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. 2026 May 12;251(3):1311–1328. doi: 10.1111/nph.71257

The algal homolog of the plant CER1 and CER3 proteins is a bifunctional hydrocarbon‐forming enzyme

Ángel Baca‐Porcel 1, Bertrand Légeret 1, Alexandre Six 1, Mallaury Cabanel 1, Mathilde Le Cossec 1, Arthur Guegano 1, Damien Sorigué 1, Yonghua Li‐Beisson 1, Florian Veillet 1,✉, Fred Beisson 1,✉
PMCID: PMC13326499  PMID: 42124393

Summary

  • The ability to synthesize and secrete hydrophobic compounds is believed to have been a pivotal event in the evolution of land plants from their aquatic green algal ancestors. The key to biosynthesis of plant surface alkanes is a heterodimeric complex consisting of two homologous membrane‐bound proteins, ECERIFERUM 3 (CER3) and ECERIFERUM 1 (CER1), which bear distinct enzyme activities. A single homolog of CER1 and CER3, referred to as CER1/3, has long been identified in some algae. However, it has remained unknown whether CER1/3 exhibits CER1 and/or CER3 activity or another ancestral activity.

  • Here we investigate the function of CER1/3 by using CRISPR‐Cas9‐mediated knockout mutants in the early‐branching chlorophyte Ostreococcus tauri and by yeast heterologous expression.

  • Genome mining shows that in green algal genomes the presence of CER1/3 is correlated with the absence of fatty acid photodecarboxylase. Knockouts provide evidence that CER1/3 is necessary for synthesizing a C21:6 alkene in Ostreococcus. Yeast expression experiments demonstrate that algal CER1/3 are bifunctional enzymes with aldehyde‐ and hydrocarbon‐forming domains, corresponding to CER3 and CER1 activities, respectively.

  • These findings support the idea that the land plant alkane‐forming CER1/CER3 complex evolved from a bifunctional hydrocarbon‐forming CER1/3 enzyme found in some of the earliest‐diverging green algal lineages.

Keywords: CER1, CER3, green algae, hydrocarbons, Ostreococcus tauri, terrestrialization

Schematic representation of the proposed reactions carried out by a CER1/3 protein from green algae. CoA, coenzyme A; CTD, C‐terminal domain; Cys, catalytic cysteine of C‐terminal domain; His, catalytic histidines of N‐terminal domain; NTD, N‐terminal domain. Only one monomer is represented, but the CER1/3 protein very likely exists as a homodimer.

graphic file with name NPH-251-1311-g001.jpg

Introduction

The colonization of terrestrial habitats by land plants has had a profound impact on the evolution of organisms and ecosystems on Earth (Bowman, 2022). This event greatly affected biogeochemical cycles, facilitating the terrestrialization of various other lineages. Land plants probably evolved from a single clade of streptophyte algae c. 470 million years ago (Ma) (Delwiche & Cooper, 2015; De Vries & Archibald, 2018). The transition from aquatic to terrestrial environments was driven by key cellular and physiological adaptations to the new environmental constraints associated with terrestrial habitats, such as desiccation, ultraviolet radiation, and pathogens. Symbiotic interactions with fungal partners are also believed to have been crucial for the earliest land plants (Delaux & Schornack, 2021).

Limiting losses of internal water was necessarily a key adaptation to terrestrial environments. This was notably achieved by building an extracellular lipid‐based structure, which is absent in algae: the cuticle (Delwiche & Cooper, 2015). In land plants, cuticle components are synthesized in the epidermal cells of aerial organs and subsequently exported onto their surface to form a waterproof layer (Yeats & Rose, 2013; Delude et al., 2016). The cuticle is made of a cutin scaffold filled and covered with cuticular waxes. Cutin is a polyester that consists mainly of hydroxy‐, epoxy‐, and dicarboxy C16‐ and C18‐fatty acids, together with glycerol (Pollard et al., 2008; Beisson et al., 2012; Kosma et al., 2025). However, it is the waxes rather than the cutin itself that confer impermeability to the cuticle. The cuticular waxes of many plants consist mainly of very‐long‐chain (VLC, > C20) aliphatic compounds (Samuels et al., 2008; Bernard & Joubès, 2013; Lee & Suh, 2015). The C16 and C18 acyl‐CoAs are substrates for the fatty acid elongase complex, which produces VLC acyl‐CoAs up to C30 and beyond (Haslam & Kunst, 2013). Following elongation, VLC acyl‐CoAs are converted into cuticular waxes via two major pathways: the alcohol‐forming pathway, yielding primary alcohols and wax esters, and the alkane‐forming pathway, which produces mainly alkanes, aldehydes, secondary alcohols, and ketones. These two pathways account for c. 20 and 80%, respectively, of the total stem wax load in Arabidopsis (Jenks et al., 1995; Li‐Beisson et al., 2013). All these wax components are then exported to the outer cell wall of epidermal cells by ATP‐binding cassette transporters of the ABCG subfamily, enabling subsequent assembly into intracuticular and epicuticular waxes (Lee & Suh, 2015). In many plant species, alkanes are the major component of cuticular waxes (Jetter et al., 2006). Alkanes are synthesized in the endoplasmic reticulum (ER) from VLC acyl‐CoAs by the ECERIFERUM proteins CER1 (Aarts et al., 1995) and CER3 (Rowland et al., 2007), which form a membrane complex with the cytochrome B5 (Bernard et al., 2012).

The CER1 and CER3 proteins of land plants are homologous and possess a bi‐domain structure consisting of an N‐terminal fatty acid hydroxylase domain, resembling the catalytic domain of the yeast sphingolipid α‐hydroxylase Scs7, and a C‐terminal WAX2 domain, which resembles the cleft region of Synechococcus elongatus acyl‐ACP reductase, an aldehyde‐producing enzyme (Wang et al., 2019; Chaudhary et al., 2021; Kojima et al., 2024). The N‐terminal domain of plant CER1 is highly conserved and possesses the histidine‐rich motifs necessary for the formation of alkanes by the CER1/CER3 complex (Bernard et al., 2012; Kojima et al., 2024), while its C‐terminus is less conserved. Conversely, the C‐terminal domain of plant CER3 is highly conserved. It contains a potentially catalytic cysteine that is likely to form a covalent bond with the acyl group (Gao et al., 2020), and a putative NADPH‐binding site, which may be involved in NADPH utilization (Kojima et al., 2024). The N‐terminal domain of plant CER3 is less conserved, and the histidine‐rich motifs present are not essential for the activity of the CER1/CER3 complex (Bernard et al., 2012; Kojima et al., 2024).

It has long been hypothesized that CER3 is a reductase that mediates the reduction of Cn VLC acyl‐CoAs into Cn aldehyde intermediates, and that CER1 is an aldehyde decarbonylase that produces the Cn‐1 alkanes (Jenks et al., 1995; Bernard et al., 2012; Pascal et al., 2019). A recent landmark study in Arabidopsis provides indirect evidence of an aldehyde‐forming activity of CER3. This study shows that CER3 also interacts with Smart‐OH1 (SOH1), a reductase that converts aldehydes into primary alcohols (Li et al., 2025). The CER1‐CER3‐SOH1 complex was proposed to participate in the regulation of transpiration through the cuticle by modulating the alkane/primary alcohol ratio of cuticular waxes in response to environmental conditions.

Given the importance of alkanes in plant cuticular waxes, the appearance of the CER1 and CER3 proteins in land plants was likely to be key to cuticle evolution and terrestrialization. Interestingly, a few algal species, such as the marine chlorophyte Ostreococcus tauri (O. tauri), possess a CER1/CER3 homolog (Sorigué et al., 2016), but always as a single‐copy gene referred to as CER1/3. Phylogenetic analyses suggest that this CER1/3 algal homolog underwent duplication in ancestral embryophytes to produce the paralogs CER1 and CER3 (Wang et al., 2019; Chaudhary et al., 2021). However, the biochemical function of the CER1/3 enzymes in the green algal lineage has not yet been examined.

In algae, the presence of VLC alkenes likely to be derived from VLC fatty acids has been known for decades in diatoms (Lee & Loeblich, 1971) and was later found in the excreted material of the green colonial microalga Botryococcus braunii (Metzger & Largeau, 2005). More recently, it has been shown in the model green microalgae Chlamydomonas reinhardtii and Chlorella variabilis NC64A that C15‐C17 alkanes and alkenes are synthesized from LC fatty acids by a light‐dependent mechanism (Sorigué et al., 2016), and the enzyme responsible for this synthesis has been identified (Sorigué et al., 2017). It is an algal‐specific photoenzyme, which converts free fatty acids into HCs in the presence of light (350–520 nm) and has been termed fatty acid photodecarboxylase (FAP). In C. reinhardtii, the FAP is predicted to be a chloroplastic enzyme, and it has been demonstrated that the alkene product of the FAP is mostly located in the thylakoid fraction (Moulin et al., 2021). The FAP activity was apparently lost in plants, but a putative FAP is present in the genome of many algae from various lineages, including some streptophyte algae, and the FAP activity was indeed shown to be conserved in several algal lineages (Moulin et al., 2021).

It has remained unknown whether the unique CER1/3 homolog in algae represents an additional system that forms HCs or bears another activity. Here, we show that CER1/3 and FAP appear to be mutually exclusive in hundred or so green algal genomes. We also demonstrate that the CER1/3 protein found in various green algal species is a bifunctional enzyme that can efficiently synthesize hydrocarbons (HCs) by producing aldehydes and converting them into HCs. These results provide experimental support to the view that an ancestral algal bifunctional CER1/3 protein has evolved into a plant CER1/CER3 protein complex, in which each original function is performed by a specialized homolog. The possible redundant or distinct roles of HCs derived from FAP and from CER1/3 are discussed.

Materials and Methods

Bioinformatic analyses

To identify algal homologs to plant CER1/CER3 proteins, BlastP searches were performed across various databases using the Arabidopsis thaliana CER1 as query protein sequence. The 67 sequences of the various algal CER1/3 and plant CER1 and CER3 proteins used for phylogenetic analysis are available in Supporting Information Table S1. These sequences were mostly selected from species with fully sequenced genomes, ensuring broad representation of the genetic diversity across green algae and land plants. For the construction of a maximum likelihood phylogenetic tree, protein sequences were first aligned with Mafft (https://www.ebi.ac.uk/jdispatcher/msa/mafft) using default parameters (Blosum 62 matrix, gap open penalty 1.53, gap extension penalty 0.123). The alignments were then trimmed in SeaView 5 using Gblocks (smaller blocks, gaps within the final block and less strict flanking positions allowed). Finally, the tree was constructed with the IQ‐TREE algorithm (Nguyen et al., 2015) using Model Finder (Kalyaanamoorthy et al., 2017) and 1000 ultrafast bootstrap approximation (Hoang et al., 2018). The best‐fit model according to the Bayesian Information Criterion was LG + R5. For protein 3D structure prediction and modeling, the AlphaFold 3 software was used (Abramson et al., 2024). The generated protein 3D structures were visualized using PyMOL software (Delano Scientific LLC). For the prediction of protein subcellular localization, the DeepLoc‐2.1 online software was used (Nielsen, 2025).

Algal strains and culture conditions

The Ostreococcus tauri RCC4221 strain was obtained from the Roscoff Culture Collection. This strain was cultured in sterile artificial seawater supplemented with F/2 medium (Sigma‐Aldrich). Cultures were maintained in T‐25 flasks with vented filter caps (CytoOne) at 22°C under an illumination of c. 50 μmol photons m−2 s−1 with a 12 h :12 h light : dark photoperiod. Cell counts were routinely performed using a Multisizer 3 (Coulter).

The Klebsormidium nitens (K. nitens) NIES‐2285 strain was obtained from the microbial culture collection of the National Institute of Environmental Studies, Japan. This strain was cultured in plates containing liquid or solid (2% agar) C medium, incubated at 22°C under c. 50 μmol photons m−2 s−1 with a 12 h :12 h light : dark photoperiod.

The Auxenochlorella UTEX 250‐A strain was obtained from the Merchant lab (Camacho et al., 2025; Craig et al., 2025). This strain was cultured in ApM1 medium at 25°C under c. 125 μmol photons m−2 s−1 with a 12 h :12 h light : dark photoperiod at 2% CO2.

Plasmid construction and yeast expression

Protein sequences from various algal CER1/3 and land plant CER1/CER3 (Table S1) were back‐translated using yeast codon optimization (Geneious software) and synthesized (TwistBioscience, San Francisco, CA, USA) for cloning into the Golden Gate MoClo system. Site‐specific mutations in the coding sequences of the different CER1, CER3, and CER1/3 proteins were introduced by PCR. Each coding sequence was assembled in a transcriptional unit under the yeast copper‐inducible promoter (pCUP1) and an appropriate terminator sequence (i.e. tPGK1, tTDH1, and tADH1) (Lee et al., 2015). Final plasmids were constructed by assembling the high‐copy 2micron yeast origin of replication and the URA3 auxotrophic selection marker (Lee et al., 2015) with at least one transcriptional unit mediating the expression of algal CER1/3 and/or land plant CER1 and CER3. Final plasmids were introduced into the Saccharomyces cerevisiae INVSc1 (ThermoFisher Scientific, Waltham, MA, USA) diploid strain (MATa his3D1 leu2 trp1‐289 ura3‐52 MAT his3D1 leu2 trp1‐289 ura3‐52) using the classic LiOAc/PEG3350 method. Yeast cells growing on plates filled with selective dropout media lacking uracil (DOM‐URA) were further checked by colony PCR. All the plasmids constructed and used in this work are detailed in Table S2. For protein expression, cultures (15 ml) were initiated at an optical density (OD600nm) of 0.2 in DOM‐URA media in 50‐ml mini bioreactor centrifuge tubes. Cells were grown in an orbital shaker at 30°C, 180 rpm. Protein expression was induced with 0.033 mg ml−1 CuSO4 4 h after starting the culture. The cultures were grown in these conditions for 48 h. The OD600nm of the different cultures was then measured before samples being collected for fatty acid and HC content analysis by GC‐MS/FID.

CRISPR‐Cas9‐mediated inactivation of the O. tauri CER1/3

The OtCER1/3 gene (ostta11g00770) was targeted using the CRISPR‐Cas9 system in order to promote the insertion of linear DNA encoding the AphI gene into the Cas9‐mediated cutting site. A 20‐bp guide RNA (5′‐ACATCAAGTGTTCCATGAAC‐3′) for the CRISPR‐SpCas9 system was designed to target the OtCER1/3 coding sequence using the CRISPOR (Concordet & Haeussler, 2018), CHOPCHOP (Labun et al., 2019), and CRISPR‐Cas9 guide RNA design checker (IDT) software. The guide RNA (Alt‐R™ CRISPR‐Cas9 crRNA) and the transactivating crRNA (Alt‐R™ CRISPR‐Cas9 tracrRNA) were synthesized (IDT) and assembled into a crRNA:tracrRNA duplex. This duplex was then mixed with the purified Cas9 enzyme (Cas9 expression plasmid: pET‐NLS‐Cas9‐6xHis) (Kim et al., 2024) to produce functional RNPs (with a guide RNA‐to‐Cas9 enzyme ratio of ≈ 2 : 1) (Table S3). The in vitro activity of the RNP was tested on PCR fragments (Table S3). Two constructs for the expression of the AphI gene which confers resistance to geneticin (G418) were built using the Golden Gate MoClo system: one was driven by the Otα‐tub (ostta12g01320) promoter and terminator, and the other by the OtUBI (ostta12g01170) promoter and terminator (Table S4), as previously described (Sanchez et al., 2019). The region of interest for both constructs was PCR amplified using the Platinum™ SuperFi II DNA Polymerase (ThermoFisher Scientific), and linear DNA was then purified using the NucleoSpin Gel and PCR Clean‐up (Macherey‐Nagel) followed by CH₃COO− Na+ precipitation. Delivery of RNPs and linear DNA into O. tauri cells was mostly performed as previously described (Sanchez et al., 2019). Briefly, the O. tauri strain RCC 4221 was grown in liquid ASW F/2 medium for 10–14 d at 22°C, 12 h : 12 h photoperiod (30–50 μE). Cells were washed twice in fresh ASW F/2 medium and resuspended at an optical density (OD600nm) of 20. For each transformation event, 250 μl of cells were mixed with RNPs, linear DNA, and 1 μl of salmon sperm DNA at 2 mg ml−1, and incubated at room temperature for a few minutes. An equal volume of PEG (4000 or 6000) 60% was added, mixed gently, and then the mixture was incubated at room temperature for 1 or 2 min (Table S3). The cells were then gently transferred into 19 ml of ASW F/2 medium and incubated for 24 h in the growth chamber. Cells were transferred into a semi‐solid medium consisting of ASW F/2 medium mixed with low‐melting agarose (Duchefa Biochemie, Haarlem, The Netherlands) at a final concentration of 0.4%, supplemented with 1 mg ml−1 geneticin. The plates were returned to the growth chamber for a few weeks, until green colonies could be isolated and grown in liquid ASW F/2 medium supplemented with 1 mg ml−1 geneticin for an extra week. The targeted insertion of the AphI cassette at the target OtCER1/3 genomic locus was detected by PCR amplification performed directly on the algal cells using the Platinum™ Direct PCR Universal Master Mix (ThermoFisher Scientific) and Sanger sequencing (Eurofins Genomics, Köln, Germany). Sequence alignments were conducted using the Geneious software. Primers used in this study for O. tauri genome editing experiments are listed in Table S5.

Analysis of intracellular HCs and fatty acids by transmethylation and saponification

For the initial identification of HCs in algae, saponification was used. Briefly, O. tauri and Auxenochlorella cell cultures were harvested after 8 and 4 d of culture, respectively. For K. nitens cells, 4‐wk‐old filaments were collected. The cells were centrifuged at 3200  g for 15 min to pellet the cells in 8‐ml glass tubes with teflon‐lined screw caps. One milliliter of NaOH 1 M was added, and the mixture was heated at 85°C for 1 h. The tubes were then allowed to reach room temperature; 250 μl of hexane was added, and the mixture was vortexed for 30 s. After centrifugation, part of the n‐hexane layer was collected and analyzed by GC‐MS/FID.

For the quantification of HCs and fatty acids in O. tauri, acid‐catalyzed transmethylation was used. Cell pellets were collected in 8‐ml glass tubes with teflon‐lined screw caps. The supernatant was discarded, and internal standards (10 μg of n‐tetracosane and 10 μg of trinonadecanoyl glycerol) were added together with 2 ml of methanol containing 5% (v/v) sulfuric acid. The mixture was heated at 85°C for 90 min. After cooling down to room temperature, 3 ml of 0.9% NaCl and 250 μl of n‐hexane were added. The samples were vortexed for 30 s, centrifuged, and part of the n‐hexane phase was collected and analyzed by GC‐MS/FID.

For the identification and quantification of HCs and fatty acids in yeast cells, direct transmethylation of cells was used. Briefly, 8 ml of cell culture at OD600nm 10 was centrifuged at 3200  g for 15 min to pellet the cells in 8‐ml glass tubes with teflon‐lined screw caps. The supernatant was discarded and internal standards (10 μg of n‐docosane and 10 μg of triheptadecanoyl glycerol) as well as 2.5 ml of methanol containing 5% (v/v) sulfuric acid were added to cell pellets. The samples were then treated and analyzed in the same way as for the algal cells, except that 200 μl of n‐hexane was used for extraction.

Synthesis of the C21:6 alkene standard

For the synthesis of the C21:6 alkene standard (all‐cis‐3,6,9,12,15,18‐heneicosahexaene) derived from all‐cis‐4,7,10,13,16,19‐docosahexaenoic acid (DHA), 100 μl of a 2.5 mg ml−1 DHA solution in ethanol and 200 μl of a 10 mg ml−1 purified recombinant FAP protein solution (Sorigué et al., 2017) from Chlorella variabilis NC64A in a buffer (150 mM NaCl, 10 mM Tris–HCl pH 8.0 with 20% glycerol) were added to a 10 ml sealed vial containing 1 ml of 50 mM Tris–HCl buffer pH 8.0. The vial was then incubated at room temperature for 1 h under 300 μmol photons m−2 s−1 of blue light to trigger the decarboxylation of DHA into C21:6 alkene by FAP. To extract the alkene, 900 μl of the aqueous reaction mixture was subjected to saponification with NaOH 1 M final. After saponification at 85°C for 1 h, the alkene was extracted by adding 500 μl of n‐hexane, followed by 5 min shaking and centrifugation at 3200  g for 2 min to facilitate phase separation. Finally, the n‐hexane phase was analyzed by GC‐MS/FID to check purity.

GC‐MS/FID analyses

For the analysis of algae and yeast fatty acid methyl esters (FAMEs) samples, an OPTIMA WAXplus column and an HP‐5MS column were used, respectively (length 30 m, internal diameter 0.25 mm, and film thickness 0.25 mm for both columns). For the analysis of the unsaponifiable material, the HP‐5MS column was used. Columns were mounted on a GC‐FID 7890B Agilent coupled to a 5977B series mass detector. Helium (1.4 ml min−1) was used as the carrier gas. One microliter of the n‐hexane phase was injected in splitless mode. The GC parameters were as follows for the HP‐5MS column: oven initial temperature, 50°C for 1 min; ramp, 10°C min−1 to 150°C and then 5°C min−1 to 310°C; hold for 8 min. Idem for the WAX column except the final temperature was 260°C for 5 min. The MS was run in full scan over 40–500 a.m.u. (electron impact ionization at 70 eV). Peaks were identified based on their retention time and mass spectrum and quantified based on the FID signal using the internal standards.

Protein–protein interaction analyses

A split‐ubiquitin yeast two‐hybrid (SUY2H) assay was performed as previously described (Bernard et al., 2012; Li et al., 2025). The OtCER1/3 coding sequence was cloned into the pBT3N bait vector using the SfiI restriction enzyme (ThermoFisher Scientific), while the OtCER1/3, AtCER1, and AtCER3 coding sequences were cloned into the pPR3N prey vector. Constructs were then introduced into the NMY51 yeast strain, and transformants were selected on plates filled with dropout media lacking leucine and/or tryptophan. Colonies were further checked by colony PCR. Interactions were assayed on dropout media lacking leucine, tryptophan, histidine, and adenine.

Statistical analyses

Data processing and statistical analyses were performed using the GraphPad Prism software. Statistical tests used and the corresponding statistical significance are indicated in the figure legends.

Results

Green algae species that possess CER1/3 lack fatty acid photodecarboxylase (FAP)

Protein sequence similarity searches in various databases using the Arabidopsis CER1 sequence as a bait allowed to identify single CER1/CER3 homologs (hereafter referred to as CER1/3) in 20 out of 99 genomes of green algae investigated (Tables 1, S6). Green algal CER1/3 proteins were present across the three phyla of the green lineage (Li et al., 2020): prasinodermophyte algae, chlorophyte algae, and streptophyte algae. Additionally, a CER1/3 homolog was also identified in the genomes of the cryptophyta algae Proteomonas sulcata and Cryptophyceae sp., both of which result from a secondary endosymbiosis. The presence of CER1/3 proteins in green algal species correlated with the lack of the plastidial FAP photoenzyme (Table 1). Some species belonging to the genus Auxenochlorella, Elliptochloris, or Helicosporidium lacked both FAP and CER1/3 homologs.

Table 1.

Presence of FAP and CER1/CER3 homologs in green algal genomes.

Phylum Class Genus No. of species or strains investigated No. of FAP genes per genome No. of CER1/CER3 homologs per genome
Prasinodermophytes Prasinodermophyceae Prasinoderma 1 0 1
Chlorophytes Mamiellophyceae Bathycoccus 1 0 1
Micromonas 2 0 1
Ostreococcus 4 0 1
Trebouxiophyceae 1 2 0
Asterochloris 1 1 0
Auxenochlorella 2 0 0
Botryococcus 1 1 0
Chlorella 5 1 0
Coccomyxa 2 1 0
Elliptochloris 1 0 0
Helicosporidium 1 0 0
Micractinium 1 1 0
Myrmecia 1 1 0
Nannochloris 2 1 0
Picochlorum 1 1 0
Symbiochloris 5 1 0
Trebouxia 1 1 0
(genus incerta) 4 1 0
Pedinophyceae Pedinomonas 1 1 0
(genus incerta) 1 1 0
Chlorodendrophyceae Tetraselmis 1 0 1
Chloropicophyceae Chloropicon 1 1 0
Picocystophyceae Picocystis 1 1 0
Chlorophyceae Astrephomene 1 1 0
Chlamydomonas 4 1 or 0 0
Chloromonas 1 2 0
Chromochloris 1 1 0
Desmodesmus 2 1 0
Dunaliella 1 1 0
Enallax 1 1 0
Flechtneria 1 1 0
Gonium 1 1 0
Haematococcus 1 1 0
Limnomonas 1 2 0
Monoraphidium 2 1 or 0 0
Pleodorina 3 1 0
Raphidocelis 1 1 0
Scenedesmus 12 1, 2 or 3 0
Tetrabaena 1 1 0
Tetradesmus 2 1 or 2 0
Volvox 4 1 0
Ulvophyceae Ulva mutabilis 1 1 0
Caulerpa 1 1 0
Ostreobium 1 1 0
Streptophytes Mesostigmatophyceae Mesostigma 2 1 0
Chlorokybophyceae Chlorokybus 1 1 0
Klebsormidiophyceae Klebsormidium 1 0 1
Charophyceae Chara 1 0 1
Zygnematophyceae Spirogloea 1 0 3
Spirogyra 1 0 1
Zygnema 4 0 1
Mesotaenium 3 0 1
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This table shows the genera and the number of species or strains whose genomes were examined for the presence of FAP and CER1/CER3 homologs. The ploidy level may have an effect on the number of genes for each genome. The full names of the species or strains used are listed in Supporting Information Table S6. The shading colour indicates the presence of either FAP (green) or CER1/3 (blue) gene(s).

Multiple alignment of algal CER1/3 proteins with land plant homologs showed that, based on conserved motifs (see the fourth section below), some species of the land plant Marchantiophytes, Briophyta, Lycophytes and Monolophytes contained, in addition to the proteins similar to the CER1/CER3 proteins of Angiosperms, some homologs that shared more structural motifs with the algal CER1/3 proteins. These land plant proteins were thus also annotated as putative CER1/3 and a phylogenetic analysis was conducted (Fig. 1). The phylogenetic tree showed that three groups could be clearly distinguished, CER1, CER3 and CER1/3. This analysis supports the view that the algal CER1/3 gene was duplicated into CER1 and CER3 genes in the common ancestor of land plants, as previously suggested based on a limited number of algal CER1/3 proteins (Chaudhary et al., 2021). However, the internal structure of each group remains uncertain due to multiple duplications (CER1 and CER3 groups) or a limited number of sequences available in algae (CER1/3 group).

Fig. 1.

Fig. 1

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Phylogenetic tree of ECERIFERUM 1 (CER1), ECERIFERUM 3 (CER3), and CER1/3 homologs in land plants and green algae. The tree was constructed using the maximum likelihood method based on 67 protein sequences from various plants and algae (see Supporting Information Table S1). Homologs were classified as CER1, CER3, or CER1/3 before tree building based on conserved motifs (see Fig. 4). The tree was rooted with the Prasinoderma coloniale CER1/3 homolog (Prasinodermophytes). Bootstrap support values from 1000 replicates are indicated. The scale bar represents 0.2 substitutions per site.

Taken together, these results indicate that the green algal CER1/3 proteins may perform the same reaction as CER1/CER3 and could potentially fulfill the same biological role as FAP.

Ostreococcus tauri and K. nitens produce C21:6 and C17:0 hydrocarbons respectively

We then assessed whether the streptophyte alga K. nitens (Klebsormidiophyceae) and the early‐branching chlorophyte alga O. tauri (Mamiellophyceae), which possess a CER1/3 protein but lack the FAP, were able to synthesize HCs that may be derived from fatty acids. We also looked in Auxenochlorella, which lacks both FAP and CER1/3. In the latter species, we could not detect any alkane or alkene but in the unsaponifiable fraction of K. nitens cells, we detected a C17 alkane (Fig. S1), a common HC found in various algae. Under our conditions, no other alkane could be found in K. nitens, including the C22 alkane reported before in this species (Kondo et al., 2016). In O. tauri cells, we show that the GC peak present in the total fatty acid analysis (Fig. 2a), which previously suggested to be a C21:6 alkene derived from DHA fatty acid (Sorigué et al., 2016), was also present in the unsaponifiable material (i.e. after removal of fatty acids) as expected for a HC (Fig. S2). To confirm the identity of the putative C21:6 alkene, we prepared an authentic C21:6 alkene standard by enzymatic decarboxylation of a pure standard of DHA fatty acid (see material and methods). The putative C21:6 alkene had indeed the same retention time and mass spectrum (Fig. 2) as the C21:6 n‐alkene FAP product (all‐cis‐3,6,9,12,15,18‐heneicosahexaene). The C21:6 alkene was the only HC that could be detected in O. tauri.

Fig. 2.

Fig. 2

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Synthesis of C21:6 alkene derived from all‐cis‐4,7,10,13,16,19‐docosahexaenoic acid (DHA) by Ostreococcus tauri RCC4221 cells. (a) Portion of the GC chromatogram showing the putative C21:6 alkene peak in the analysis of total fatty acids from O. tauri wild‐type cells. The putative C21:6 alkene peak is indicated by an arrow (11.13 min). It coelutes with an unknown compound (11.17 min). (b) GC peak of the C21:6 alkene standard obtained through the in vitro decarboxylation of a pure DHA fatty standard using purified CvFAP. (c) Same GC chromatogram portion shown in (a) for the total fatty acid analysis of the cer1/3 mutants (pool of the six mutants). (d) Mass spectrum for the putative C21:6 alkene peak from O. tauri wild‐type cells shown in (a). (e) Mass spectrum of the C21:6 standard peak shown in (b).

OtCER1/3 is necessary for C21:6 alkene synthesis in O. tauri

In order to study the in vivo role of algal CER1/3 enzymes in HC biosynthesis, we developed a genome editing approach in O. tauri. CRISPR‐Cas9 RNPs targeting the OtCER1/3 gene, along with linear DNA encoding the geneticin resistance gene AphI, were co‐delivered into O. tauri cells using the PEG method (Fig. 3a). Geneticin resistant colonies were isolated and then genotyped by PCR (Fig. 3a). Six independent cer1/3 mutants that possess the AphI cassette inserted into the OtCER1/3 gene (Figs 3b, S3), likely through the non‐homologous end joining mechanism, were selected and grown for further functional characterization (Fig. 3c). The C21:6 alkene accumulation was totally abolished in all six cer1/3 mutants (Figs 2c, 3d). The absence of the C21:6 alkene in the cer1/3 mutants did not affect the levels of individual FAMEs, including the level of DHA, which is very likely to be the precursor of C21:6 alkene (Fig. 3e). These results clearly indicated that the OtCER1/3 protein is a functional enzyme in vivo that is responsible for C21:6 alkene biosynthesis in O. tauri cells. However, several possibilities remained as to the exact activity of CER1/3: only aldehyde‐forming activity (i.e. CER3 activity), only alkane‐forming activity (i.e. CER1 activity or yet another activity not using an aldehyde intermediate), or both CER1 and CER3 activities (use of an aldehyde intermediate). Since these activities may have varied during algal evolution, we next decided to study in greater detail the sequence and activity of four CER1/3 proteins present in distantly related green algae: for the Prasinodermophytes, PcCER1/3 from Prasinoderma coloniale CCMP1413; for the Chlorophytes, OtCER1/3 from O. tauri RCC4221; and for the Streptophytes, KnCER1/3 from K. nitens NIES‐2285 (Klebsormidiophyceae) and MkCER1/3 from Mesotaenium kramstae Lemmermann NIES‐657 (Zygnematophyceae). TheCER1/3 homolog from the cryptophyceae lineage (PsCER1/3 from P. sulcata CCMP1175) was also investigated.

Fig. 3.

Fig. 3

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CRISPR‐mediated inactivation of the OtCER1/3 gene in Ostreococcus tauri RCC4221 results in impaired C21:6 alkene biosynthesis. (a) Schematic representation of the CRISPR‐mediated insertion of the geneticin resistance cassette in the OtCER1/3 genomic locus and PCR genotyping for the six O. tauri cer1/3 mutants selected. (b) Schematic representation of the OtCER1/3 locus in the six O. tauri cer1/3 mutants selected. (c) Growth curve of the O. tauri wild‐type (WT) strain and the six cer1/3 mutants. Data are mean ± SE of 4 independent cultures. (d) Heneicosahexaene (C21:6 alkane) content in O. tauri WT strain and the six cer1/3 mutants. Data are represented as individual values and mean ± SE of 4 independent cultures. Asterisks indicate significant differences between the WT strain and each cer1/3 mutant according to a Kruskal‐Wallis test with multiple comparisons with an uncorrected Dunn's test (*, P < 0.01). (e) Fatty acid methyl ester (FAMP) relative abundance in O. tauri WT strain and the six cer1/3 mutants. Data are mean ± SE of 4 independent cultures.

The algal CER1/3 homologs exhibit the key features of both land plant CER1 and CER3

The land plant proteins CER1 and CER3 have a bi‐domain structure consisting of an N‐terminal fatty acid hydroxylase domain (NTD) and a C‐terminal WAX2 domain (CTD). CER1's catalytic activity is believed to be mediated by histidine‐rich motifs in the NTD. By contrast, CER3's activity is thought to be associated with a potentially catalytic cysteine residue and a putative NADPH‐binding site (Bernard et al., 2012; Kojima et al., 2024). Aligning five different algal CER1/3 protein sequences revealed that their NTDs and CTDs display a high degree of similarity to the key motifs of land plant CER1 and CER3, respectively (Fig. 4a). The analysis showed that all five studied algal CER1/3 homologs possess the three highly conserved histidine‐rich motifs in their NTDs, the conserved putative NADPH‐binding site, and the putative catalytic cysteine in their CTDs (Fig. 4a).

Fig. 4.

Fig. 4

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Conservation of putative catalytic motifs and global structure of algal ECERIFERUM 1/3 (CER1/3) proteins compared to their land plant counterparts. (a) Alignment of sequences from five algal CER1/3 proteins and already characterized land plant CER1 and CER3 proteins. Histidine‐rich motifs are shown in red, while the putative NADPH‐binding site and the putative catalytic cysteine are shown in dark blue and pink, respectively. At, Arabidopsis thaliana; Kn, Klebsormidium nitens; Mk, Maesotaenium kramstrae; No, Nymphaea odorata; Ot, Ostreococcus tauri; Pc, Prasinoderma coloniale; Ps, Proteomonas sulcata. (b) Superposition of OtCER1/3 predicted structure with AtCER1 (upper) and AtCER3 (lower). Histidine‐rich motifs are shown in red, while the putative NADPH‐binding site and the putative catalytic cysteine are shown in dark blue and pink, respectively.

We then superposed the predicted 3D structure of the OtCER1/3 protein, as predicted by AlphaFold 3, onto the predicted 3D structures of Arabidopsis CER1 and CER3. This analysis suggested that the OtCER1/3 structure is highly similar to the structures of AtCER1 and AtCER3 (Fig. 4b). The Arabidopsis CER1 and CER3 proteins are embedded in the ER membrane due to the presence of transmembrane domains (Bernard et al., 2012). Sequence analysis of the OtCER1/3 protein showed a predicted localization to the ER and the possible presence of transmembrane domains (DeepLoc 2.1), suggesting that algal CER1/3 may localize to the same subcellular compartment as their land plant counterparts. Taken together, these results suggest that green algal CER1/3 homologs possess structural features that allow them to produce HCs independently.

Algal CER1/3 proteins are functional HC‐forming enzymes

To determine if algal CER1/3 proteins are HC‐forming enzymes or exhibit CER1‐ or CER3‐like activity, the genes encoding five algal CER1/3 proteins were cloned and expressed under a copper‐inducible promoter (pCUP1) in the yeast strain S. cerevisiae INVSc1 (Fig. 5a,b; Tables S1, S2). We chose S. cerevisiae as a model for our heterologous expression studies because it does not naturally produce fatty acid‐derived HCs, yet it has VLC fatty acids and acyl‐CoAs with up to 26 carbons, as well as traces of fatty acids with 28 carbons (Denic & Weissman, 2007; Bernard et al., 2012; Wang et al., 2016). To validate our experimental approach, we first expressed the known Arabidopsis CER1 and/or CER3 proteins (Bernard et al., 2012). Although expressing AtCER1 and AtCER3 individually did not result in HC production, coexpressing them led to the accumulation of C17:0 and C25:0 alkanes (Fig. 5c). Substitution of amino acids in the NTD of AtCER1 (second histidine‐rich motif) or the CTD of AtCER3 (putative NADPH‐binding site or catalytic cysteine) impaired HC production (Fig. 5c), confirming the importance of these domains for catalytic activity.

Fig. 5.

Fig. 5

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Expression of land plant ECERIFERUM 1/3 (CER1/3) and algal CER1/3 proteins in yeast. (a) Scheme showing the genetic construct strategy used for the heterologous expression of land plant CER1/CER3 and algal CER1/3 proteins in yeast. (b) Example of copper‐inducible expression in yeast (addition of 10 μM CuSO4 was followed by 24 h of growth on a solid plate) using a construct expressing the mCherry2 gene under the control of the yeast copper‐inducible promoter (CUP1). (c) Hydrocarbon content in yeast cells expressing different land plant CER1 and/or CER3 proteins. Data are mean + SE of independent cultures: n = 4 (AtCER1 + AtCER3‐C587A, AtCER1 + AtCER3‐A461N‐T462L), n = 6 (AtCER1, AtCER3, AtCER1‐H159A + AtCER3), n = 22 (EV, AtCER1 + AtCER3). (d) Hydrocarbon content in yeast cells expressing different algal CER1/3 proteins. Data are mean + SE of independent cultures: n = 4 (PsCER1/3, MkCER1/3), n = 12 (PcCER1/3 and KnCER1/3), n = 34 (OtCER1/3), n = 44 (EV). (e) Hydrocarbon content in yeast cells expressing different versions of the OtCER1/3 protein. Data are mean + SE of independent cultures: n = 6 (each mutated version of OtCER1/3), n = 12 (EV, OtCER1/3). EV, Empty vector; At, A. thaliana; Ps, Proteomonas sulcata; Pc, Prasinoderma coloniale; Ot, Ostreococcus tauri; Kn, Klebsormidium nitens; Mk, Mesotaenium kramstae.

Using the same experimental approach, we next expressed algal CER1/3 proteins. Interestingly, expressing each protein alone was sufficient to enable the production of HCs (Fig. 5d). The cryptophyte PsCER1/3 protein and the prasinodermophyte PcCER1/3 protein both produced 8‐heptadecene and n‐heptadecane, while the chlorophyte OtCER1/3 protein produced three different LC HCs, which were identified as 7‐pentadecene, 8‐heptadecene, and n‐heptadecane (Figs 5d, S4). Expression of the streptophyte KnCER1/3 and MkCER1/3 proteins only produced n‐heptadecane or 8‐heptadecene, respectively (Figs 5d, S4). Notably, no VLC HCs (i.e. > C20) were detected in any of the strains expressing the different algal CER1/3 homologs.

To determine if the key motifs of land plant CER1 and CER3 proteins are important for catalytic activity in algal CER1/3 proteins, we produced versions of OtCER1/3 with specific amino acid changes. Substituting any of the histidine‐rich motifs (H142A, H155A, or H243A) with a single amino acid abolishes HC production, as do substitutions in the putative NADPH‐binding (A455N and T456L) or catalytic (C581A) sites (Fig. 5e).

Taken together, these results demonstrate that algal CER1/3 proteins are functional HC‐forming enzymes with the conserved catalytic motifs of both land plant proteins CER1 and CER3, suggesting a potential bifunctional activity.

Single algal CER1/3s are bifunctional enzymes that perform the same catalytic reactions as the plant CER1/CER3 complex

Since algal CER1/3 proteins can produce HCs when expressed alone in yeast, it is possible that these proteins are bifunctional, possessing both CER3‐like activity (converting acyl‐CoAs into aldehydes) and CER1‐like activity (reducing aldehydes to alkanes). Alternatively, algal CER1/3 proteins may operate via an intermediate distinct from aldehydes or use a different mechanism to convert acyl‐CoAs directly into alkanes.

We first evaluated whether OtCER1/3 can form both homodimers and heterodimers with its Arabidopsis homologs using a SUY2H approach. In contrast to yeast cells that coexpress Cub‐OtCER1/3 and NubG (the negative control), yeast cells coexpressing Cub‐OtCER1/3 and NubG‐OtCER1/3 were able to grow on a selective media lacking histidine and adenine, as observed for the positive control (NubWT) (Fig. 6). Furthermore, yeast cells coexpressing Cub‐OtCER1/3 and NubG‐AtCER1 or ‐AtCER3 were able to grow on the same selective medium (Fig. 6). These results strongly suggest that OtCER1/3 enzymes homodimerize but also heterodimerize with AtCER1 and AtCER3.

Fig. 6.

Fig. 6

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OtCER1/3 interacts with itself and with Arabidopsis CER1 and CER3. Yeast cells cotransformed with Cub‐OtCER1/3 as bait and NubWT, NubG, NubG‐AtCER1, and NubG‐AtCER3 as prey were grown on selective media lacking leucine and tryptophan (−LT) or lacking leucine, tryptophan, histidine, and adenine (‐LTHA). OD600nm dilution of yeast cells, from left to right: 1, 10−1, 10−2, 10−3. These results were independently replicated twice with similar outcomes. CER1, ECERIFERUM 1; CER3, ECERIFERUM 3.

Then, to evaluate whether aldehydes are possible substrates of different algal CER1/3 protein homologs, we coexpressed OtCER1/3 with AtCER3 (or AtCER1 as a control) in yeast. AtCER3 has been suggested to produce C24–C32 aldehydes, while AtCER1 is thought to form C27–C31 alkanes from C28–C32 aldehydes (Bernard et al., 2012; Pascal et al., 2019; Kojima et al., 2024). Interestingly, the HC profile was significantly impacted by the co‐expression of algal CER1/3 and AtCER3 compared to algal CER1/3 alone (Fig. 7a). Co‐expression of all five algal CER1/3 with AtCER3 resulted in the production of VLC alkanes (e.g. C23 and/or C25 alkanes). Furthermore, coexpressing AtCER3 with PsCER1/3, OtCER1/3, or MkCER1/3 increased LC HC accumulation. By contrast, co‐expression of OtCER1/3 with AtCER1 did not yield VLC alkanes and even reduced LC alkanes, producing only 8‐heptadecene and small amounts of n‐heptadecane.

Fig. 7.

Fig. 7

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Co‐expression of algal ECERIFERUM 1/3 (CER1/3) with Arabidopsis CER1/CER3 proteins in yeast. (a) Hydrocarbon content in yeast cells expressing different algal CER1/3 and land plant CER3 or CER1 proteins. Data are mean + SE of independent cultures: n = 4 (PsCER1/3, PsCER1/3 + AtCER3, MkCER1/3, MkCER1/3 + AtCER3), n = 6 (OtCER1/3 + AtCER1, PcCER1/3, PcCER1/3 + AtCER3, KnCER1/3, KnCER1/3 + AtCER3), n = 16 (OtCER1/3, OtCER1/3 + AtCER3), n = 20 (EV). (b) Hydrocarbon content in yeast cells expressing OtCER1/3 and different versions of AtCER3. Data are mean + SE of independent cultures: n = 4 (OtCER1/3 + AtCER3‐C587A, OtCER1/3 + AtCER3‐A461N‐T462L), n = 6 (OtCER1/3 + AtCER3, OtCER1/3 + AtCER3‐H147A, OtCER1/3 + AtCER3‐H161A, OtCER1/3 + AtCER3‐H250A), n = 10 (EV). (c) Hydrocarbon content in yeast cells expressing different versions of OtCER1/3 and AtCER3. Data are mean + SE of 4 independent cultures. EV, Empty vector; At, Arabidopsis thaliana; Ps, Proteomonas sulcata; Pc, Prasinoderma coloniale; Ot, Ostreococcus tauri; Kn, Klebsormidium nitens; Mk, Mesotaenium kramstae.

To determine which domain(s) of AtCER3 are responsible for C25:0 alkane formation when coexpressed with OtCER1/3, AtCER3 variants mutated in the histidine‐rich motifs (i.e. H147A, H161A or H250A), the putative catalytic cysteine (i.e. C587A), or the putative NADPH‐binding site (i.e. A461N and T462L) were coexpressed with native OtCER1/3. The production of the C25:0 alkane was unaffected by mutations in the different histidine‐rich motifs of AtCER3 (Fig. 7b). On the other hand, mutations in the putative NADPH‐binding site or the putative catalytic cysteine of AtCER3 resulted in the abolition of the C25:0 alkane production and a considerable decrease in the alkane content compared to the control construct. These results clearly demonstrate that OtCER1/3 uses the LC and VLC aldehyde intermediates produced by the reductase activity of the CTD of AtCER3, indicating that the decarbonylase activity of algal CER1/3 exhibits broad substrate specificity.

Finally, we coexpressed the native AtCER3 with different mutated versions of the OtCER1/3 protein that are impaired for HC production (Fig. 5e). OtCER1/3 proteins mutated in the histidine‐rich motifs could not be rescued by the co‐expression with AtCER3 (Fig. 7c). However, co‐expression of AtCER3 with the OtCER1/3 protein mutated in the putative catalytic cysteine led to HC production similar to the control construct. This result suggests that AtCER3 has a broad substrate specificity, ranging from C16 to C26 acyl‐CoA. The co‐expression of AtCER3 with the OtCER1/3 protein mutated in the NADPH‐binding site resulted in the accumulation of low levels of LC and VLC HCs. This result may be explained by partially impaired protein–protein interaction, as mutations in the putative NADPH‐binding site reduce the ability to dimerize with the native OtCER1/3 protein, contrary to the effect of mutations in the histidine‐rich motifs or the putative catalytic cysteine (Fig. S5).

Taken together, these data demonstrate that, like AtCER1, the algal CER1/3 proteins have the ability to use the VLC aldehyde intermediates produced by AtCER3 to synthesize VLC alkanes. It is therefore likely that the algal CER1/3 proteins are bifunctional enzymes that perform the same catalytic reactions as the plant CER1/CER3 complex.

Discussion

Algal CER1/3 proteins are functional HC‐forming enzymes

Here, we establish that the presence of FAP or CER1/3 genes in the 99 green algal genomes investigated is mutually exclusive (Table 1). CER1/3 homologs were also identified in some cryptophyte algae. Additionally, algal species lacking the FAP enzyme but possessing a CER1/3 homolog, such as O. tauri and K. nitens, were able to produce HCs (Fig. 2), suggesting that these algal CER1/3 proteins function as HC‐producing enzymes. After performing multiple sequence alignments and analyzing the AlphaFold‐predicted 3D structures of various algal CER1/3 proteins, as well as Arabidopsis CER1 and CER3 proteins (Fig. 4), we observed that algal CER1/3 proteins possess in their NTD the key motifs of land plant CER1 (histidine‐rich motifs), and in their CTD, the key motifs of land plant CER3 (a putative NADPH‐binding site and a putative catalytic cysteine). This suggests not only that algal CER1/3 proteins may be HC‐producing enzymes, but also that the enzyme may be bifunctional, performing both CER1 and CER3 activities.

Using heterologous expression in yeast, we further demonstrated that green algal CER1/3 s are functional LC HC‐forming enzymes (Fig. 5). These enzymes exhibit different substrate specificities across the large panel of species investigated, which may reflect different in vivo functions. Contrary to Arabidopsis CER1 and CER3, none of the algal CER1/3 enzymes used in this study produce VLC HCs (i.e. C25 HCs) when expressed in yeast (Fig. 5). This observation likely reflects the different substrate specificities of algal CER1/3 and land plant CER1 and CER3 enzymes. The key amino acid residues controlling substrate specificity are yet to be identified.

Algal CER1/3 proteins are bifunctional enzymes

As demonstrated in this study and in previous work (Bernard et al., 2012; Kojima et al., 2024), the key motifs of the CTD of land plant CER3 (the putative NADPH‐binding site and the putative catalytic cysteine) and the key motifs of the NTD of land plant CER1 (the histidine‐rich motifs) are essential for HC production, as deletion of these motifs impairs HC synthesis. Individual mutations in the corresponding motifs in the green algal OtCER1/3 resulted in impaired HC production, suggesting that algal CER1/3 may operate via a similar mechanism as land plant CER1/CER3 complexes. Furthermore, OtCER1/3 enzymes expressed in yeast can form homodimers, but also heterodimers with their two distantly related Arabidopsis homologs (Fig. 6). This result suggests that OtCER1/3 enzymes may form a hydrocarbon‐forming complex in algal cells. It remains to be deciphered whether other partners are involved in this complex, as they are in land plants (Bernard et al., 2012; Huang et al., 2024).

Co‐expression of various algal CER1/3 proteins with A. thaliana CER3, an aldehyde‐producing enzyme, altered the HC product profile (Fig. 7). In addition to LC HCs, VLC HCs (mainly C25:0 alkane) were produced in all tested algal CER1/3 s, suggesting that AtCER3 provides VLC aldehydes that can be further processed into HCs by the NTD of algal CER1/3s. When OtCER1/3 was coexpressed together with AtCER3 mutated in the histidine‐rich motifs, the strain was still capable of producing LC and VLC HCs. However, when OtCER1/3 was coexpressed together with different AtCER3 versions individually mutated in the putative NADPH‐binding site or the putative catalytic cysteine, only LC HCs were produced. In addition, coexpressing the native AtCER3 together with the OtCER1/3 version mutated in the putative catalytic cysteine restored the HC biosynthesis profile (Fig. 7). Our results strongly suggest that LC and VLC aldehydes are provided by the CTD of AtCER3 to the NTD of OtCER1/3 for further decarbonylation into LC and VLC HCs. Furthermore, these results, along with the fact that there is only one single CER1/3 homolog in the algal genomes, support the hypothesis that algal CER1/3 proteins are bifunctional enzymes with both CER1‐ and CER3‐like activities (Fig. 8). We hypothesize that the CTD of algal CER1/3 proteins converts a LC acyl‐CoA into a substrate intermediate, very likely an aldehyde, which is then decarbonylated into a LC HC by their NTD. As proposed for the AAR enzyme, it is possible that the acyl‐CoA first forms a thioester bond with the putative catalytic cysteine of CER1/3 CTD, and then NADPH provides a hydride to break this bond, releasing the acyl chain as an aldehyde (Gao et al., 2020). This intermediate molecule would then be decarbonylated into HC in the active site of CER1/3 NTD, potentially through the involvement of histidine residues in the coordination of metal atoms and the formation of a dimetal center, similar to what has been described in the yeast sphingolipid α‐hydroxylase (Zhu et al., 2015). Further experiments are now needed to describe this biosynthetic system in more detail.

Fig. 8.

Fig. 8

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Schematic representation of the proposed reactions carried out by a ECERIFERUM 1/3 (CER1/3) protein from green algae. CoA, coenzyme A; CTD, C‐terminal domain; Cys, catalytic cysteine of C‐terminal domain; ER, endoplasmic reticulum; His, catalytic histidines of N‐terminal domain; NTD, N‐terminal domain. Only one monomer is represented, but the CER1/3 protein very likely exists as a homodimer.

Land plant CER1 and CER3 genes evolved from an ancestral algal CER1/3 gene

In algal CER1/3 s, both the CTD and the NTD motifs are essential for HC production, whereas in land plants, only the NTD or CTD motifs are essential for CER1 or CER3 activity, respectively. These findings, together with the phylogenetic analysis (Fig. 1), support the view that the land plant CER1 and CER3 genes originated from the duplication and differentiation of an ancestral algal CER1/3 gene (Wang et al., 2019; Chaudhary et al., 2021), which existed before the divergence of the three green algal phyla (prasinodermophyte, chlorophyte, and streptophyte algae). The presence of functional CER1/3 proteins in the current streptophyte algae K. nitens and M. kramstae, which are close relatives of the land plant common ancestor (Delwiche & Cooper, 2015; De Vries & Archibald, 2018), further reinforces this evolutionary scenario. As proposed in another work (Kojima et al., 2024), it seems that after gene duplication, land plant CER1 and CER3 proteins carried out different processes of differentiation. The NTD of CER1 remained enzymatically active for aldehyde decarbonylation, whereas the CTD may have lost its catalytic activity for aldehyde formation. Conversely, the NTD of CER3 may have lost its decarbonylation activity while its CTD remained enzymatically active for the reduction of acyl‐CoA into aldehydes. Such a protein specialization may have provided early land plants with new regulatory mechanisms. Interestingly, a recent study showed that Arabidopsis CER1 and CER3 form a dynamic complex with the aldehyde reductase SOH1, which also localizes to the ER (Li et al., 2025). In this regulatory complex, CER3 provides aldehydes that are then channeled into either the alcohol‐ (via SOH1) or the alkane‐ (via CER1) forming pathways in response to environmental conditions, independently of the CER4 fatty acyl‐CoA reductase pathway, which forms primary fatty alcohols in the waxes of many plants (Keyl et al., 2024). While the alcohol accumulation is associated with high cuticular transpiration, high alkane content is associated with low cuticular transpiration and therefore higher resistance to drought conditions (Li et al., 2025). Therefore, the duplication of the green algal CER1/3 gene followed by specialization of each gene into CER1 and CER3 likely constitutes a key evolutionary event that helped land plants to colonize terrestrial environments more easily by providing a fine‐tuning of the alkane/fatty alcohol ratio and the possibility to produce and release fatty aldehydes more efficiently than with CER1/3s. The specific role of fatty aldehydes on plant surfaces is still to be elucidated.

The role of algal CER1/3‐derived HCs still needs to be deciphered

Although the role of fatty acid‐derived HCs has been well documented in land plants, their biological and physiological functions in algae remain elusive. In the green microalga C. reinhardtii, it has been shown that the FAP enzyme is a soluble protein bound to the thylakoid membrane that is responsible for the synthesis of only one type of HC (7‐heptadecene) found mostly in the thylakoid fraction (Moulin et al., 2021). While inactivation of the C. reinhardtii FAP gene leads to transient photosynthetic impairment under cold conditions when light intensity varies (Moulin et al., 2021), the exact role of FAP‐derived HCs remains elusive. In this study, we developed a CRISPR‐Cas9‐based strategy for producing cer1/3 mutants in O. tauri, which are the first mutants of an algal CER1/3 gene. This strategy enabled us to demonstrate that the OtCER1/3 enzyme is responsible for the biosynthesis of heneicosahexaene (C21:6 alkene), the only HC found in O. tauri cells. The precise intracellular localization of algal CER1/3 enzymes, which are potentially embedded in the ER membrane like their land plant counterparts (Bernard et al., 2012; Pascal et al., 2019), and their HC product are important questions yet to be answered in order to identify the biological role of OtCER1/3‐derived HCs. Further work is also needed to elucidate the evolutionary role of algal CER1/3 enzymes. For example, it would be interesting to assess the in vivo function of other algal CER1/3 enzymes, in particular in K. nitens, which only produces a C17:0 alkane. Whether the different HCs produced by phylogenetically distant algal CER1/3 enzymes are reflected by different cellular and/or physiological functions constitutes an interesting open question.

Competing interests

None declared.

Author contributions

AB‐P, BL, AS, MC, MLC, AG, FV and FB carried out the experiments. AB‐P, DS, FV and FB contributed to the design and implementation of the research. AB‐P, BL, FV and FB contributed to the analysis of the results. AB‐P wrote the original draft. FV and FB wrote the final manuscript, with contributions from DS and YL‐B to the editing and revision of the final manuscript. All authors approved the final manuscript.

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Supporting information

Fig. S1 Mass spectrum of the C17 alkane detected in Klebsormidium nitens unsaponifiable cell material.

Fig. S2 Portion of the GC chromatogram showing the putative C21:6 alkene peak from Ostreococcus tauri wild‐type cells.

Fig. S3 Sanger chromatograms of the Ostreococcus tauri wild‐type (WT) strain and the six cer1/3 mutants.

Fig. S4 Mass spectrum of hydrocarbons produced in yeast experiments.

Fig. S5 Dimerization of mutated OtCER1/3 proteins with the native OtCER1/3 protein.

NPH-251-1311-s002.pdf (1.1MB, pdf)

Table S1 List of protein sequences used in this study for phylogenetic analysis, sequence alignments and heterologous expression in yeast.

Table S2 List of plasmids used in this study for yeast (INVSc1 strain) heterologous expression.

Table S3 Assembly of CRISPR‐Cas9 RNPs, in vitro cleavage assay and PEG‐mediated transformation of Ostreococcus tauri cells.

Table S4 Linear DNA constructs used for the transformation of Ostreococcus tauri cells.

Table S5 List of primers used in this study for Ostreococcus tauri genome editing experiments.

Table S6 List of genomes from different species or strains of green algae investigated for the presence of FAP and CER1/CER3 homologs.

Please note: Wiley is not responsible for the content or functionality of any Supporting Information supplied by the authors. Any queries (other than missing material) should be directed to the New Phytologist Central Office.

NPH-251-1311-s001.xlsx (1.1MB, xlsx)

Acknowledgements

We thank Louise Ardouin for technical help and Dr Thierry Desnos and Dr Zhongze Li for helpful discussions. We also thank Dr Xavier Bailly for providing the T. convolutae CER1/3 sequence, the Merchant lab for providing the Auxenochlorella UTEX 250‐A strain, and Dr Shiyou Lü for kindly providing the NMY51 yeast strain and the pBT3N and pPR3N vectors. This work was funded in part by the CEA internal program ‘Economie Circulaire du Carbone’, the Exploratory Program of CEA and the Horizon Prize, and in part by the Agence Nationale de la Recherche project Snapshot (ANR‐18‐CE11‐0021) and Brown Alkaenes (ANR‐24‐CE20‐3101‐01). The funding provided by the European Union Regional Development Fund (ERDF), the Région Sud, the French Ministry of Research, and the CEA to the HelioBiotec platform is also acknowledged. ABP was supported by a CEA‐funded PhD fellowship. Open access publication funding provided by COUPERIN CY26.

Contributor Information

Florian Veillet, Email: florian.veillet@cea.fr.

Fred Beisson, Email: frederic.beisson@cea.fr.

Data availability

The data that support the findings of this study are included as part of the manuscript or as Supporting Information. Accession numbers of algal and land plant proteins used in this study can be found in Table S1.

References

  1. Aarts MG, Keijzer CJ, Stiekema WJ, Pereira A. 1995. Molecular characterization of the CER1 gene of Arabidopsis involved in epicuticular wax biosynthesis and pollen fertility. Plant Cell 7: 2115–2127. [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Abramson J, Adler J, Dunger J, Evans R, Green T, Pritzel A, Ronneberger O, Willmore L, Ballard AJ, Bambrick J et al. 2024. Accurate structure prediction of biomolecular interactions with AlphaFold 3. Nature 630: 493–500. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Beisson F, Li‐Beisson Y, Pollard M. 2012. Solving the puzzles of cutin and suberin polymer biosynthesis. Current Opinion in Plant Biology 15: 329–337. [DOI] [PubMed] [Google Scholar]
  4. Bernard A, Domergue F, Pascal S, Jetter R, Renne C, Faure J‐D, Haslam RP, Napier JA, Lessire R, Joubès J. 2012. Reconstitution of plant alkane biosynthesis in yeast demonstrates that Arabidopsis ECERIFERUM1 and ECERIFERUM3 are core components of a very‐long‐chain alkane synthesis complex. Plant Cell 24: 3106–3118. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Bernard A, Joubès J. 2013. Arabidopsis cuticular waxes: advances in synthesis, export and regulation. Progress in Lipid Research 52: 110–129. [DOI] [PubMed] [Google Scholar]
  6. Bowman JL. 2022. The origin of a land flora. Nature Plants 8: 1352–1369. [DOI] [PubMed] [Google Scholar]
  7. Camacho DJ, Moseley JL, Merchant SS. 2025. A defined growth medium for trace metal limitation studies in Auxenochlorella sp. (UTEX 250‐A) during mixotrophic growth with glucose. bioRxiv doi: 10.1101/2025.05.09.653207. [DOI]
  8. Chaudhary K, Geeta R, Panjabi P. 2021. Origin and diversification of ECERIFERUM1 (CER1) and ECERIFERUM3 (CER3) genes in land plants and phylogenetic evidence that the ancestral CER1/3 gene resulted from the fusion of pre‐existing domains. Molecular Phylogenetics and Evolution 159: 107101. [DOI] [PubMed] [Google Scholar]
  9. Concordet J‐P, Haeussler M. 2018. CRISPOR: intuitive guide selection for CRISPR/Cas9 genome editing experiments and screens. Nucleic Acids Research 46: W242–W245. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Craig RJ, Dueñas MA, Camacho DJ, Gallaher SD, Avendaño‐Monsalve MC, Lin Y‐T, Blaby‐Haas CE, Moseley JL, Merchant SS. 2025. Targeted genetic manipulation and yeast‐like evolutionary genomics in the green alga Auxenochlorella. Plant Cell 37: koaf259. [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. De Vries J, Archibald JM. 2018. Plant evolution: landmarks on the path to terrestrial life. New Phytologist 217: 1428–1434. [DOI] [PubMed] [Google Scholar]
  12. Delaux P‐M, Schornack S. 2021. Plant evolution driven by interactions with symbiotic and pathogenic microbes. Science 371: eaba6605. [DOI] [PubMed] [Google Scholar]
  13. Delude C, Moussu S, Joubès J, Ingram G, Domergue F. 2016. Plant surface lipids and epidermis development. In: Nakamura Y, Li‐Beisson Y, eds. Lipids in plant and algae development. Cham, Switzerland: Springer International Publishing, 287–313. [DOI] [PubMed] [Google Scholar]
  14. Delwiche CF, Cooper ED. 2015. The evolutionary origin of a terrestrial flora. Current Biology 25: R899–R910. [DOI] [PubMed] [Google Scholar]
  15. Denic V, Weissman JS. 2007. A molecular caliper mechanism for determining very long‐chain fatty acid length. Cell 130: 663–677. [DOI] [PubMed] [Google Scholar]
  16. Gao Y, Zhang H, Fan M, Jia C, Shi L, Pan X, Cao P, Zhao X, Chang W, Li M. 2020. Structural insights into catalytic mechanism and product delivery of cyanobacterial acyl‐acyl carrier protein reductase. Nature Communications 11: 1525. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Haslam TM, Kunst L. 2013. Extending the story of very‐long‐chain fatty acid elongation. Plant Science 210: 93–107. [DOI] [PubMed] [Google Scholar]
  18. Hoang DT, Chernomor O, Von Haeseler A, Minh BQ, Vinh LS. 2018. UFBoot2: improving the ultrafast bootstrap approximation. Molecular Biology and Evolution 35: 518–522. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Huang H, Wang Y, Yang P, Zhao H, Jenks MA, Lü S, Yang X. 2024. The Arabidopsis cytochrome P450 enzyme CYP96A4 is involved in the wound‐induced biosynthesis of cuticular wax and cutin monomers. The Plant Journal 118: 1619–1634. [DOI] [PubMed] [Google Scholar]
  20. Jenks MA, Tuttle HA, Eigenbrode SD, Feldmann KA. 1995. Leaf epicuticular waxes of the eceriferum mutants in Arabidopsis. Plant Physiology 108: 369–377. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Jetter R, Kunst L, Samuels AL. 2006. Composition of plant cuticular waxes. In: Riederer M, Müller C, eds. Annual Plant Reviews Volume 23: Biology of the Plant Cuticle. London, UK: John Wiley & Sons. doi: 10.1002/9780470988718.ch4. [DOI] [Google Scholar]
  22. Kalyaanamoorthy S, Minh BQ, Wong TKF, Von Haeseler A, Jermiin LS. 2017. ModelFinder: fast model selection for accurate phylogenetic estimates. Nature Methods 14: 587–589. [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Keyl A, Herrfurth C, Pandey G, Kim RJ, Helwig L, Haslam TM, De Vries S, De Vries J, Gutsche N, Zachgo S et al. 2024. Divergent evolution of the alcohol‐forming pathway of wax biosynthesis among bryophytes. New Phytologist 242: 2251–2269. [DOI] [PubMed] [Google Scholar]
  24. Kim M, Jorge GL, Aschern M, Cuiné S, Bertrand M, Mekhalfi M, Putaux J‐L, Yang J‐S, Thelen JJ, Beisson F et al. 2024. The DYRKP1 kinase regulates cell wall degradation in Chlamydomonas by inducing matrix metalloproteinase expression. Plant Cell 36: 4988–5003. [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. Kojima H, Yamamoto K, Suzuki T, Hayakawa Y, Niwa T, Tokuhiro K, Katahira S, Higashiyama T, Ishiguro S. 2024. Broad chain‐length specificity of the alkane‐forming enzymes NoCER1A and NoCER3A/B in Nymphaea odorata . Plant and Cell Physiology 65: 428–446. [DOI] [PMC free article] [PubMed] [Google Scholar]
  26. Kondo S, Hori K, Sasaki‐Sekimoto Y, Kobayashi A, Kato T, Yuno‐Ohta N, Nobusawa T, Ohtaka K, Shimojima M, Ohta H. 2016. Primitive extracellular lipid components on the surface of the charophytic alga Klebsormidium flaccidum and their possible biosynthetic pathways as deduced from the genome sequence. Frontiers in Plant Science 7: 952. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Kosma DK, Graça J, Molina I. 2025. Update on the structure and regulated biosynthesis of the apoplastic polymers cutin and suberin. Plant Physiology 197: kiae653. [DOI] [PubMed] [Google Scholar]
  28. Labun K, Montague TG, Krause M, Torres Cleuren YN, Tjeldnes H, Valen E. 2019. CHOPCHOP v3: expanding the CRISPR web toolbox beyond genome editing. Nucleic Acids Research 47: W171–W174. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Lee ME, DeLoache WC, Cervantes B, Dueber JE. 2015. A highly characterized yeast toolkit for modular, multipart assembly. ACS Synthetic Biology 4: 975–986. [DOI] [PubMed] [Google Scholar]
  30. Lee RF, Loeblich AR. 1971. Distribution of 21:6 hydrocarbon and its relationship to 22:6 fatty acid in algae. Phytochemistry 10: 593–602. [Google Scholar]
  31. Lee SB, Suh MC. 2015. Advances in the understanding of cuticular waxes in Arabidopsis thaliana and crop species. Plant Cell Reports 34: 557–572. [DOI] [PubMed] [Google Scholar]
  32. Li L, Wang S, Wang H, Sahu SK, Marin B, Li H, Xu Y, Liang H, Li Z, Cheng S et al. 2020. The genome of Prasinoderma coloniale unveils the existence of a third phylum within green plants. Nature Ecology & Evolution 4: 1220–1231. [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Li S, Zhang X, Huang H, Yin M, Jenks MA, Kosma DK, Yang P, Yang X, Zhao H, Lü S. 2025. Deciphering the core shunt mechanism in Arabidopsis cuticular wax biosynthesis and its role in plant environmental adaptation. Nature Plants 11: 165–175. [DOI] [PubMed] [Google Scholar]
  34. Li‐Beisson Y, Shorrosh B, Beisson F, Andersson MX, Arondel V, Bates PD, Baud S, Bird D, DeBono A, Durrett TP et al. 2013. Acyl‐lipid metabolism. The Arabidopsis Book 2010: e0133. [DOI] [PMC free article] [PubMed] [Google Scholar]
  35. Metzger P, Largeau C. 2005. Botryococcus braunii: a rich source for hydrocarbons and related ether lipids. Applied Microbiology and Biotechnology 66: 486–496. [DOI] [PubMed] [Google Scholar]
  36. Moulin SLY, Beyly‐Adriano A, Cuiné S, Blangy S, Légeret B, Floriani M, Burlacot A, Sorigué D, Samire P‐P, Li‐Beisson Y et al. 2021. Fatty acid photodecarboxylase is an ancient photoenzyme that forms hydrocarbons in the thylakoids of algae. Plant Physiology 186: 1455–1472. [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Nguyen L‐T, Schmidt HA, Von Haeseler A, Minh BQ. 2015. IQ‐Tree: a fast and effective stochastic algorithm for estimating maximum‐likelihood phylogenies. Molecular Biology and Evolution 32: 268–274. [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. Nielsen H. 2025. Practical applications of language models in protein sorting prediction: SignalP 6.0, DeepLoc 2.1, and DeepLocPro 1.0. In: Dukka KC, ed. Large language models (LLMs) in protein bioinformatics. New York, NY, USA: Springer US, 153–175. [DOI] [PubMed] [Google Scholar]
  39. Pascal S, Bernard A, Deslous P, Gronnier J, Fournier‐Goss A, Domergue F, Rowland O, Joubès J. 2019. Arabidopsis CER1‐LIKE1 functions in a cuticular very‐long‐chain alkane‐forming complex. Plant Physiology 179: 415–432. [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Pollard M, Beisson F, Li Y, Ohlrogge JB. 2008. Building lipid barriers: biosynthesis of cutin and suberin. Trends in Plant Science 13: 236–246. [DOI] [PubMed] [Google Scholar]
  41. Rowland O, Lee R, Franke R, Schreiber L, Kunst L. 2007. The CER3 wax biosynthetic gene from Arabidopsis thaliana is allelic to WAX2/YRE/FLP1. FEBS Letters 581: 3538–3544. [DOI] [PubMed] [Google Scholar]
  42. Samuels L, Kunst L, Jetter R. 2008. Sealing plant surfaces: cuticular wax formation by epidermal cells. Annual Review of Plant Biology 59: 683–707. [DOI] [PubMed] [Google Scholar]
  43. Sanchez F, Geffroy S, Norest M, Yau S, Moreau H, Grimsley N. 2019. Simplified transformation of Ostreococcus tauri using polyethylene glycol. Genes 10: 399. [DOI] [PMC free article] [PubMed] [Google Scholar]
  44. Sorigué D, Légeret B, Cuiné S, Blangy S, Moulin S, Billon E, Richaud P, Brugière S, Couté Y, Nurizzo D et al. 2017. An algal photoenzyme converts fatty acids to hydrocarbons. Science 357: 903–907. [DOI] [PubMed] [Google Scholar]
  45. Sorigué D, Légeret B, Cuiné S, Morales P, Mirabella B, Guédeney G, Li‐Beisson Y, Jetter R, Peltier G, Beisson F. 2016. Microalgae synthesize hydrocarbons from long‐chain fatty acids via a light‐dependent pathway. Plant Physiology 171: 2393–2405. [DOI] [PMC free article] [PubMed] [Google Scholar]
  46. Wang H, Ni X, Harris‐Shultz K. 2019. Molecular evolution of the plant ECERIFERUM1 and ECERIFERUM3 genes involved in aliphatic hydrocarbon production. Computational Biology and Chemistry 80: 1–9. [DOI] [PubMed] [Google Scholar]
  47. Wang M, Wang Y, Wu H, Xu J, Li T, Hegebarth D, Jetter R, Chen L, Wang Z. 2016. Three TaFAR genes function in the biosynthesis of primary alcohols and the response to abiotic stresses in Triticum aestivum . Scientific Reports 6: 25008. [DOI] [PMC free article] [PubMed] [Google Scholar]
  48. Yeats TH, Rose JKC. 2013. The formation and function of plant cuticles. Plant Physiology 163: 5–20. [DOI] [PMC free article] [PubMed] [Google Scholar]
  49. Zhu G, Koszelak‐Rosenblum M, Connelly SM, Dumont ME, Malkowski MG. 2015. The crystal structure of an integral membrane fatty acid α‐hydroxylase. Journal of Biological Chemistry 290: 29820–29833. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Fig. S1 Mass spectrum of the C17 alkane detected in Klebsormidium nitens unsaponifiable cell material.

Fig. S2 Portion of the GC chromatogram showing the putative C21:6 alkene peak from Ostreococcus tauri wild‐type cells.

Fig. S3 Sanger chromatograms of the Ostreococcus tauri wild‐type (WT) strain and the six cer1/3 mutants.

Fig. S4 Mass spectrum of hydrocarbons produced in yeast experiments.

Fig. S5 Dimerization of mutated OtCER1/3 proteins with the native OtCER1/3 protein.

NPH-251-1311-s002.pdf (1.1MB, pdf)

Table S1 List of protein sequences used in this study for phylogenetic analysis, sequence alignments and heterologous expression in yeast.

Table S2 List of plasmids used in this study for yeast (INVSc1 strain) heterologous expression.

Table S3 Assembly of CRISPR‐Cas9 RNPs, in vitro cleavage assay and PEG‐mediated transformation of Ostreococcus tauri cells.

Table S4 Linear DNA constructs used for the transformation of Ostreococcus tauri cells.

Table S5 List of primers used in this study for Ostreococcus tauri genome editing experiments.

Table S6 List of genomes from different species or strains of green algae investigated for the presence of FAP and CER1/CER3 homologs.

Please note: Wiley is not responsible for the content or functionality of any Supporting Information supplied by the authors. Any queries (other than missing material) should be directed to the New Phytologist Central Office.

NPH-251-1311-s001.xlsx (1.1MB, xlsx)

Data Availability Statement

The data that support the findings of this study are included as part of the manuscript or as Supporting Information. Accession numbers of algal and land plant proteins used in this study can be found in Table S1.

Articles from The New Phytologist are provided here courtesy of Wiley

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