responsible for DGTS synthesis were identified by Riekhof et al. (2005a, b), new phylogenetic analysis of DGTS biosynthesis enzymes have been possible (Murakami et al., 2018; Bolik et al., 2023), as well as genome searches for DGTS synthase genes, confirming the absence of DGTS synthesis enzymes in the genomes of seed plants. DGTS biosynthesis was first discovered in bacteria and is realized in two steps by the two enzymes BtaA and BtaB whereas in *Chlamydomonas*, in fungi, and more generally in eukaryotes, it is catalysed by a single bifunctional enzyme, Bta1, containing both the BtaA and BtaB domains. BtaA uses diacylglycerol (DAG) and S-adenosylmethionine (SAM) as substrates to produce diacylglycerol-O-homoserine, which is subsequently used by BtaB to produce DGTS through the sequential addition of three methyl groups to the nitrogen atom using SAM as the methyl donor (Riekhof et al., 2005a, b). The DGCC synthesis pathway is not yet known, and DGTA is hypothesized to be synthesized from DGTS, but the enzyme responsible for this conversion remains unidentified (Sato, 1991).

The goal of this work is to investigate the effect of betaine lipid production on seed plant development and Pi starvation stress response, to understand why the capacity to synthesize betaine lipids was lost during land plant evolution. We chose to produce DGTS in flowering plants because (i) DGTS is the most widely distributed betaine lipid and is present in lower
plants, and (ii) it is the only betaine lipid for which the genes and synthesis pathways have been characterized in different organisms. We generated DGTS in plants either by producing transgenic Arabidopsis lines expressing *Bta1* under the control of a phosphate starvation inducible promoter, pMGD3 (Shimojima et al., 2015), or by transiently overexpressing *Bta1* under the 35S promoter in *Nicotiana benthamiana*. Our results indicate that DGTS accumulates in extraplastidial membranes, leading to ER proliferation. DGTS production does not seem to improve growth or affect lipid remodelling in phosphate starvation.

## Materials and methods

### Plant materials and growth conditions

*Arabidopsis thaliana* ecotype Col-0 plants were used as the wild type. Two independent T-DNA insertion lines in the Col-0 background were obtained with the pMGD3-Sec63TM-Mg Bta1-HAx3 construction (Supplementary Fig. S1) using the *Bta1* gene from *Microchloropsis gaditana* (Naga_100016g36). The presence of the T-DNA was verified by PCR using a forward primer (number 1, Supplementary Table S1) located on the *MgBta1* gene and a reverse primer (number 2, Supplementary Table S1) located on the junction between the *MgBta1* gene and the HAx3 tag.

Calli were obtained from mesophyll tissue of 2-week-old Arabidopsis leaves, and grown on Murashige and Skoog (MS) (MSP09, Caisson Laboratories, Inc.) agar plates supplemented with 3% (w/v) sucrose, 1.2 mg l⁻¹ 2,4-dichlorophenoxyacetic acid and 0.8% (w/v) agar. After formation, the calli were maintained in 200 ml of liquid MS (MS without KH₂PO₄, Duchefa Biochemie) supplemented with 0.5 mM of phosphate, 1.5% (w/v) sucrose, and 1.2 mg l⁻¹ 2,4-dichlorophenoxyacetic acid. The calli were maintained under continuous light (100 µmol m⁻² s⁻¹) at 22 °C, agitated by rotary shaking at 125 rpm, and subcultured every 7 d. For phosphate starvation experiments, 5 ml of calli suspension were washed three times with 50 ml of MS medium -Pi (MS without KH₂PO₄, Duchefa Biochemie) with 0.5 mM Pi or without Pi before being resuspended in 200 ml of liquid MS with 0.5 mM or without Pi and grown for 4 or 8 d.

The Arabidopsis mutant and Col-0 seedling lines were grown *in vitro* on MS medium (MS without KH₂PO₄, Duchefa Biochemie) supplemented with 1% (w/v) sucrose, 1.2% (w/v) agar (A7921, Sigma-Aldrich), and 1 mM Pi for the Pi replete condition and 0.005 mM Pi for the Pi depleted condition. Seedlings were grown for 21 d under long day conditions (16 h/8 h light-dark photoperiod) at 20-22 °C with a photon flux density of 30-60 µmol m⁻² s⁻¹. The leaf portion above the hypocotyl was harvested from 21-day-old Arabidopsis seedlings, frozen in liquid nitrogen, and stored at -80 °C.

*Nicotiana benthamiana* wild type plants were grown under long day conditions (16 h/8 h light-dark photoperiod) at 18-20 °C with 60-70% humidity and a photon flux density of 100-120 µmol m⁻² s⁻¹. Plants were grown for 4 to 5 weeks on soil [potting soil/ vermiculite/ biological fungicide (Prestop, Lallemand) 35/7/10 (v/v/w in l/l/g)] and watered three times per week before agroinfiltration.

### Arabidopsis transgenic line construction

Genes were cloned into the pFP108 plant expression vector (To et al., 2006) in frame with 3xHA tags added in the C-terminal (pFP108-3xHA) and the pMGD3 promoter. Expression of genes from this vector were under the control of a NOS terminator (Supplementary Fig. S1A). The plasmids were transferred into *Agrobacterium tumefaciens* strain C58C1 and

used for stable transformation of Col-0 plants by floral dip. Transformed plants were selected using Basta® and two independent lines expressing the constructs were selected for further analysis which we named pMGD3-MgBTA1-1 and pMGD3-MgBTA1-2. Leaves from F₂ plants of these two lines were used for callus formation.

### Cloning of Bta1 fused proteins used for transient expression in *Nicotiana benthamiana*

*Bta1* sequences from *Kluyveromyces lactis* (KLLA0_F22198g, KlBta1), *Marchantia polymorpha* (MARPO_0023s0088, MpBta1), and *Chlamydomonas reinhardtii* (CHLRE_07g324200v5, CrBta1) were synthesized with codon optimization for expression in *A. thaliana* using the GeneArt synthesis service (ThermoFischer). The optimized sequences are provided in Supplementary Dataset S1. Forward/reverse primers are described in Supplementary Table S1.

For constructs without green fluorescent protein (GFP): all amplified *Bta1* genes were then cloned into the pFP108 plant expression vector in frame with 3xHA tag by Gibson assembly (Gibson et al., 2009) (Supplementary Fig. S1A). For the MgBTA1-GFP and nf-BTA1Pt-GFP constructs, amplified *Bta1* genes were cloned into the pFP108 plant expression vector (To et al., 2006) in frame with 3xHA tags and the GFP sequence at the C-terminus and the Sec63 (AtERDJ2B, AT4G21180) transmembrane fragment (aa 1-233) at the N-terminus as shown in Supplementary Fig. S1B. For the PtBTA1-GFP construct, the amplified *Bta1* gene was cloned into the pFP108 plant expression vector in frame with the GFP sequence at the C-terminus and Sec63 (AtERDJ2B, AT4G21180) transmembrane fragment (aa 1-233) at the N-terminus (Supplementary Fig. S1B). The expression of the genes from this vector was under the control of two 35S promoter and a NOS terminator. The plasmids were transferred into *Agrobacterium tumefaciens* C58C1 strain and used for transient transformation of *N. benthamiana* leaves by agroinfiltration.

### *Nicotiana benthamiana* agroinfiltration

*Agrobacterium tumefaciens* carrying *Bta1* plasmids were cultivated in 10 ml of luria broth (LB) medium containing the appropriate antibiotics supplemented with acetoxyringone (20 µM) for 24 h at 28 °C with 180 rpm agitation. The *A. tumefaciens* strain expressing the P19 protein was used in addition to all agroinfiltrations under the same conditions as the *A. tumefaciens* transformants. Cultures were centrifuged at 2880 g for 10 min. The pellets were resuspended in 5 ml of agroinfiltration buffer (1 mM MgCl₂, 200 µM acetoxyringone) and further diluted to obtain 10 ml of cell suspensions at OD₆₀₀ nm =1. Each transformed *Agrobacterium* strain was mixed with the P19 strain in a 1:1 (v:v) ratio. P19 alone was also used as a control. Infiltration was performed with a 1 ml syringe on the abaxial side of leaves of 4-5-week-old *N. benthamiana* plants. Plants were maintained for 4 or 8 d after agroinfiltration and excised for microscopy and chloroplast purification or ground in liquid nitrogen for western blot and lipid analysis.

### Confocal microscopy

Laser scanning confocal microscopy was performed on the platform µLife hosted by the Laboratory of Plant and Cell Physiology (LPCV) with a Zeiss LSM880 microscope equipped with a ×63/1.4 oil-immersion Plan-Apochromat objective, using Zen 2.3 SP1 acquisition software. Excitation/emission wavelengths were 488/493 to 570 nm for GFP and 561/582 to 622 nm for red fluorescent protein (RFP). Images were visualized and processed using ImageJ 1.53a software with the Fiji image processing package. For the fluorescence quantification, 30 images per sample from three independent experiments were analysed.
## Starch quantification

Starch quantification was performed using the Starch Assay kit from Sigma-Aldrich (SA20). Leaves of *N. benthamiana* plants were harvested 4 and 8 d after agroinfiltration and directly frozen in liquid nitrogen. After grinding, the samples were supplemented with 5 ml of 80% ethanol and heated for 5 min at 85 °C in a water bath. The samples were mixed and supplemented with another 5 ml of 80% ethanol before centrifugation at 1000× g for 10 min. The following steps were performed according to the manufacturer's instructions. For the glucose assay, a standard curve was generated using 10, 20, 30, 40, and 50 µg of the kit's glucose standard reagent, and used to calculate the amount of glucose in µg per mg of fresh weight. The experiment was repeated on three independent cultures of *N. benthamiana*.

## Photosynthetic analysis

Chlorophyll fluorescence measurements were conducted on agroinfiltrated *N. benthamiana* plants on day 4 (four plants per measurement) at room temperature using a Speedzen 3 chlorophyll fluorescence imaging set-up (JBeamBio). Samples were dark-adapted for 30 min prior to analysis. In this set-up, actinic light and saturating pulses were generated by red LEDs peaking at 630 nm, while measurement pulses (duration of 250 ms) were emitted by blue LEDs peaking at 470 nm. The detection time after each measurement pulse was set to 15 µs and no binning was applied. The reported fluorescence value is an average of 10 different points on different leaves, each consisting of four pixels. The maximum quantum efficiency of PSII was calculated as $F_v/F_m=(F_m-F_o)/F_m$, where $F_o$ and $F_m$ are the fluorescence emissions measured before and at the end of a 200 ms saturating pulse in dark-adapted leaves, respectively. $\Phi$PSII was calculated as $(F_m-F)/F_m$, where F is the steady-state fluorescence reached in the presence of illumination with actinic light and $F_m$ is the fluorescence emission measured at the end of a 200 ms saturating pulse, in light-adapted leaves. The electron transport rate (ETR) was calculated as $0.6 \times \text{PAR} \times \Phi$PSII where 0.6 correspond to the average ratio of PSII reaction centres to PSI reaction centres and PAR is the irradiation light level in the PAR range (400 to 700 nm) in µmol photons m⁻² s⁻¹. Non-photochemical quenching (NPQ) was recorded during 5 min of illumination at 672 µmol photons m⁻² s⁻¹ followed by 5 min of darkness. NPQ was calculated as $(F_m-F')/F_m$ (Maxwell and Johnson, 2000). Data were obtained from three independent experiments with three biological samples.

## Chloroplast purification

Agroinfiltrated plants were pre-incubated for 12 h at 4 °C before chloroplast purification. Sixteen leaves (3–4 g fresh weight) from eight plants were homogenized in a blender in 70 ml ice-cold buffer containing 0.4 M sorbitol, 20 mM tricine/KOH pH 8.4, 10 mM EDTA pH 8.0, 10 mM NaHCO₃, and 0.15% bovine serum albumin (BSA) (the supernatant was collected as the total fraction and stored at –80 °C). A crude chloroplast pellet was obtained by centrifugation at 2070× g for 2 min and further purified in 0.8 M sorbitol, 40 mM Hepes/KOH pH 7.6, 10 mM MgCl₂, 5 mM EDTA pH 8.0, and 20 mM NaHCO₃ (washing buffer) on a Percoll gradient previously prepared by centrifugation of 100% (v/v) Percoll in a JA20 rotor at 39 191× g for 55 min. After centrifugation at 9798× g for 10 min, intact chloroplasts were recovered from the lower phase. The chloroplasts were washed with 38 ml of washing buffer and centrifuged at 1482× g for 5 min in a JA20 rotor. The pellet was resuspended in 1 ml of washing buffer and stored at –80 °C for further analysis as the purified chloroplast fraction. The supernatant obtained after the first centrifugation was collected and centrifuged for 1 h at 4 °C at 255 530× g in a SW40 rotor. The pellet was resuspended in 1 ml of washing buffer and stored at –80 °C as the extraplastidial membrane fraction.

## Protein extraction and western blot analysis

Proteins from *N. benthamiana* agroinfiltrated leaves were extracted from ground tissue with 200 µl of protein buffer [8 M urea, 50 mM Tris-HCl pH 6.8, 1 mM ethylene glycol-bis(β-aminoethyl ether)-N,N,N',N'-tetraacetic acid (EGTA), 10 mM dithiothreitol (DTT)] and proteins from chloroplast purification fractions were extracted with 500 µl of protein buffer. Proteins were heated at 70 °C under 500 rpm agitation for 5 min then quantified by Bradford assay (Invitrogen). Ten micrograms of protein were used for migration on denaturing SDS-PAGE. After migration, the proteins were transferred to nitrocellulose membranes for 1 h at 25 V in transfer buffer [Bolt MES SDS Running Buffer (Invitrogen)]. The membranes were stained with Ponceau red [Ponceau red 0.2% (w/v) and trichloroacetic acid 3% (v/v)] and used for western blots after destaining. Immunodetection was performed with anti-GFP-HRP (130-091-833, Miltenyi Biotec) and anti-BIP2 (AS09481, Agrisera) antibodies. An anti-LHCP antibody was kindly provided by Olivier Vallon (IBPC, Paris).

## Transmission electronic microscopy

*Nicotiana benthamiana* leaves were prepared as previously described (Flori *et al.*, 2018). Briefly, leaf samples were fixed in 0.1 M phosphate buffer (PB) (pH 7.4) containing 2.5% (v/v) glutaraldehyde for 1 h at room temperature, under vacuum using a desiccator and then stored overnight at 4 °C. The samples were then washed five times in 0.1 M PB (pH 7.4), and the buffer was removed directly by pipetting. Samples were fixed by incubation for 2 h on ice in 0.1 M PB (pH 7.4) containing 2% osmium tetroxide and 1.5% potassium ferricyanide before washing five times with 0.1 M PB (pH 7.4). Samples were incubated in 0.1 M PB (pH 7.4) containing 0.1% tannic acid and incubated for 30 min in the dark at room temperature. The samples were again washed five times with 0.1 M PB (pH 7.4), dehydrated in ascending sequences of ethanol, and infiltrated with ethanol/EMBed-812 resin mixture. Finally, the samples were embedded in EMBed-812. Ultrathin sections (70–100 nm) were cut with a diamond knife on a PowerTome ultramicrotome (RMC Products) and collected on 200 mesh copper grids. Leaf samples were visualized by scanning transmission electron microscopy (STEM) using a FEI Tecnai microscope (FEI) set up at 200 kV (Fig. 7A; Supplementary Fig. S8). For Supplementary Videos S1, S2 and Fig. 7B, C, focused ion beam scanning electron microscopy (FIB-SEM) tomography was performed using a Zeiss CrossBeam 550 microscope, equipped with Fibics Atlas 3D tomography software as previously described (Uwizeye *et al.*, 2021). The resin block containing the cells was fixed on a stub with silver paste, and the surface abraded with a diamond knife in a microtome to obtain a perfectly flat and clean surface. The entire sample was metallized with 4 nm of platinum to avoid charging during observation. Inside the FIB-SEM, a second layer of platinum (1–2 µm) was deposited locally on the analysed area to mitigate possible curtaining artefacts. The sample was then abraded slice by slice with the $Ga^+$ ion beam (generally with a current of 700 nA at 30 kV). Each freshly exposed surface was imaged by scanning electron microscopy (SEM) at 1.5 kV and with a current of ~1 nA using the in-column EsB backscatter detector. The simultaneous milling and imaging mode was used for better stability, with an hourly automatic correction of focus and astigmatism. For each slice, a thickness of 6 nm was removed, and SEM images were acquired with a pixel size of 6 nm, resulting in an isotropic voxel size of 6×6×6 nm³. From the aligned FIB-SEM stack, the region of interest denoting the cell was cropped to remove unnecessary background using the Fiji software (https://imagej.net/Fiji). The 3D segmentation, reconstruction, and visualization was achieved using Dragonfly software (www.theobjects.com/dragonfly). Surface areas and volumes were calculated directly from the Dragonfly software.
## Lipid analysis

All lipid experiments were done at the LIPANG platform hosted by the LPCV.

Glycerolipids were extracted from plant samples frozen in liquid nitrogen immediately after harvest. Once freeze-dried, plants were crushed and suspended in 4 ml boiling ethanol for 5 min to prevent lipid degradation, and lipids were extracted by adding 2 ml methanol and 8 ml chloroform at room temperature according to Simionato et al. (2013). The mixture was then saturated with argon and stirred for 1 h at room temperature. After filtration through glass wool, the cell debris was rinsed with 3 ml chloroform/methanol 2/1 (v/v) and 5 ml 1% NaCl was added to the filtrate to initiate biphase formation. The chloroform phase was dried under argon before the lipid extract was solubilized in pure chloroform. Total glycerolipids were quantified from their fatty acids: a known amount of C15:0 was added to an aliquot fraction and the fatty acids present were converted to fatty acid methyl esters (FAMEs) by a 1 h incubation in 3 ml 2.5% H₂SO₄ in pure methanol at 100 °C (Jouhet et al., 2003). The reaction was stopped by adding 3 ml water and 3 ml hexane. The hexane phase was analysed by gas chromatography coupled to a flame ionization detector (GC-FID) (Perkin Elmer) on a BPX70 (SGE) column. FAMEs were identified by comparison of their retention times with those of standards (Sigma) and quantified by the surface peak method using C15:0 for calibration.

Lipid extracts corresponding to 25 nmol of total fatty acids were dissolved in 100 µl of chloroform/methanol 2/1 (v/v) containing 125 pmol of each internal standard. The internal standards used were PE 18:0-18:0 and DAG 18:0-22:6 from Avanti Polar Lipid and SQDG 16:0-18:0 extracted from spinach thylakoids (Demé et al., 2014) and hydrogenated as described in Buseman et al. (2006). The lipids were then separated by HPLC and quantified by MS/MS.

The HPLC separation method was adapted from Rainteau et al. (2012). Lipid classes were separated on an Agilent 1260 Infinity II HPLC system using a 150 mm ×3 mm (length × internal diameter) 5 µm diol column (Macherey-Nagel) at 40 °C. The mobile phases consisted of hexane/isopropanol/water/1 M ammonium acetate, pH 5.3 [625/350/24/1 (v/v/v/v)] (A) and isopropanol/water/1 M ammonium acetate, pH 5.3 [850/149/1 (v/v/v)] (B). The injection volume was 20 µl. After 5 min, the percentage of B was increased linearly from 0% to 100% over 30 min and remained at 100% for 15 min. This elution sequence was followed by a return to 100% A in 5 min and equilibration with 100% A for 20 min before the next injection, resulting in a total run time of 70 min. The flow rate of the mobile phase was 200 µl min⁻¹. The different glycerophospholipid classes were eluted sequentially as a function of the polar head group.

Mass spectrometric analysis was performed on a 6470 triple quadrupole mass spectrometer (Agilent) equipped with a Jet stream electrospray ion source with the following settings: drying gas heater: 230 °C, drying gas flow 10 1 min⁻¹, sheath gas heater: 200 °C, sheath gas flow: 10 1 min⁻¹, nebulizer pressure: 25 psi, capillary voltage: ±4000 V, nozzle voltage ±2000. Nitrogen was used as the collision gas. Quadrupoles Q1 and Q3 were operated at widest and unit resolution, respectively. DGTS and PC analysis was performed in positive ion mode by scanning for precursor ions at 236 m/z and 184 m/z at a collision energy (CE) of 55 and 35 eV, respectively. SQDG analysis was performed in negative ion mode by scanning for precursor ion at -225 m/z at a CE of -55 eV. Phosphatidylethanolamine (PE), phosphatidylinositol (PI), phosphatidylglycerol (PG), phosphatidylinositol (PS), monogalactosyldiacylglycerol (MGDG), and digalactosyldiacylglycerol (DGDG) measurements were performed in positive ion mode by scanning for neutral losses of 141 Da, 277 Da, 189 Da, 185 Da, 179 Da, and 341 Da at CEs of 29 eV, 21 eV, 25 eV, 21eV, 8 eV, and 11 eV, respectively. Quantification was performed by multiple reaction monitoring (MRM) with a 30 ms dwell time. DAG and triacylglycerol (TAG) species were identified and quantified by MRM as singly charged ions [M+NH₄]⁺ at a CE of 19 and 26 eV, respectively,

with a 30 ms dwell time. DPG species were quantified by MRM as singly charged ions [M-H]⁻ at a CE of -54 eV with a 50 ms dwell time. Mass spectra were processed using MassHunter Workstation software (Agilent) for identification and quantification of lipids. Lipid contents (pmol) were corrected for response differences between internal standards and endogenous lipids and by comparison with a quantified control (QC) using an in-house software developed by the LIPANG platform (Jouhet et al., 2017). The QC extract corresponds to a known mixed lipid extract from *Chlamydomonas* and Arabidopsis cell cultures qualified and quantified by thin layer chromatography (TLC) and GC-FID as described by (Jouhet et al., 2017).

## Statistical analysis

For photosynthesis measurements, normality was tested using Skewness and Kurtosis tests. Since the data followed a normal distribution, the statistical differences between samples were analysed using the parametric two-way ANOVA followed by Tukey's multiple comparison test.

The fluorescence quantification and lipid analysis data did not follow a normal distribution, so a non-parametric test was used for the analyses. The Kruskal-Wallis pairwise comparison test followed by the Benjamini, Krieger and Yekutieli post-hoc test was used to detect significant differences between samples. As the Benjamini, Krieger and Yekutieli post-hoc test is more conservative, it was used to minimize the risk of false positive results.

## Results

### Selection of DGTS synthase genes for DGTS production in plants

Bta1 genes from various species were identified using a BLAST search with the protein sequence of *Chlamydomonas reinhardtii* BTA1. To identify BTA1 enzymes that are functional in higher plants, five Bta1 sequences from different lineages were selected for functional tests by transient expression in *Nicotiana benthamiana*. Therefore, Bta1 cDNA from *Microchloropsis gaditana* (*MgBta1*, eustigmatophyte), *Phaeodactylum tricornutum* (*nf-PtBta1*, diatom), *Kluveromyces lactis* (*KIBta1*, fungus), *Marchantia polymorpha* (*MpBta1*, bryophyte), and *Chlamydomonas reinhardtii* (*CrBta1*, green algae) were cloned under the control of the 35S promoter and transiently overexpressed in *N. benthamiana* leaves by agroinfiltration (Supplementary Fig. S1A). Eight days after infiltration, lipids were extracted from the leaves and separated by thin layer chromatography (TLC) (Fig. 1). A lipid spot corresponding to DGTS was clearly identified when the enzymes of *Microchloropsis gaditana*, *K. lactis*, *Marchantia polymorpha*, and *C. reinhardtii* were expressed, showing that DGTS could be produced in higher plants by expressing a BTA1 protein. These results also confirmed that the Bta1 genes of *Marchantia polymorpha* and *Microchloropsis gaditana* encode functional enzymes. However, we did not detect DGTS production in leaves expressing the Bta1 construct from *P. tricornutum*. Upon further analysis of the annotated sequence retrieved from the Joint Genome Institute (JGI), we noticed an intron, not initially annotated, present in the N-terminus sequence that was cloned from the genomic DNA of *P. tricornutum*.
(Supplementary Fig. S1B). Consequently, we cloned the correct version of the *Phaeodactylum* gene from cDNA. To ensure that the absence of DGTS synthesis was not due to a defect in ER addressing, we fused both *P. tricornutum* and *M. gaditana* constructs to the N-terminal ER transmembrane domains of Sec63. We then transiently expressed both constructs in *N. benthamiana* leaves. To better separate DGTS from endogenous lipids, we performed 2D-TLC with lipids extracted from non-transformed plants or plants transformed with the new *P. tricornutum* BTA1 construction (thereafter named PtBTA1) and MgBTA1 as a positive control (Supplementary Fig. S2). This experiment showed that PtBta1 was indeed functional but produced DGTS at a lower level in *N. benthamiana* compared to MgBta1, probably due to differences in expression levels of the two proteins (see below). Thus, the presence of the N-terminal intron leads to the production of a non-functional enzyme that we named nf-PtBTA1. For further experiments, MgBta1, PtBta1, and nf-PtBta1 constructs were chosen for strong, moderate, and inefficient DGTS production in plants, respectively.

## Analysis of Arabidopsis seedlings producing DGTS under phosphate starvation

We initially attempted to transform Arabidopsis with the highly active *MgBta1* under the control of the 35S promoter. However, we were unable to obtain transformed seeds expressing the transgene, suggesting that high expression of *Bta1* in *A. thaliana* might have a deleterious effect. We then decided to express MgBta1 under a phosphate starvation inducible promoter, pMGD3 (Supplementary Fig. S1C) (Shimojima et al., 2015). To ensure that the production of DGTS was localized in the ER, where it is supposed to occur in algae (Riekhof et al., 2005a; Zhang et al., 2016) and where PC synthesis takes place (Nakamura, 2021), we targeted the construction to the ER by adding the transmembrane fragment of Sec63. We selected two independent lines from separate transformation events that contained the transgene (Supplementary Fig. S3) and synthesized DGTS in response to phosphate starvation (see below). These independent lines were named pMGD3-*MgBta1-1* and pMGD3-*MgBta1-2* and we characterized them on two different systems: seedlings grown on plates and callus cell cultures. Non-transformed Col-0 plants were used as controls.

Col-0, pMGD3-*MgBta1-1*, and pMGD3-*MgBta1-2* seeds were grown for 21 d *in vitro* on MS medium with or without Pi (Fig. 2A). As expected, the absence of Pi severely slowed down plant growth, but no major phenotypic difference was detected between Col-0 and the lines expressing BTA1 under both conditions. Lipids were extracted from the rosette leaves and analysed by HPLC-MS/MS (Fig. 2B, C). Under +Pi conditions, lipid content was similar in both Col-0 and the BTA1 overexpressing lines (Fig. 2B), and no significant differences in lipid composition were observed between Col-0 and the mutants (Fig. 2C). Furthermore, DGTS was barely detected

in the mutants, confirming that genes under the control of pMGD3 were not expressed under +Pi conditions. Under -Pi conditions, the plant lipid content slightly decreased compared to +Pi conditions, but no significant difference between Col-0 and the mutants was observed (Fig. 2B). DGTS synthesis was induced in the mutants, reaching up to 4% of total glycerolipids, but the rest of the glycerolipidome was not significantly affected. Phospholipids were degraded and DGDG production significantly increased during Pi starvation, as previously observed (Fig. 2C) (Jouhet et al., 2004). DGTS production did not seem to affect glycerolipid remodelling, likely because the induction of DGTS synthesis was not very strong.

We also analysed the distribution of DGTS molecules under -Pi, as DGTS was barely detectable under +Pi conditions (Fig. 2D). Each molecule is described first by the number of carbons in the acyl groups and secondly by the number of unsaturations. For example, DGTS 36:2 is a 1,2-diacyl-*sn*-glycero-3-trimethylhomoserine in which the acyl groups at *sn*-1 and *sn*-2 positions contain 36 carbons in total and two double bonds. DGTS molecule distribution and desaturation patterns were similar to those of PC molecules (Fig. 2E), with a slight enrichment of the C34 diacylglycerol backbone, corresponding to a 16 C and a 18 C fatty acid, compared to PC, which is enriched in the C36 diacylglycerol backbone, corresponding to two C18 fatty acids. Because *Microchloropsis* is an alga containing very few fatty acids with C18 (Billey et al., 2021), this difference could be due to the enzyme specificity. Chloroplast lipids (MGDG, DGDG, SQDG, and PG) have distinctive lipid compositions (Fig. 2F; Supplementary Fig. S4A-C) that differ from DGTS, and because DGTS composition is similar to PC, DGTS is likely produced in the ER by the Kennedy pathway. Thus, the BTA1 enzyme seems to use the same DAG pool as PC (Karki et al., 2019).

## Analysis of DGTS production in Arabidopsis calli under phosphate starvation

To confirm this hypothesis and to test the impact of DGTS production on the kinetics of lipid remodelling under phosphate starvation, calli were generated from the same Col-0, pMGD3-*MgBTA1-1*, and pMGD3-*MgBTA1-2* lines and cultivated under phosphate replete conditions (+Pi) and phosphate starved conditions (-Pi) for 4 and 8 d (Fig. 3). Lipids were extracted and analysed by HPLC-MS/MS. Since calli are non-photosynthetic cells, their phospholipid-to-galactolipid ratio is much higher than in leaves, making lipid remodelling defects more noticeable in calli. After 8 d of phosphate starvation, a slight but non-significant reduction in the total glycerolipid content was observed (Fig. 3A), alongside phospholipid degradation and an increase in DGDG and SQDG levels (Fig. 3B). In the transgenic lines, DGTS levels increased more than 100 times (Fig. 3C), yet represented less than 5% of the total glycerolipid content, even after 8 d of starvation. Similar to what we observed in leaves, DGTS synthesis had no major
<figure><img src="image_2.png" /><figcaption>**Fig. 1.** TLC analysis of lipids extracted from *Nicotiana benthamiana* leaves transiently expressing different *Bta1* genes. Lipid control for migration: DGTS, PC and PE. The arrow in red indicates the DGTS position, the arrows in black the endogenous glycerolipids (MGDG, PE, PG, DGDG, PC, and PI). Ctrl corresponds to leaf extract without *Bta1*, all other lines (Mg, nf-Pt, KI, Mp, Cr) correspond to leaf extracts expressing different genes of *Bta1*. Mg: *Microchloropsis gaditana*; nf-Pt: non-functional *Phaeodactylum tricornutum*; KI: *Kluveromyces lactis*; Mp: *Marchantia polymorpha*; Cr: *Chlamydomonas reinhardtii*. MGDG, monogalactosyldiacylglycerol; DGDG, digalactosyldiacylglycerol; PG, phosphatidylglycerol; PI, phosphatidylinositol; PE, phosphatidylethanolamine; PC, phosphatidylcholine.</figcaption></figure>

effect on the kinetics of lipid remodelling, except for a slight, non-significant delay in the reduction of the glycerolipid content, potentially due to the neosynthesis of DGTS (Fig. 3C).

When examining fatty acid composition and glycerolipid molecular distribution (Supplementary Fig. S5A), the mutant cells showed an enrichment in 18:1 fatty acid, a phenomenon observed in +Pi condition, suggesting it was independent of DGTS production. This enrichment was more likely due to differences in cell division kinetics, as previously reported (Meï *et al.*, 2015), and not to the transgene expression. This pattern was reflected in the glycerolipid composition (Supplementary Fig. S5C, D, E). In terms of DGTS composition (Supplementary Fig. S5B), as in leaves, DGTS molecular distribution was similar to that of PC, with a slight enrichment in C34 species, contrasting with the DGDG composition. This suggests that DGTS was synthesized from a distinct pool of DAG than the one dedicated to extraplastidial lipid synthesis.

## Analysis of DGTS overproduction in *Nicotiana benthamiana* leaves

Given the low levels of DGTS production in our stable *A. thaliana* lines, we used the transient expression system in *N.*

*benthamiana* described earlier to achieve significantly higher DGTS levels and assess the impact on the lipidome and membrane structure of higher plants. Protein overexpression through agroinfiltration can influence leaf metabolism by inhibiting chloroplast gene expression and activating oxidative stress responses (Hamel *et al.*, 2024). To ensure that the observed leaf phenotypes were due to DGTS production and not to protein overexpression or agroinfiltration stresses, three DGTS synthases targeted to the ER with different amount of DGTS production were selected: MgBTA1 for high DGTS production, PtBTA1 for moderate production, and nf-PtBTA1 for ineffective production (Supplementary Fig. S1B).

The three enzymes were fused to the Sec63 transmembrane domains at the N-terminus and to GFP at the C-terminus (Supplementary Fig. S1B). To validate their ER localization, GFP fluorescence was observed using confocal microscopy (Fig. 4). For all three enzymes, the GFP signal matched the ER membrane pattern in the epidermal cells, exhibiting the standard reticulated network pattern and nuclear envelope staining (Fig. 4A). As variations in the GFP signal intensities were reproducibly observed between the three constructions, we quantified the level of GFP fluorescence within cells from independent agroinfiltration experiments. MgBTA1-GFP and nf-PtBTA1-GFP showed consistently
<figure><img src="image_3.png" /><figcaption>**Fig. 2.** pMGD3-MgBta1 seedling analysis. (A) Pictures of pMGD3-MgBta1-1, pMGD3-MgBta1-2, and Col-0 seedlings grown for 21 d on MS medium supplemented with 1 mM Pi (+Pi) or 0.005 mM Pi (-Pi). (B–F) Lipid analysis of pMGD3-MgBta1-1, pMGD3-MgBta1-2, and Col-0 seedlings leaves grown for 21 d on MS medium supplemented with 1 mM Pi (+Pi) or 0.005 mM Pi (-Pi). Values shown correspond to the median of five biological replicates and the error bars represent the range between minimum and maximum values. (B) Quantity of glycerolipids in nmol per mg of rosette fresh weight. (C–F) Glycerolipid distribution in mol % (C) and molecular distribution within DGTS (D), PC (E), and DGDG (F) classes determined by HPLC-MS/MS. The first number corresponds to the sum of the carbon numbers of the two fatty acids, and the second number to the number of unsaturations. Asterisks indicate significant differences (*P*<0.05) in comparison with the +Pi condition by a Kruskal–Wallis test. SQDG, sulfoquinovosyldiacylglycerol; MGDG, monogalactosyldiacylglycerol; DGDG, digalactosyldiacylglycerol; PG, phosphatidylglycerol; DPG, diphosphatidylglycerol; PI, phosphatidylinositol; PS, phosphatidylserine; PE, phosphatidylethanolamine; PC, phosphatidylcholine; PA, phosphatidic acid; DGTS, diacylglyceryl-*N*,*N*,*N*-trimethyl-homoserine; DAG, diacylglycerol; TAG, triacylglycerol.</figcaption></figure>

higher fluorescence intensities compared to PtBTA1-GFP (at least twice as high), indicating lower expression levels of the PtBTA1-GFP enzyme (Fig. 4B). These results can explain the lower DGTS production observed for PtBTA1 overexpressing line in the TLC analysis (Supplementary Fig. S2). To confirm this result, we tried to quantify the BTA1-GFP signal by western blot analysis. However, whatever the protein extraction methods and the protease inhibitors used,

no signal corresponding to the intact protein was detected but a signal corresponding to the GFP alone was measured (Supplementary Fig. S6). Because the fluorescence signal was not cytosolic as expected for GFP alone *in planta*, the BTA1-GFP fusion was likely preserved *in vivo*. The intensity of the GFP band corresponded to the fluorescence levels: very high for MgBTA1-GFP and *nf*-PtBTA1-GFP, and barely detectable for PtBTA1-GFP. To validate the ER localization,
<figure><img src="image_4.png" /><figcaption>**Fig. 3.** Lipid analysis of pMGD3-MgBta1 calli. (A–D) Lipid analysis of pMGD3-MgBta1-1, pMGD3-MgBta1-2, and Col-0 calli grown in phosphate replete (+Pi) and deplete (–Pi) conditions and harvested at day 4 (D4) and day 8 (D8). Values shown correspond to the median of six biological replicates and the error bars represent the range between minimum and maximum values. (A) Adjusted glycerolipid content per mg of fresh weight, (B) glycerolipid composition in mol %, (C) DGTS content and (D) total lipid content, all determined by HPLC-MS/MS. Asterisks indicate significant differences (\*\*P<0.01; \*P<0.05) in comparison with the +Pi (D4) condition by a Kruskal–Wallis test. Abbreviations as in Fig. 2.</figcaption></figure>

MgBTA1-GFP and nf-PtBTA1-GFP were co-expressed with an ER marker fused to RFP (Fig. 4C). GFP and RFP co-localized, indicating that the constructed proteins were indeed restricted to the ER and no significant degradation of the fusion releasing GFP alone occurred in planta.

Lipid analysis of leaves expressing P19 only (infiltration control), nf-PtBTA1-GFP, PtBTA1-GFP, and MgBTA1-GFP fused proteins at 4 and 8 d post-infiltration was performed using HPLC-MS/MS (Fig. 5; Supplementary Fig. S7). Interestingly, nf-PtBTA1-GFP produced a small amount of DGTS, undetectable by TLC (Fig. 1), suggesting that the enzyme with an extended N-terminus was not entirely inactive (Fig. 5A). However, DGTS synthesis was negligible compared to leaves expressing PtBTA1-GFP and MgBTA1-GFP, despite a protein expression level similar to that of MgBTA1 (Fig. 4B). In the PtBTA1 and MgBTA1 overexpressing lines,

DGTS accumulated at a very high level, even equivalent to MGDG content at D8 in the MgBTA1 overexpressing leaves (Fig. 5A, B), reflecting the difference of expression of the two proteins. Indeed, DGTS accounted for up to 30 mol% of the glycerolipid content in leaves overexpressing MgBTA1-GFP (Fig. 5B). When looking at the DGTS molecular composition, as in Arabidopsis, the DGTS molecule distribution was comparable to the PC distribution, with a preference for a C34 DAG backbone, distinct from the DGDG distribution (Supplementary Fig. S7A–C). This C34 enrichment in DGTS can be seen at the level of the total fatty acid composition in MgBTA1 agroinfiltrated leaves with a global enrichment of 16:0 (Supplementary Fig. S7D).

Despite the high DGTS amount, no significant changes in global glycerolipid content were observed, except maybe for a slight non-significant increase of lipid content in the MgBTA1
<figure><img src="image_5.png" /><figcaption>**Fig. 4.** Subcellular localizations of BTA1 proteins overexpressed in *Nicotiana benthamiana*. (A) Subcellular localization of MgBTA1-GFP, PtBTA1-GFP, and nf-PtBTA1-GFP proteins transiently expressed in *N. benthamiana* leaves 8 d after infiltration. Scale bar, 10 µm. (B) Fluorescence quantification from confocal images of MgBTA1-GFP, PtBTA1-GFP, and nf-PtBTA1-GFP fused proteins transiently expressed in *N. benthamiana* leaves 8 d after infiltration. Determination of corrected total cell fluorescence (CTCF) was done with ImageJ software with a total of 30 images per construction, taken from three independent plants. CTCF = integrated density – (area of selected cell × mean fluorescence of background readings). Asterisks indicate significant differences (\*\*P<0.01; \*\*\*P<0.001) in pairwise comparisons by Kruskal–Wallis test. (C) Subcellular localization of MgBTA1-GFP and nf-PtBTA1-GFP proteins transiently co-expressed with the ER marker ER-RFP in *N. benthamiana* leaves. ER-RFP: RFP protein fused to an HDEL retention signal at the C-terminus. Scale bar, 10 µm.</figcaption></figure>

overexpressing line (Fig. 5C). Looking at the glycerolipid pro-
portion within the leaves (Fig. 5B) although the differences
were non-significant, galactolipid levels appeared to be affected
when DGTS production exceeded 10% of the glycerolipid
content, while SQDG remained unaffected. PA and PC levels,
as well as PG at D8, also seemed to decrease in MgBTA1-
overexpressing leaves, possibly to accommodate the increase
in DGTS.

Given the high DGTS production in the MgBTA1 overex-
pressing leaves and the minimal impact on other glycerolipids,
we investigated the subcellular localization of DGTS, particu-
larly regarding chloroplast membranes, representing more than
70% of the cell membranes. Chloroplasts were purified from D4
agoinfiltrated leaves either with the nf-PtBTA1-GFP vector
as a control with a high amount of protein overexpression and
a poor level of DGTS production, or with the MgBTA1-GFP
vector as a high producer of DGTS, more than 20% of total

glycerolipids (Fig. 6A, B). Chloroplast fractions were character-
ized by a high proportion of glycolipids and a very low amount
of PE and PC in comparison to the total fractions, whereas
the opposite was true for the endomembrane fractions. The
chloroplast fractions for both overexpressing lines were pure,
with an enrichment of LHCP (a thylakoid marker) and no
labelling of BIP2 (an ER marker) and GFP (a BTA1 marker)
(Fig. 6C). In these fractions, DGTS was detected at the same
low level (<1.5%) in both nf-PtBTA1 and MgBTA1 agroinfil-
trated leaves (Fig. 6B). The extraplastidial fractions were con-
taminated by chloroplasts with a high amount of galactolipids
and the presence of LHCP; however, they were enriched in
endomembranes with a strong BIP2 signal and an enrichment
in PE absent from chloroplast membranes. These fractions con-
tained roughly the same amount of DGTS as the total extracts.
Overall, this experiment indicates that DGTS was absent from
the chloroplast membranes and was restricted to extraplastidial
<figure><img src="image_6.png" /><figcaption>**Fig. 5.** Lipid analysis of *Nicotiana benthamiana* leaves agroinfiltrated with BTA1 constructs. (A–D) Lipid analysis of *N. benthamiana* leaves agroinfiltrated with P19 only, nf-PtBTA1-GFP, PtBTA1-GFP, and MgBTA1-GFP fused proteins. Leaves were harvested 4 (D4) and 8 (D8) d after agroinfiltration. Control: *N. benthamiana* agroinfiltrated with P19 protein (negative control). Each replicate corresponds to two leaves that were pooled. Represented values correspond to the median of three biological replicates and the error bars represent the range between minimum and maximum values. Asterisks indicate significant differences (\*\*P<0.01; \*P<0.05) in comparison to control D4 and D8 by a Kruskal–Wallis test. (A) Quantity of each class of glycerolipids determined by HPLC-MS/MS. (B) Total glycerolipid quantity determined by HPLC-MS/MS. (C) Log₂ fold change in lipid quantities between BTA1 constructions (nf-PtBTA1-GFP, PtBTA1-GFP, and MgBTA1-GFP) and P19. (D) Glycerolipid composition. Abbreviations as in Fig. 2.</figcaption></figure>

membranes, highlighting the specificity of the lipid transport pathway towards chloroplasts.

To assess the impact of DGTS accumulation on extra-plastidial membranes, leaves agroinfiltrated at D4 with P19, nf-PtBTA1-GFP, PtBTA1-GFP, and MgBTA1-GFP were observed by electronic microscopy (Fig. 7A; Supplementary Fig. S8). Chloroplast and thylakoid ultrastructure appeared unaffected, except maybe for an increase in starch content in MgBTA1 overexpressing lines (Fig. 7A, first lane). To validate whether this was an artefact, starch content was measured in different leaf extracts from different independent plants and leaves. The starch content was highly variable depending on the experiment and seemed to correlate more with the level of protein overexpression than with DGTS production, nf-PtBTA1 and MgBTA1 having the highest level of protein expression and starch content with very different DGTS amounts (Supplementary Fig. S9A). Photosynthesis efficiency was also measured (Supplementary Fig. S9B, C). As expected, agroinfiltration had a negative impact on the electron transfer rate (ETR) and maybe a slight delay on the relaxation time of the non-photochemical quenching (NPQ) (Hamel et al., 2024). No difference was observed between

P19 and the BTA1 overexpressing lines in terms of NPQ. However, the ETR seemed to be slightly related to the DGTS amount with an ETR value following the DGTS proportion: ETR<sub>MgBTA1</sub>≤ETR<sub>PtBTA1</sub>≤ETR<sub>nf-PtBTA1</sub> and ETR<sub>P19</sub>. Further investigations would be needed to confirm if this finding is related to the observed galactolipid reduction.

In the MgBTA1 overexpressing lines, membrane proliferations were observed in the cytosol possibly related to the extraplastidial localization of DGTS (Fig. 7A; Supplementary Fig. S8). These structures were absent in the leaves overexpressing nf-PtBTA1, indicating a link to DGTS production rather than protein overexpression. The architecture of these membrane structures was analysed in 3D by focused ion beam scanning electron microscopy (FIB-SEM) in comparison with what could be observed in the nf-PtBTA1 overexpressing line (Supplementary Videos S1, S2). A snapshot of the reconstituted cell architecture showing the cell wall, the mitochondria, the peroxisome, and the ER without the plastid is presented Fig. 7B. In both reconstructions, the ER was surrounding mitochondria and peroxisomes that were embedded within ER tubules. Furthermore, reconstructed ER from MgBTA1-GFP overexpressing line was constituted of numerous tubule layers
A

<figure><img src="image_7.png" /></figure>

B

Glycerolipid composition

<figure><img src="image_8.png" /><figcaption>**Fig. 6.** Localization of DGTS in *Nicotiana benthamiana* leaves overexpressing either MgBTA1-GFP or nf-PtBTA1-GFP. (A) Strategy used to isolate chloroplast and extraplastidial fractions from 3–4 g of *N. benthamiana* leaves (representing eight different plants with two leaves per plant independently infiltrated) 4 d after agroinfiltration with MgBTA1-GFP and nf-PtBTA1-GFP fused proteins. Curved arrows represent the centrifugation steps. (B) Proportion of glycerolipids determined by HPLC-MS/MS in each fraction. Total, total fraction;; chloroplast, purified chloroplast fraction. (C) Western blot analysis of purified chloroplast and endomembrane fractions using the BIP2 antibody for ER, LHCP for thylakoids, and GFP for BTA1 fused proteins.</figcaption></figure>

C

<figure><img src="image_9.png" /></figure>

but with a membrane organization very similar to the standard ER architecture observed in the nf-PtBTA1 overexpressing line (Fig. 7C). In reference to cell wall volume, the volume of the ER in the MgBTA1-GFP overexpressing line was eight times greater than the volume of the ER in the nf-PtBTA1-GFP overexpressing line. Overall, these results indicate that DGTS was probably synthesized in the ER and its accumulation stimulated ER proliferation at the expense of the other membranes.

Discussion

Betaine lipids are absent in seed plants but can be found in chlorophytes, bryophytes, and pteridophytes (Vogel and Eichenberger, 1992; Rozentsvet, 2004). Since the identification of the genes responsible for DGTS synthesis (Riekhof

*et al.*, 2005a, b), genome analyses and phylogenetic reconstructions have confirmed the absence of the *BTA1* gene in seed plants—gymnosperms and angiosperms—and have linked its evolution to the habitat, whether freshwater or marine (Bolik *et al.*, 2023). By expressing *BTA1* genes derived from marine or freshwater organisms in *Nicotiana benthamiana* (Fig. 1), we demonstrated that plants retain the ability to produce DGTS and that the absence of the *BTA1* gene is solely responsible for the loss of their capacity to synthetize DGTS. However, in our hands, we never managed to obtain stable *Arabidopsis thaliana* lines that overexpressed DGTS under the control of the constitutive 35S promoter. Previous studies have indicated that DGTS bilayers are thicker than PC bilayers and that the DGTS bilayer thickness could be sensitive to hydration (Bolik *et al.*, 2023). Seed plants lost their desiccation tolerance at the vegetative stage compared to more primitive plants and produced seeds representing a very dry stage (less than 20%
<figure><img src="image_10.png" /><figcaption>**Fig. 7.** Impact of BTA1 proteins transient expression on *Nicotiana benthamiana* leaf cell architecture. (A) Electronic microscopy observations of the architecture of cellular membranes in *N. benthamiana* leaves agroinfiltrated with BTA1 fused proteins (nf-PtBTA1-GFP, PtBTA1-GFP, and MgBTA1-GFP) or P19 as the control at day 4. Scale bar is 2 µm for the top four images and 500 nm for the rest of the images. Red asterisks indicate membrane ER proliferation. (B–C) Representative 3D architecture of the cell (B) and the ER alone (C) from FIB-SEM imaging from *N. benthamiana* leaves agroinfiltrated with MgBTA1-GFP or nf-PtBTA1-GFP. Blue, the cell wall; red, the ER; yellow, the mitochondria; green, the peroxisomes. These pictures are extracted from Supplementary Video S1 for MgBTA1-GFP and Supplementary Video S2 for nf-PtBTA1-GFP.</figcaption></figure>

protoplasmic water) (Oliver et al., 2000). The disappearance of the *BTA1* gene coincides with these two parameters. Further experiments are needed to determine whether this is a mere correlation or a causal relationship.

Betaine lipids are known to substitute for extraplastidial phospholipids in various organisms, such as bacteria, fungi, and algae under phosphate starvation (Van Mooy et al., 2009; Riekhof et al., 2014; Abida et al., 2015; Mühlroth et al., 2017; Murakami et al., 2018; Oishi et al., 2022). In seed plants, however, this function is performed by DGDG, which is synthesized in plastids and exported outside of the chloroplast (Jouhet et al., 2004; Andersson et al., 2005). To investigate this remodeling, we stably expressed the *MgBTA1* gene in *A. thaliana* under the control of a phosphate starvation inducible promoter, pMGD3 (Shimojima et al., 2015). Under phosphate starvation, DGTS was synthesized at very low levels in vegetative tissues without affecting plant growth or lipid remodelling: the increase of DGDG content and phospholipid degradation were similar in wild type plants and overexpressing lines.

However, the DGTS synthesis remained low, resulting in a level of DGTS representing less than 5% of total glycerolipids. The pMGD3 promoter has been shown to be induced more than 10-fold under phosphate starvation compared to control conditions (Shimojima et al., 2015), whereas the 35S promoter is known to induce gene expression more than 100-fold for the same genes (Demski et al., 2020). This difference in promoter strength likely explains the low DGTS production.

DGTS synthesis occurs in two steps: the first one uses DAG and SAM as substrates to produce diacylglycerol-O-homoserine which then undergoes successive methylation steps from the SAM donor to form DGTS (Riekhof et al., 2005a, b). During phosphate starvation, DAG is supposed to be released from PC for recycling into DGDG (Jouhet et al., 2003; Nakamura et al., 2009); however, the exact localization of DAG release remains unknown. In our experiments, DGTS production under phosphate starvation did not impact DGDG synthesis or its molecular composition (Figs 2–3; Supplementary Fig. S5). We hypothesize that
pMGD3-*MgBTA1* lines no. 1 and 2 grown 21 d in presence of 1 mM (+Pi) or 0.005 mM (-Pi) of phosphate.

Fig. S5. Lipid analysis of *Arabidopsis thaliana* calli from Col-0 and pMGD3-*MgBTA1* lines no. 1 and 2 cultivated for 4 d in the presence of 1 mM (+Pi (D4)) or 4 d (D4) and 8 d (D8) in absence of phosphate (-Pi).

Fig. S6. Western blot analysis of agroinfiltrated *N. benthamiana* leaves.

Fig. S7. Glycerolipid distribution of *Nicotiana benthamiana* leaves agroinfiltrated with or without (control) BTA1-GFP constructions.

Fig. S8. Electron micrographs of agroinfiltrated *N. benthamiana* leaves.

Fig. S9. Starch and photosynthesis efficiency analysis on *N. benthamiana* leaves 4 d after agroinfiltration with BTA1-GFP constructions.

Table S1. List of the primers used for this study.

Dataset S1. Optimized sequences of the *BTA1* gene for plant expression.

## Acknowledgements

The authors would like to thank Sylvie Figuet for the maintenance of the plant growth facility and Dimitri Tolleter for his help with the photosynthesis measurement.

## Author contributions

SS generated the plant material and carried out the cloning and most of the experiments; MS carried out the lipid analysis; SR developed the software for reprocessing the GC-FID and LC-MS data; CA undertook the chloroplast purification; GSL, DF, and P-HJ carried out the electronic microscopy experiments; MM undertook the BTA1 expression in the ER, designed the cloning strategy and produced stable transgenic *A. thaliana* lines; JJ designed and supervised the work and wrote the manuscript; and all authors revised the manuscript.

## Conflict of interest

The authors declare no conflict of interest.

## Funding

SS was supported by an INRAE PhD fellowship. JJ was supported by Blink ANR (ANR-18-CE92-0015). The LIPANG (Lipid analysis in Grenoble) platform, utilized in this work, is supported by the Rhône-Alpes Region, the funds FEDER and GRAL, and is further financed within the University Grenoble Alpes graduate school (Ecoles Universitaires de Recherche) CBH-EUR-GS (ANR-17-EURE-0003). We thank the microscopy facility MuLife of IRIG/DBSCI, funded by CEA Nanobio and GRAL LabEX (ANR-10-LABX-49-01) also financed within the University Grenoble Alpes graduate school CBH-EUR-GS (ANR-17-EURE-0003).

## Data availability

All primary data are included in this paper.

## References

Abida H, Dolch L-J, Meï C, et al. 2015. Membrane glycerolipid remodeling triggered by nitrogen and phosphorus starvation in *Phaeodactylum tricornutum*. Plant Physiology **167**, 118–136.

Andersson MX, Larsson KE, Tjellström H, Liljenberg C, Sandelius AS. 2005. Phosphate-limited oat. The plasma membrane and the tonoplast as major targets for phospholipid-to-glycolipid replacement and stimulation of phospholipases in the plasma membrane. The Journal of Biological Chemistry **280**, 27578–27586.

Billey E, Magneschi L, Leterme S, et al. 2021. Characterization of the Bubblegum acyl-CoA synthetase of *Microchloropsis gaditana*. Plant Physiology **185**, 815–835.

Bolik S, Albrieux C, Schneck E, Demé B, Jouhet J. 2022. Sulfoquinovosyldiacylglycerol and phosphatidylglycerol bilayers share biophysical properties and are good mutual substitutes in photosynthetic membranes. Biochimica et Biophysica Acta (BBA) - Biomembranes **1864**, 184037.

Bolik S, Schlaich A, Mukhina T, Amato A, Bastien O, Schneck E, Demé B, Jouhet J. 2023. Lipid bilayer properties potentially contributed to the evolutionary disappearance of betaine lipids in seed plants. BMC Biology **21**, 275.

Buseman CM, Tamura P, Sparks AA, et al. 2006. Wounding stimulates the accumulation of glycerolipids containing oxophytodienoic acid and dinor-oxophytodienoic acid in Arabidopsis leaves. Plant Physiology **142**, 28–39.

Cañavate JP, Armada I, Hachero-Cruzado I. 2017. Interspecific variability in phosphorus-induced lipid remodelling among marine eukaryotic phytoplankton. New Phytologist **213**, 700–713.

Cañavate JP, Armada I, Ríos JL, Hachero-Cruzado I. 2016. Exploring occurrence and molecular diversity of betaine lipids across taxonomy of marine microalgae. Phytochemistry **124**, 68–78.

Chi X, Zhang X, Guan X, Ding L, Li Y, Wang M, Lin H, Qin S. 2008. Fatty acid biosynthesis in eukaryotic photosynthetic microalgae: identification of a microsomal delta 12 desaturase in *Chlamydomonas reinhardtii*. Journal of Microbiology **46**, 189–201.

Cuyas L, David P, de Craieye D, et al. 2023. Identification and interest of molecular markers to monitor plant Pi status. BMC Plant Biology **23**, 401.

Dembitsky VM. 1996. Betaine ether-linked glycerolipids: chemistry and biology. Progress in Lipid Research **35**, 1–51.

Demé B, Cataye C, Block MA, Maréchal E, Jouhet J. 2014. Contribution of galactoglycerolipids to the 3-dimensional architecture of thylakoids. FASEB Journal **28**, 3373–3383.

Demski K, Łosiewska A, Jasieniecka-Gazarkiewicz K, Klińska S, Banaś A. 2020. Phospholipid:diacylglycerol acyltransferase1 overexpression delays senescence and enhances post-heat and cold exposure fitness. Frontiers in Plant Science **11**, 611897.

Eichenberger W, Araki S, Müller DG. 1993. Betaine lipids and phospholipids in brown algae. Phytochemistry **34**, 1323–1333.

Flori S, Jouneau P-H, Gallet B, Estrozi LF, Moriscot C, Schoehn G, Finazzi G, Falconet D. 2018. Imaging plastids in 2D and 3D: confocal and electron microscopy. In: Maréchal E, ed. Plastids: methods and protocols. New York, NY: Springer US, 113–122.

Gibson DG, Young L, Chuang R-Y, Venter JC, Hutchison CA, Smith HO. 2009. Enzymatic assembly of DNA molecules up to several hundred kilobases. Nature Methods **6**, 343–345.

Hamel L-P, Tardif R, Poirier-Gravel F, et al. 2024. Molecular responses of agroinfiltrated *Nicotiana benthamiana* leaves expressing suppressor of silencing P19 and influenza virus-like particles. Plant Biotechnology Journal **22**, 1078–1100.
Hoffmann DY, Shachar-Hill Y. 2023. Do betaine lipids replace phosphatidylcholine as fatty acid editing hubs in microalgae? Frontiers in Plant Science **14**, 1077347.

Hölzl G, Dörmann P. 2019. Chloroplast lipids and their biosynthesis. Annual Review of Plant Biology **70**, 51–81.

Jouhet J, Lupette J, Clerc O, Magneschi L, Bedhomme M, Collin S, Roy S, Maréchal E, Rébeillé F. 2017. LC-MS/MS versus TLC plus GC methods: consistency of glycerolipid and fatty acid profiles in microalgae and higher plant cells and effect of a nitrogen starvation. PLoS One **12**, e0182423.

Jouhet J, Maréchal E, Baldan B, Bligny R, Joyard J, Block MA. 2004. Phosphate deprivation induces transfer of DGDG galactolipid from chloroplast to mitochondria. The Journal of Cell Biology **167**, 863–874.

Jouhet J, Maréchal E, Bligny R, Joyard J, Block MA. 2003. Transient increase of phosphatidylcholine in plant cells in response to phosphate deprivation. FEBS Letters **544**, 63–68.

Kalisch B, Dörmann P, Hölzl G. 2016. DGDG and glycolipids in plants and algae. In: Nakamura Y, Li-Beisson Y, eds. Lipids in plant and algae development. Cham: Springer International Publishing, 51–83.

Kanduč M, Schlaich A, de Vries AH, Jouhet J, Maréchal E, Demé B, Netz RR, Schneck E. 2017. Tight cohesion between glycolipid membranes results from balanced water-headgroup interactions. Nature Communications **8**, 14899.

Karki N, Johnson BS, Bates PD. 2019. Metabolically distinct pools of phosphatidylcholine are involved in trafficking of fatty acids out of and into the chloroplast for membrane production. The Plant Cell **31**, 2768–2788.

Kato M, Sakai M, Adachi K, Ikemoto H, Sano H. 1996. Distribution of betaine lipids in marine algae. Phytochemistry **42**, 1341–1345.

Künzler K, Eichenberger W. 1997. Betaine lipids and zwitterionic phospholipids in plants and fungi. Phytochemistry **46**, 883–892.

Künzler K, Eichenberger W, Radunz A. 1997. Intracellular localization of two betaine lipids by cell fractionation and immunomicroscopy. Zeitschrift fur Naturforschung C: Journal of Biosciences **52**, 487–495.

Li-Beisson Y, Beisson F, Riekhof W. 2015. Metabolism of acyl-lipids in *Chlamydomonas reinhardtii*. The Plant Journal **82**, 504–522.

Makewicz A, Gribi C, Eichenberger W. 1997. Lipids of *Ectocarpus fasciculatus* (Phaeophyceae). Incorporation of [I-¹⁴C]Oleate and the role of TAG and MGDG in lipid metabolism. Plant and Cell Physiology **38**, 952–962.

Maxwell K, Johnson GN. 2000. Chlorophyll fluorescence—a practical guide. Journal of Experimental Botany **51**, 659–668.

Meï C, Michaud M, Cussac M, Albrieux C, Gros V, Maréchal E, Block MA, Jouhet J, Rébeillé F. 2015. Levels of polyunsaturated fatty acids correlate with growth rate in plant cell cultures. Scientific Reports **5**, 15207.

Mühlroth A, Winge P, El Assimi A, Jouhet J, Maréchal E, Hohmann-Marriott MF, Vadstein O, Bones AM. 2017. Mechanisms of phosphorus acquisition and lipid class remodeling under P limitation in a marine microalga. Plant Physiology **175**, 1543–1559.

Murakami H, Nobusawa T, Hori K, Shimojima M, Ohta H. 2018. Betaine lipid is crucial for adapting to low temperature and phosphate deficiency in *Nannochloropsis*. Plant Physiology **177**, 181–193.

Nakamura Y. 2021. Headgroup biosynthesis of phosphatidylcholine and phosphatidylethanolamine in seed plants. Progress in Lipid Research **82**, 101091.

Nakamura Y, Koizumi R, Shui G, Shimojima M, Wenk MR, Ito T, Ohta H. 2009. Arabidopsis lipins mediate eukaryotic pathway of lipid metabolism and cope critically with phosphate starvation. Proceedings of the National Academy of Sciences, USA **106**, 20978–20983.

Nelson NO, Janke RR. 2007. Phosphorus sources and management in organic production systems. HortTechnology **17**, 442–454.

Oishi Y, Otaki R, Iijima Y, Kumagai E, Aoki M, Tsuzuki M, Fujiwara S, Sato N. 2022. Diacylglycerol-*N*,*N*,*N*-trimethylhomoserine-dependent lipid remodeling in a green alga, *Chlorella kessleri*. Communications Biology **5**, 19.

Oliver MJ, Tuba Z, Mishler BD. 2000. The evolution of vegetative desiccation tolerance in land plants. Plant Ecology **151**, 85–100.

Petroutsos D, Amiar S, Abida H, et al. 2014. Evolution of galactoglycerolipid biosynthetic pathways--from cyanobacteria to primary plastids and from primary to secondary plastids. Progress in Lipid Research **54**, 68–85.

Poirier Y, Bucher M. 2002. Phosphate transport and homeostasis in Arabidopsis. The Arabidopsis Book **1**, e0024.

Poirier Y, Thoma S, Somerville C, Schiefelbein J. 1991. Mutant of Arabidopsis deficient in Xylem loading of phosphate 1. Plant Physiology **97**, 1087–1093.

Rainteau D, Humbert L, Delage E, et al. 2012. Acyl chains of Phospholipase D transphosphatidylation products in arabidopsis cells: a study using multiple reaction monitoring mass spectrometry. PLoS One **7**, e41985.

Riekhof WR, Andre C, Benning C. 2005a. Two enzymes, BtaA and BtaB, are sufficient for betaine lipid biosynthesis in bacteria. Archives of Biochemistry and Biophysics **441**, 96–105.

Riekhof WR, Naik S, Bertrand H, Benning C, Voelker DR. 2014. Phosphate Starvation in Fungi Induces the Replacement of Phosphatidylcholine with the phosphorus-free betaine lipid diacylglycerol-*N*,*N*,*N*-trimethylhomoserine. Eukaryotic Cell **13**, 749–757.

Riekhof WR, Sears BB, Benning C. 2005b. Annotation of genes involved in glycerolipid biosynthesis in *Chlamydomonas reinhardtii*: discovery of the betaine lipid synthase BTA1Cr. Eukaryotic Cell **4**, 242–252.

Rozentsvet OA. 2004. Comparative examination of distribution of phospholipids and a betaine lipid DGTS in tropical fern species. Biochemical Systematics and Ecology **32**, 303–311.

Sato N. 1991. Lipids in *Cryptomonas* CR-1. II. Biosynthesis of betaine lipids and galactolipids. Plant and Cell Physiology **32**, 845–851.

Sato N. 1992. Betaine lipids. The Botanical Magazine Tokyo **105**, 185–197.

Shimojima M, Madoka Y, Fujiwara R, Murakawa M, Yoshitake Y, Ikeda K, Koizumi R, Endo K, Ozaki K, Ohta H. 2015. An engineered lipid remodeling system using a galactolipid synthase promoter during phosphate starvation enhances oil accumulation in plants. Frontiers in Plant Science **6**, 664.

Simionato D, Block MA, La Rocca N, Jouhet J, Maréchal E, Finazzi G, Morosinotto T. 2013. The response of *Nannochloropsis gaditana* to nitrogen starvation includes *de novo* biosynthesis of triacylglycerols, a decrease of chloroplast galactolipids, and reorganization of the photosynthetic apparatus. Eukaryotic Cell **12**, 665–676.

Smit AL, Bindraban PS, Schröder JJ, Conijn JG, Meer HG. 2009. Phosphorus in agriculture: global resources, trends and developments. Wageningen: Plant Research International B.V., 1–33.

Tiessen H. 2008. Phosphorus in the global environment. In: White PJ, Hammond JP, eds. The ecophysiology of plant-phosphorus interactions. Dordrecht: Springer Netherlands, 1–7.

To A, Valon C, Savino G, Guilleminot J, Devic M, Giraudat J, Parcy F. 2006. A network of local and redundant gene regulation governs Arabidopsis seed maturation. The Plant Cell **18**, 1642–1651.

Uwizeye C, Decelle J, Jouneau P-H, et al. 2021. Morphological bases of phytoplankton energy management and physiological responses unveiled by 3D subcellular imaging. Nature Communications **12**, 1049.

Vanhercke T, Dyer JM, Mullen RT, Kilaru A, Rahman MM, Petrie JR, Green AG, Yurchenko O, Singh SP. 2019. Metabolic engineering for enhanced oil in biomass. Progress in Lipid Research **74**, 103–129.

Van Mooy BAS, Fredricks HF, Pedler BE, et al. 2009. Phytoplankton in the ocean use non-phosphorus lipids in response to phosphorus scarcity. Nature **458**, 69–72.

Vaskovsky VE, Khotimchenko SV, Boolugh EM. 1998. Distribution of diacylglycerotrimethylhomoserine and phosphatidylcholine in mushrooms. Phytochemistry **47**, 755–760.

Vitousek PM, Naylor R, Crews T, et al. 2009. Nutrient imbalances in agricultural development. Science **324**, 1519–1520.
**Vogel G, Eichenberger W.** 1992. Betaine lipids in lower plants. Biosynthesis of DGTS and DGTA in *Ochromonas danica* (Chrysophyceae) and the possible role of DGTS in lipid metabolism. *Plant and Cell Physiology* **33**, 427–436.

**Yang W, Moroney JV, Moore TS.** 2004. Membrane lipid biosynthesis in *Chlamydomonas reinhardtii*: ethanolaminephosphotransferase is capable of synthesizing both phosphatidylcholine and phosphatidylethanolamine. *Archives of Biochemistry and Biophysics* **430**, 198–209.

**Yin C, Andersson MX, Zhang H, Aronsson H.** 2015. Phosphatidylcholine is transferred from chemically-defined liposomes to chloroplasts through

proteins of the chloroplast outer envelope membrane. FEBS Letters **589**, 177–181.

**Zhang Y, He J, Jia L-J, Yuan T-L, Zhang D, Guo Y, Wang Y, Tang W-H.** 2016. Cellular tracking and gene profiling of *Fusarium graminearum* during maize stalk rot disease development elucidates its strategies in confronting phosphorus limitation in the host apoplast. *PLoS Pathogens* **12**, e1005485.

**Zhou X-R, Bhandari S, Johnson BS, Kotapati HK, Allen DK, Vanhercke T, Bates PD.** 2020. Reorganization of acyl flux through the lipid metabolic network in oil-accumulating tobacco leaves. *Plant Physiology* **182**, 739–755.