PDAT regulates PE as transient carbon sink alternative to triacylglycerol in Nannochloropsis

Abstract

Triacylglycerols (TAGs) are the main storage lipids in photosynthetic organisms under stress. In the oleaginous alga Nannochloropsis oceanica, while multiple acyl CoA:diacylglycerol (DAG) acyltransferases (NoDGATs) are involved in TAG production, the role of the unique phospholipid:DAG acyltransferase (NoPDAT) remains unknown. Here, we performed a functional complementation assay in TAG-deficient yeast (Saccharomyces cerevisiae) and an in vitro assay to probe the acyltransferase activity of NoPDAT. Subcellular localization, overexpression, and knockdown (KD) experiments were also conducted to elucidate the role of NoPDAT in N. oceanica. NoPDAT, residing at the outermost plastid membrane, does not phylogenetically fall into the clades of algae or plants and uses phosphatidylethanolamine (PE) and phosphatidylglycerol with 16:0, 16:1, and 18:1 at position sn-2 as acyl-donors in vivo. NoPDAT KD, not triggering any compensatory mechanism via DGATs, led to an ∼30% decrease of TAG content, accompanied by a vast accumulation of PEs rich in 16:0, 16:1, and 18:1 fatty acids (referred to as “LU-PE”) that was positively associated with CO₂ availability. We conclude that the NoPDAT pathway is parallel to and independent of the NoDGAT pathway for oil production. LU-PE can serve as an alternative carbon sink for photosynthetically assimilated carbon in N. oceanica when PDAT-mediated TAG biosynthesis is compromised or under stress in the presence of high CO₂ levels.

The oleaginous alga Nannochloropsis oceanica stores specific phosphatidyldiacylethanolamines, substrates of phospholipid:diacylglycerol acyltransferase, for triacylglycerol production under stress.

Introduction

Generally, two distinct routes, an acyl-CoA-dependent Kennedy pathway and an acyl-CoA-independent pathway, contribute to triacylglycerol (TAG) biosynthesis in many organisms including yeast, land plants, and algae (Sorger and Daum, 2003; Chapman and Ohlrogge, 2012; Xu et al., 2018). Acyl CoA:diacylglycerol (DAG) acyltransferase (DGAT) catalyzes the last acylation step in Kennedy pathway for TAG production. The acyl-CoA-independent pathway for TAG biosynthesis is mediated by phospholipid:DAG acyltransferase (PDAT), transferring an acyl moiety from the sn-2 position of a membrane glycerolipid, usually a phospholipid, to the sn-3 position of DAG (Dahlqvist et al., 2000). In the yeast Saccharomyces cerevisiae, knockout and overexpression (OE) of the PDAT-encoding gene lecithin cholesterol acyltransferase-related open-reading frame (ORF) LRO1 supported its indispensable contribution to TAG biosynthesis (Oelkers et al., 2000), together with the type II DGAT-encoding gene DGA1 (Oelkers et al., 2002; Sandager et al., 2002). Yet the relative contributions of PDAT and DGA1 are growth stage-dependent: PDAT contributes more at the exponential growth stage while DGA1 contributes more at the stationary stage (Oelkers et al., 2002). Albeit showing activities on a broad range of polar lipids, S. cerevisiae PDAT prefers the zwitterionic phospholipids phosphatidylethanolamine (PE) and phosphatidylcholine (PC), particularly the former, as acyl donor for TAG biosynthesis (Dahlqvist et al., 2000; Ghosal et al., 2007; Feng et al., 2019). In contrast to the presence of a single copy of PDAT gene in yeast and algae, many land plants experienced gene expansion events during evolution and contain multiple PDAT copies (Pan et al., 2015; Falarz et al., 2020). Unlike in S. cerevisiae, neither OE nor knockout of PDAT1 in Arabidopsis (Arabidopsis thaliana) affects the seed lipid profiles (Ståhl et al., 2004; Mhaske et al., 2005). Nevertheless, in a dgat1 null mutant of Arabidopsis, PDAT1 knockdown (KD) leads to a more severe decrease in seed TAG level, suggesting that PDAT1 and DGAT1 have overlapping activities at least in seeds (Zhang et al., 2009). In contrast, in actively growing tissues such as developing leaves, PDAT1 functions independently and contributes more than DGAT1 to TAG biosynthesis (Fan et al., 2013a, 2013b). Arabidopsis PDAT1 is believed to possess similar substrate specificity as that of S. cerevisiae PDAT (Fan et al., 2013a, 2013b).

While well documented in yeast and land plants, PDAT from algae has been less scrutinized. Most detailed biochemical and physiological characterization are restricted to green algae, especially Chlamydomonas reinhardtii (Yoon et al., 2012; Liu et al., 2016b). Chlamydomonas reinhardtii PDAT can use a broad range of substrates including phospholipids, glycolipids, and DAG as acyl donors. Still, in vitro enzymatic assays highlighted an apparent preference of C. reinhardtii PDAT for anionic phospholipids such as phosphatidylinositol (PI), phosphatidylserine (PS), and phosphatidylglycerol (PG; Yoon et al., 2012). Intriguingly, it is believed that C. reinhardtii PDAT mediates turnover of chloroplast membrane lipids for TAG biosynthesis particularly under nonstress (favorable growth) conditions (Yoon et al., 2012). In contrast, DGATs in C. reinhardtii are predominantly active under stress rather than favorable growth conditions (Liu et al., 2016a).

Altogether, the roles of PDAT and DGAT seem evolutionarily conserved, namely, PDAT mediates membrane lipid turnover and functions primarily under favorable growth conditions, such as in S. cerevisiae and C. reinhardtii or developing leaves of Arabidopsis, while DGAT function is apparently predominant following growth arrest. The context in which PDAT might operate, that is, in close relation with the endoplasmic reticulum (ER) or the plastid, is unknown in such clades as Stramenopiles/Heterokonts, whose cells contain plastids originating from a secondary endosymbiosis, bounded by four membranes, with the outermost being related to the endomembrane system. This outermost membrane is known as the chloroplast ER (cER) or epiplastid membrane, and its lipid composition has not yet been determined (Petroutsos et al., 2014; Flori et al., 2016; Dolch et al., 2017).

Unlike C. reinhardtii, the stramenopile alga Nannochloropsis oceanica, harboring over 10 DGATs (Vieler et al., 2012b; Li et al., 2014), has been recognized as an industrially important single-cell factory for lipid production because of its fast growth and high lipid content (Vieler et al., 2012a). Although both type I and type II N. oceanica DGATs (NoDGATs) show activities for TAG biosynthesis in vitro, their roles in vivo depend on growth conditions (Wei et al., 2017a; Xin et al., 2017, 2019). The differences in the number of functional DGATs between N. oceanica and C. reinhardtii and the presence of DGATs that contribute to nonstress associated TAG biosynthesis in N. oceanica raise the questions whether N. oceanica PDAT (NoPDAT) is critical, and under what conditions and to what extent it contributes to TAG biosynthesis.

Here, we addressed the function and physiological role of NoPDAT in N. oceanica. NoPDAT resides at the cER, where both plastid and ER lipids are presumed to be present. In vitro assays supported that NoPDAT could use phospholipids and glycolipids and suggested an apparent preference for PG as acyl donor. Genetic analysis showed that NoPDAT contributed at least 30% to TAG biosynthesis under nitrogen limited conditions and seemed determinant in the relative proportion of 16:0 and 16:1 at position sn-3 of TAG. In the cellular context, NoPDAT KD caused an increase, regulated by CO₂ availability, of PE and PG with 16:0, 16:1, or 18:1 at position sn-2 of the lipid glycerol backbone. Our study highlights the role of NoPDAT in mobilizing membrane lipids, particularly PE and PG, in the massive biosynthesis of TAG in N. oceanica, which is parallel and independent to the Kennedy pathway. Moreover, the PE acylated with saturated/monounsaturated fatty acids (FAs) can serve as a potential lipid sink when PDAT-mediated TAG biosynthesis is compromised or under stress with high CO₂ level.

Results

Cloning and bioinformatics analysis of the NoPDAT gene

Assisted with the 5′- and 3′-RACE data, we identified the full-length cDNA of NoPDAT (Supplemental Figure S1). Analysis of the polypeptide sequence of NoPDAT indicated that this protein belongs to the membrane-bound O-acyltransferase family and contains a PC-sterol O-acyltransferase domain, with the presence of a transmembrane domain at the N-terminus (Supplemental Figure S2).

Based on preliminary phylogenetic inferences (Supplemental Methods S1), red algal PDATs were used as outgroup to root the tree. Bayesian and maximum-likelihood (ML) tree topologies were in perfect accordance on the end branches with angiosperms PDAT1 (ML support 100, Bayesian posterior probability [PP] 1) and PDAT2 (98, 1), fungi (100, 1), and green algae (60, 0.98) clustering accordingly together in robust individual clades. Interestingly, the Klebsormidium nitens putative PDAT sequence clustered in both inferences in a basal position to the angiosperms (100, 1), in accordance with Streptophyta’s evolutionary history (Hori et al., 2014; Wang et al., 2020). Surprisingly, stramenopile PDAT sequences did not cluster together, namely, N. oceanica and Microchloropsis gaditana grouped in a sister clade to Phytophthora infestans/Plasmopara halstedii clade with a weak support (37, 0.74); for diatoms, ML did not resolve supraspecific level, while Bayesian analysis robustly (PP = 0.99) clustered diatoms in basal position to the clade grouping plants, fungi, and the other stramenopiles. The polyphyly of stramenopile PDATs may reveal different evolutionary history in diatoms versus eustigmatophyte/oomycetes (Figure 1).

Figure 1.

Bayesian and ML phylogenetic analyses of PDATs from different phyla. The Bayesian tree only is represented here and is drawn to scale, with branch lengths measured in the number of substitutions per site. The PP from the Bayesian analysis and the bootstrap value from the ML analysis are reported on the nodes. The nodes where PP equals 1.0 and ML support equals 100 were removed. – = node not present in ML. Rhodophyte sequences (in the box) were used as outgroup to root both Bayesian and ML trees.

Protein sequence alignment suggested that NoPDAT harbors domains conserved in PDAT polypeptides (Supplemental Figure S2). The use of HECTAR, a method to predict subcellular targeting in Heterokonts/Stramenopiles (Gschloessl et al., 2008), together with other prediction tools such as SignalP, TargetP, and ChloroP, or Mitoprot did not support any localization in chloroplast or mitochondrion. The absence of any targeting sequence to chloroplast appears consistent with NoPDAT sequence grouping also with nonphotosynthetic Stramenopile sequences such as PDAT from oomycota.

Functional complementation of NoPDAT in the TAG-deficient yeast and in vitro assay

The prominent TAG spot resolved by thin-layer chromatography (TLC) (Figure 2A) and green fluorescence following BODIPY staining (Figure 2B) in the TAG-deficient yeast expressing NoPDAT (Supplemental Figure S3) indicated that NoPDAT is functional for TAG biosynthesis and yeast subcellular membranes contain substrates that NoPDAT can possibly utilize. Among the 10 tested polar lipids for the substrate specificity assay of NoPDAT, PG led to the highest activity for TAG formation, while others showed either a low (sulfoquinovosyldiglyceride [SQDG], monogalactosyldiacylglycerol [MGDG], and digalactosyldiacylglycerol [DGDG]), very low or no apparent activity (Figure 2C). This in vitro analysis confirmed that NoPDAT is a glycerolipid acyl transferase, which may accept other substrates than phospholipids, for example, glycolipids.

Figure 2.

Characterization of NoPDAT activity. A and B, Functional complementation of NoPDAT in TAG-deficient S. cerevisiae H1246 cells as determined by TLC analysis of lipid extracts (A) and Bodipy staining (B) of NoPDAT-expressing H1246 cells. Empty vector (EV) and CrDGTT3 (Liu et al., 2016a) were used as negative and positive controls, respectively. C and D, Substrate preference of NoPDAT for polar lipids (C) and DAGs (D). DAG (18:1/18:1) was used as the acyl acceptor in (C) and PG was used as the acyl donor in (D). Data represent mean ± sd (n = 3). Bar, 5 μm. FFA, free FA; Ori, origin of migration; and PA, phosphatidic acid.

We also tested the preference of NoPDAT for the structure of the DAG acceptor, including seven 1,2-DAG molecular species and one 1,3-DAG, using PG as the acyl donor (Figure 2D). NoPDAT could operate on a broad spectrum of DAG acceptors with a higher apparent activity on 18:1/18:1, 16:0/18:1, and 18:1/16:0. We did not evaluate the main DAG donor from N. oceanica, 16:1/16:0 (Nobusawa et al., 2017), which was not commercially available.

Subcellular localization of NoPDAT at the cER

Laser confocal scanning microscopy analyses showed that in the control transformants the enhanced green fluorescent protein (eGFP) signal (green) was present in the cytosol, not overlapping with the plastid autofluorescence (PAF, red) (Figure 3, A and D). In contrast, in the NoPDAT-eGFP transformants, the green eGFP signal formed irregular layers surrounding the plastid (Figure 3B), which was further supported by a 3D visualization of eGFP and PAF signals (Figure 3E). The distribution pattern of NoPDAT-eGFP was similar to that of endogenous ER stained with ER tracker, and the signal was visualized mainly around the plastid (Figure 3C; Supplemental Figure S4), consistent with the reported ER- or cER-localized proteins (Moog et al., 2015; Wei et al., 2017a, 2017b; Shi et al., 2021). We also introduced NoPDAT-eGFP into Phaeodactylum tricornutum, a Heterokont alga with bigger cells, in which there is experimental demonstration of cER-localized proteins. Clearly, the eGFP signal surrounded the plastid and nucleus (blue; Figure 3F), resembling the localization pattern of P. tricornutum hDer1-2, a confirmed cER-localized protein (Hempel et al., 2009). These results suggest that NoPDAT resides at the cER.

Figure 3.

Subcellular localization of NoPDAT in N. oceanica and P. tricornutum cells. A–C, Nannochloropsis oceanica cells transformed with the EV (eGFP, A) and NoPDAT-eGFP (B, C). D and E, A 3D visualization of PAF+GFP in (A) and (B), respectively. F, Phaeodactylum tricornutum cells transformed with NoPDAT-eGFP. BF, bright field; PAF, plastid autofluoresence; H, Hoechst. Red indicates the PAF, green indicates the GFP signal, yellow indicates the staining of ER with ER-tracker red dye, while blue indicates the staining of nucleus with Hoechst (H). Arrowheads indicate co-localization of ER and GFP fluorescence. Bar, 3 μm.

NoPDAT OE promotes TAG accumulation and suggests a consumption of 16:0 and 16:1 in polar lipids

Two lines, PDAT-OE3 and PDAT-OE5, which showed the maximal increase (>10-fold) in NoPDAT transcripts compared with wild-type (WT), were selected from 10 NoPDAT-overexpressing lines for subsequent analysis (Figure 4A). Immunoblot analysis revealed that NoPDAT protein abundance was higher in the two transformants than in WT (Figure 4B; Supplemental Figure S5). During a 10-day batch culture period, no difference in the cell density was observed between the transformants and WT (Figure 4C), suggesting that NoPDAT OE had no effect on the growth of N. oceanica. Accordingly, the nitrate concentration in the culture medium showed no difference between the transformants and WT and was almost consumed within 6 days (Figure 4D). Fluorescence intensity assay of Nile red-stained algal cells indicated that N. oceanica accumulated TAG after 4 days of culture and the transformants had a higher TAG content than WT during the late culture days (Days 6–10; Figure 4E). Gas chromatography–mass spectrometry (GC–MS) quantification revealed that NoPDAT OE enhanced TAG by 28%–33%, yet having few consequences on the content of total FA (TFA) (P > 0.05) (Figure 4F). NoPDAT OE had a limited effect on the FA profile of TAG, whereas a slight decrease of 16:0 (P < 0.05 for PDAT-OE3) and increase of 20:5 (P < 0.05) were noticed in TFA (Figure 4, G and H).

Figure 4.

Characterization of NoPDAT OE lines of N. oceanica. A, NoPDAT mRNA level in WT and the two NoPDAT OE lines (PDAT-OE3 and PDAT-OE5), as determined by RT-qPCR. The NoPDAT expression in WT on Day 2 was used as the reference (set as 1). B, NoPDAT protein level in WT and the two NoPDAT OE lines, as determined by immunoblotting. C, Growth (static condition) curve as determined by cell density. D, Nitrate concentration in the culture medium. E, Fluorescence intensity (a.u.: arbitrary units) of Nile Red-stained algal cells. f, Content (DW: dry weight) of TFA and TAG from the 8-day cultures. G and H, Relative FA abundance of TFA (G) and TAG (H) from the 8-day cultures. Data in (A and C–H) represent mean ± sd (n = 3). * and different letters above the bars indicate statistically significant differences (P < 0.05) by t test (A) and one-way ANOVA (F–H), respectively.

NoPDAT KD attenuates TAG accumulation and decreases the proportion of TAG with 16:0 and 16:1

Two out of ˃10 RNAi lines, PDAT-KD2 and PDAT-KD4 that had the lowest NoPDAT transcript abundance (<10% of that in WT on Day 8), were chosen for subsequent experiments (Figure 5A). Similar to the OE, NoPDAT KD had little impact on the growth of N. oceanica (Figure 5B). Nile red staining and fluorescence intensity assay indicated that the KD lines had a TAG content lower than WT, particularly during the late culture days (Figure 5C). GC–MS quantification confirmed the decrease in TAG (reduced by 26%–38%) caused by NoPDAT KD, whereas TFA content showed little change (Figure 5D). NoPDAT KD increased the relative abundance of 16:0 (P < 0.05) and 16:1 (P < 0.05 for PDAT-KD2) and decreased that of 20:4 and 20:5 (P < 0.05) in TFA; changes of FA profile of TAG were observed in line PDAT-KD2, and the relative abundance of 16:1 and 18:1, in particular, increased (P < 0.05) (Figure 5, E and F). Assuming that TFA reflects FA contained in membrane lipids and TAG, the differences observed in TFA should reflect an impact on membrane glycerolipids. Thus, a KD of NoPDAT clearly leads to the accumulation of 16:0 and 16:1 in membrane glycerolipids.

Figure 5.

Characterization of NoPDAT KD lines of N. oceanica. A, NoPDAT mRNA level in WT and the two NoPDAT KD lines (PDAT-KD2 and PDAT-KD4), as determined by RT-qPCR. The NoPDAT expression in WT on Day 2 was used as the reference (set as 1). B, Growth (static condition) curve as determined by cell density. C, Fluorescence intensity of Nile Red-stained algal cells. D, Content of TFA and TAG from the 8-day cultures. E and F, Relative FA abundance of TFA (E) and TAG (F) from the 8-day cultures. Data represent mean ± SD (n = 3). * and different letters above the bars indicate statistically significant differences (P < 0.05) by t test (A) and one-way ANOVA (D–F), respectively.

To uncover the NoPDAT KD-associated global changes of TAG species in a time-resolved manner, TAG quantification by electrospray ionization MS (ESI–MS) analysis was conducted. There was no difference in total TAG content between PDAT-KD4 and WT on Day 3, a time point before massive TAG accumulation in N. oceanica cells; in contrast, PDAT-KD4 showed a significantly lower total TAG content than WT on Days 5 and 7 (Figure 6). These results suggest that in N. oceanica, NoPDAT had little impact on the basal levels of TAG accumulation under favorable growth conditions, but likely acts under nutrient stress, opposite to the function of C. reinhardtii PDAT (Yoon et al., 2012).

Figure 6.

Heat map illustrating the variation of TAG species. The data are expressed as log₂(fold change) values (n = 3) for WT (Days 5 and 7, versus Day 3), NoPDAT KD line (PDAT-KD4) (Days 5 and 7, versus Day 3), PDAT-KD4 versus WT (Days 3, 5, and 7), and PDAT-KD4 on Day 5 (static versus Air and CO₂ versus Air). The circles at the left of the heat maps designate the content of individual TAG species for WT on Day 3 (first circle column), PDAT-KD4 on Day 3 (second circle column), and PDAT-KD4 on Day 5 (third circle column). Static, static culture; Air, culture aerated with air; CO₂, culture aerated with 2% CO₂. Significant difference (Student’s t test, P < 0.01) is indicated with an asterisk.

More than 50 TAG molecular species were identified in N. oceanica (Figure 6; Supplemental Dataset 1). Over time (e.g. on Days 5 and 7), most TAG species followed a similar accumulation pattern and increased considerably in WT and PDAT-KD4. Albeit having no impact on total TAG content in N. oceanica on Day 3, NoPDAT KD modulated the content of individual TAG species, which were not initially visible when simply considering the TAG FA profile. Four major TAG species, that is, 16:1/16:1/16:1, 16:1/16:1/16:0, 14:0/16:1/16:1, and 16:1/16:0/14:0, decreased following NoPDAT KD, highlighting mainly a defect in transfer of 16:0 and 16:1 from membrane lipids. On Day 3, this trend was also observed for minor TAG species and conversely, the proportion of TAG species containing 20:4 or 20:5 at positions sn-1 or sn-3 increased. On Day 5, all TAG species showed a significant decrease in the NoPDAT KD lines, consistent with a lowering of the TAG production, and the capacity of NoPDAT to transfer other acyls than 16:0 or 16:1.

NoPDAT KD leads to an accumulation of PE and PG enriched in 16:0, 16:1, and 18:1

Through the semi-quantitative TLC analysis of polar lipids, one band clearly accumulated in the NoPDAT KD lines (Figure 7A, left). This band was recovered, subjected to MS analysis, and characterized as PE enriched in saturated/monounsaturated FAs. Henceforth, the TLC spot characterized by such PE molecular species is referred to as “less-unsaturated PE” or LU-PE. Quantification via ESI–MS analysis was then conducted for WT and PDAT-KD4 cultures harvested on Days 3, 5, and 7 under air condition. WT showed a decrease in the overall PE content in the course of cultivation (Figure 7B) and the PE profile was predominantly more unsaturated, with 20:4/20:4, 20:5/20:4, 20:4/20:3, and 20:5/20:5 accounting for over 80% of PE (Figure 7C), consistent with previous observation in Nannochloropsis species (Alboresi et al., 2016; Nobusawa et al., 2017). In PDAT-KD4, the consumption of PE was arrested (Figure 7B). Thus, although NoPDAT had no apparent activity on PE in in vitro enzymatic assay (Figure 2C), PDAT-KD4 accumulated a much higher level of this membrane lipid than the WT (∼2.2-fold) regardless of the culture time points (Figure 7B). In PDAT-KD4, the molecular species of PE, which showed an accumulation, were those detected by TLC, that is, 16:0/16:1 (>100-fold higher than that in WT), followed by 16:1/18:1, 16:1/16:1, 16:0/18:1, 16:0/16:0, 14:0/16:0, and 14:0/16:1 (Figure 7C), corresponding to LU-PE. Considering the sn-2 position in LU-PE molecular species, PE appears therefore as a major donor of 16:0, 16:1, and 18:1, for NoPDAT action.

Figure 7.

Effect of NoPDAT KD on polar lipids of N. oceanica. A, TLC analysis of lipid extracts from WT and NoPDAT KD lines (PDAT-KD2 and PDAT-KD4) aerated with air (left) and PDAT-KD4 under different CO₂ availability conditions (right). B, Content of polar lipids. C–E, Content of PE species (C), PG species (D), and PME species (E). Static, static culture; Air, culture aerated with air; CO₂, culture aerated with 2% CO₂. Data represent mean ± sd (n = 3). * and different letters above the bars indicate statistically significant differences (P < 0.05) by t test (WT and PDAT-KD4) and one-way ANOVA (PDAT-KD4 under different culture conditions), respectively. MU-PE: “more-unsaturated PE,” PE enriched in 20:3, 20:4, and 20:5 FAs; LU-PE: “less-unsaturated PE,” PE enriched in saturated or monounsatured FAs.

In line with the strong in vitro activity of NoPDAT on PG (Figure 2C), the PDAT-KD4 line was marked by a 50%–80% higher (P < 0.05) level of this lipid class compared to the WT (Figure 7B). A total of nine PG species were detected, and like for PE, those harboring a 16:0 or 16:1 at position sn-2 accumulated in the PDAT-KD4 line (Figure 7D). However, different from PE that tended to increase in PDAT-KD4 line, PG decreased gradually in both WT and PDAT-KD4 cells as the culture persisted.

Minor quantitative variations (P > 0.05) and even decreases (P < 0.05) were noticeable for phospholipids such as PC or PI (Figure 7B), consistent with in vitro tests (Figure 2C), except for a 25% increase (P < 0.05) of PC at Day 3 (Figure 7B). Contrary to the results obtained with plastid glycolipids in in vitro assays, MGDG levels at Days 3 and 7 or DGDG levels at Day 7 changed little (P > 0.05) in the PDAT-KD4 line compared to the WT, however, DGDG levels increased by 14%–17% (P < 0.05) at Days 3 and 5 (Figure 7B). As for the FA profiles of MGDG, DGDG, or PC, a slight enrichment in molecular species measured in the PDAT-KD4 line concerned 16:0 or 16:1 at the sn-2 position, which could be related to a utilization by NoPDAT (Supplemental Figure S6). These findings suggest that NoPDAT may have poor activity on PC, PI, MGDG, or DGDG in in vivo conditions, either by a lack of specificity or due to a location of the enzyme with limited access to these lipids. Diacylglyceryl-N,N,N-trimethylhomoserine (DGTS) relative quantity was evaluated based on the staining intensity on TLC plate, showing no great change except for Day 3 in the PDAT-KD4 line (Supplemental Figure S7). Eventually, plasmenylethanolamine (PME) was detected in trace amount in N. oceanica and accumulated considerably (∼15-fold) following NoPDAT KD, showing that this unusual phospholipid, structurally close to PE, could also serve as acyl donor for NoPDAT (Figure 7E).

Transcriptional response of lipid metabolic genes to NoPDAT KD

To investigate the effect of NoPDAT KD on lipid metabolism network at the transcriptional level, reverse transcription quantitative PCR (RT-qPCR) was conducted for 55 lipid metabolic genes in WT and PDAT-KD4 (Figure 8; Supplemental Dataset S2). Enzymes involved in de novo FA biosynthesis were transcriptionally downregulated in response to NoPDAT KD, which is consistent with a slightly lower TFA content in NoPDAT KD lines. The Δ6-FA desaturase (Δ6 FAD) that introduces a double bond between carbons 6 and 7 on C18 acyls and FA elongase2 (FAE2) that catalyzes the formation of C20 acyls from C18 acyls were also downregulated in the NoPDAT KD line. This may partially explain that the KD strains had lower percentage of very long-chain FAs 20:4 and 20:5 (Figure 5E). However, transcript levels of enzymes involved in the Kennedy pathway (glycerol-3-phosphate acyltransferase: GPAT, lysophosphatidic acid acyltransferases: LPATs, and DGATs) were little impacted by NoPDAT KD. This indicates that a lack of NoPDAT does not trigger any compensatory mechanism via the acyl-CoA-dependent pathway, consistent with the observed impact of OE and KD on TAG levels. The PDAT pathway appears therefore independent from the Kennedy pathway. In addition, although PE, PG, and PME increased considerably (Figure 7), no transcriptional upregulation of their biosynthesis was observed in response to NoPDAT KD.

Figure 8.

The effect of NoPDAT KD on lipid metabolism network in N. oceanica. Four time points were selected for RT-qPCR. Genes were considered to be significantly differentially expressed if: the expression level fold change between NoPDAT KD line (PDAT-KD4) and WT was no less than two-fold with P < 0.01 for at least one time point or no ˂1.5-fold with P < 0.01 for at least two time points. The genes in blue and red were downregulated and upregulated, respectively. AAD, acyl-ACP desaturase; ACC, acetyl-CoA carboxylase; ADHAPR, acyl/alkyl-dihydroxyacetone phosphate reductase; ADHAPS, alkyl-dihydroxyacetone phosphate synthase; CDIPT, CDP-DAG-inositol 3-phosphatidyltransferase; CDS, CDP-DAG synthase; Δ12 FAD, Δ12-FA desaturase; Δ5 FAD, Δ5-FA desaturase; Δ6 FAD, Δ6-FA desaturase; Δ9 FAD, stearoyl-CoA 9-desaturase; DGDGS, digalactosyldiacylglycerol synthase; DGK, diacylglycerol kinase; ENR, enoyl-ACP reductase; FAE, FA elongase; GPAT, glycerol-3-phosphate acyltransferase; HAD, hydroxyacyl-ACP dehydrase; IPCS, inositol phosphorylceramide synthase; KAR, ketoacyl-ACP reductase; KAS, ketoacyl-ACP synthase; LACS, long-chain acyl-CoA synthetase; LPAT, lysophosphatidic acid acyltransferases; LPCAT, lysophosphatidylcholine acyltransferase; LPLAT, lysophospholipid acyltransferase; MCAT, malonyl-CoA:ACP transacylase; PAP, phosphatidic acid phosphatase; PC-PLD, phosphatidylcholine-specific phospholipase D,; PDH, pyruvate dehydrogenase; PEAMT, phosphoethanolamine N-methyltransferase; PGPP, phosphatidylglycerolphosphate phosphatase; PLA, phospholipase A; PLMT, phospholipid N-methyltransferase.

CO₂ availability affects the NoPDAT KD-associated accumulation of PE enriched in 16:0, 16:1, and 18:1

The accumulation of LU-PE was observed in NoPDAT KD cultures aerated with air and was visible by TLC analysis (Figure 7A, left), however, it was absent in PDAT-KD4 line grown at static condition (Figure 7A, right). For the WT, the LU-PE band was not visible even with the aeration of 2% CO₂ until Day 7, let alone the static (Supplemental Figure S8a) and air conditions (Figure 7A, left). Accordingly, PDAT-KD4 aerated with 2% CO₂ accumulated more LU-PE than air condition as shown on TLC plate (Figure 7A, right).

On Day 5, nitrate in the medium was nearly exhausted under static, air, and 2% CO₂ conditions (Supplemental Figure S9), and the comparable cell density (Figure 5; Supplemental Figure S10) was achieved in the PDAT-KD4. To gain the detailed changes in response to CO₂ availability, quantification of glycerolipids in 5th-day PDAT-KD4 via ESI–MS was performed. Unlike those PE enriched in 20:4/20:4, 20:5/20:4, and 20:4/20:3 that had the highest level in cultures aerated with air, LU-PE species were all associated with the CO₂ availability and their levels followed the order of static cultures < cultures aerated with air < cultures aerated with 2% CO₂(P < 0.05) (Figure 7C). Similarly, the PG enriched in 16:0, 16:1, and 18:1 in PDAT-KD4 were positively associated with CO₂ availability (Figure 7D). PME species were also increased by CO₂ availability: the higher CO₂ supply, the higher level of PME (Figure 7E). Under static condition, LU-PE maintained a rather low level in PDAT-KD4 line on Day 5, though it was higher than that on Day 5 in WT aerated with air. Moreover, LU-PE content of PDAT-KD4 under static condition was comparable to that on Day 3 and less than a half of that on Day 7 in WT aerated with air, and it was ˂10% and 5% of that of PDAT-KD4 aerated with air and 2% CO₂, respectively.

PDAT mRNA level gradually increased in the WT culture aerated with air and maintained the highest from Days 5–7 (Supplemental Figure S8b). Immunoblotting analysis showed that increasing CO₂ supply (from air to 2% and 5% CO₂) in the culture enhanced PDAT expression (Supplemental Figure S8c). It was evident that accumulation of LU-PE in WT cells with elevated CO₂ supply was not attributed to the downregulated PDAT, showing that LU-PE may act as carbon sink under stress with high CO₂ level. In fact, aeration with 5% CO₂ led to a visible LU-PE band not only in WT but also in PDAT OE strains (Supplemental Figure S8a).

Although TAG species responded differentially to CO₂ availability in PDAT-KD4, the major more-unsaturated TAG species such as 52:6 (20:5/16:0/16:1), 52:5 (20:4/16:1/16:0), and 52:4 (20:3/16:0/16:1) were associated with CO₂ supply and their levels followed the order of static cultures < cultures aerated with air < cultures aerated with 2% CO₂(P < 0.05) (Figure 6). These data suggest under high CO₂ supply conditions, acyl-donors with 16:0 and 16:1 are likely partitioned preferentially to PE (maybe also PG) while acyl-donors with 20:4 and 20:5 are partitioned preferentially to TAG, with LU-PE and TAG serving as carbon sinks.

LU-PE accumulation is independent from genes involved in PE homeostasis

The analysis of the NoPDAT KD line suggests that this enzyme is responsible for the specific consumption of LU-PE. We wondered whether other enzymes involved in the production or catabolism of PE could also control LU-PE. We performed OE and/or KD of a series of genes (Figure 9; Supplemental Figure S11), including those encoding the putative CPT/EPT (the bifunctional enzyme responsible for the last biosynthetic step of PE and PC), a PSS that converts PE into PS, a PE N-methyltransferase (PEMT) and a phospholipid N-methyltransferase (PLMT) that catalyze the methylation of PE to form PC, and the phosphoethanolamine N-methyltransferase (PEAMT) that converts phosphoethanolamine to phosphocholine for subsequent PC synthesis (Zienkiewicz et al., 2020). TLC was used to screen phenotypes, based on the detection of the specific LU-PE band, used as readout. Overexpressing of each of the two CPT/EPT genes did not induce any accumulation of LU-PE in N. oceanica (Supplemental Figure S12). The turnover of LU-PE species is therefore unlikely counterbalanced by an increase in PE production. Likewise, OE of PSS or KD of PSS, PEMT, PLMT, or PEAMT did not lead to any LU-PE accumulation (Supplemental Figure S12). Altogether, these results indicated that LU-PE accumulation in N. oceanica is not linked to the genes involved in PE homeostasis and that NoPDAT is solely responsible for the observed profile change.

Figure 9.

A hypothesized working model illustrating the role of NoPDAT in lipid metabolism in N. oceanica. Downregulated genes and decreased metabolite by PDAT KD are illustrated in blue, and upregulated gene, increased metabolites, and carbon flux by PDAT KD in red. KD and OE of genes involved in PE homeostasis that were carried out in the study are also shown in the model with KD or OE symbols below gene names. PGA, 3-phosphoglyceric acid; G3P, glycerol-3-phosphate; DHAP, dihydroxyacetone phosphate; CDP-Etn, cytidine diphosphate-ethanolamine; P-Etn, phosphoethanolamine; P-Cho, phosphocholine; P-MMEtn, phospho-monomethyl-ethanolamine; CPT, cholinephosphotranferase; EPT, ethanolaminephosphotranferase; PLMT, phospholipid N-methyltransferase; PEAMT, phosphoethanolamine methyltransferase; LU-PE, “less-unsaturated PE,” PE enriched in saturated or monounsaturated FAs.

Discussion

NoPDAT is a canonical glycerolipid:DAG acyltransferase using mainly PG and PE as acyl donors in vivo

PDATs can utilize various substrates as acyl donors, yet the activity and substrate preference are dependent on the PDAT sources. Both S. cerevisiae and Arabidopsis PDATs prefer to transfer the acyl moiety from PE and PC, particularly the former, to DAG for TAG biosynthesis (Dahlqvist et al., 2000; Ghosal et al., 2007; Fan et al., 2013a, 2013b; Feng et al., 2019). Chlamydomonas reinhardtii PDAT, on the other hand, exhibits less activity on PE and PC for in vitro (Yoon et al., 2012).

Here, our in vitro results showed that NoPDAT having no detectable activity on PE or PC exhibited a high activity for PG with the apparent preference for molecular species harboring 16:0 and 16:1 at the sn-2 position (Figure 2C; Supplemental Figure S13). Interestingly, NoPDAT KD increased not only PG but also PE levels considerably in N. oceanica (Figure 7B). It must be noticed that the lipidic substrates, obtained from commercial suppliers, contained molecular species differing from those occurring naturally in N. oceanica. Considering all in vitro and in vivo results, NoPDAT is a canonical glycerolipid:DAG acyltransferase, which can accept certain acyl-donors with 16:0, 16:1 and to some extent 18:1 at position sn-2, and the main substrates of NoPDAT in vivo appear to be PE and PG.

The substrate variation of PDATs may arise from their diversification during evolution. Green and red algae originated from a primary endosymbiosis, whereas Stramenopile/Heterokonts emerged from a secondary endosymbiosis, in which eukaryotic hosts engulf a red alga (Yoon et al., 2002; Füssy and Oborník, 2018). Although both N. oceanica and P. tricornutum belong to Heterokont algae, the balance between secondary host versus endosymbiont genetic importance differed over the course of speciation. In N. oceanica, the symbiont nucleus likely dominated over the host nucleus, whereas in P. tricornutum the symbiont nucleus was probably discarded (Guo et al., 2019). It is possible that N. oceanica and P. tricornutum PDATs were inherited from the symbiont and host nuclei, respectively. Consistent with this hypothesis, the PDAT sequences from Eustigmatophytes and Bacillariophyceae did not group together in reconstructed phylogenetic tree (Figure 1). It has been proposed that PDATs evolution could be driven by different selection constraints and that positive selection may play critical roles in PDAT function (Falarz et al., 2020). More functional data need to be gathered in the Heterokont/Stramenopile phylum to investigate this hypothesis.

NoPDAT contributes to membrane lipid turnover during stress-induced TAG biosynthesis

In the batch cultures, N. oceanica showed a rapid growth within the first 4 days and then the algal growth slowed down (Figures 4C and 5B), indicative of the onset of nutrient limitation stress. Consistently, the nitrate content in the culture was drastically consumed within the first 4 days (Figure 4D) and thereafter TAG began to accumulate (Figures 4E and 5C). Neither NoPDAT OE nor NoPDAT KD caused any important change in TAG content of N. oceanica before Day 4, thus considered as the favorable growth (nutrient replete) condition with a basal level of synthesized TAG. In contrast, after 4 days of cultivation, when the algal cells were exposed to nutrients’ limitation, a striking increase of TAG was observed in NoPDAT overexpressing lines and an opposite trend was observed in NoPDAT KD lines (Figures 4–6). These results suggest a substantial contribution of NoPDAT to TAG production at the onset of the stationary phase. In line with this deduction, NoPDAT transcripts in the WT showed a moderate increase on Days 5 and 8 compared with Day 2 (Figures 4A and 5A). Therefore, NoPDAT may function as a TAG contributor in N. oceanica under stress, but not under favorable fast-growing conditions, differing from C. reinhardtii and S. cerevisiae or developing leaves of Arabidopsis in which PDAT contributes mainly to TAG accumulation under favorable growth conditions (Oelkers et al., 2002; Yoon et al., 2012; Fan et al., 2013b). Nevertheless, the analysis of KD lines showed that even in early cultivation stages, the acyl-donors of PDAT, that is, LU-PE, accumulated. In addition, we observed a decrease of several TAG species caused by NoPDAT KD on Day 3, though the total TAG level showed little change (Figure 6). Thus, we cannot rule out the possibility that NoPDAT operates on a basal level and contributes to nonstress associated TAG synthesis as well, but not detectable compared with the bulk produced via the Kennedy pathway, as several NoDGATs have been demonstrated to contribute mainly to TAG biosynthesis in N. oceanica under nonstress conditions (Xin et al., 2017; Wei et al., 2017a).

Parallel and possibly independent TAG production via NoPDAT and NoDGAT pathways

Of the three NoDGATs that function mainly under stress, KD of NoDGAT1A, NoDGAT2A, and NoDGAT2D leads to TAG decrease by 25%, 26%, and 47%, respectively (Wei et al., 2017a; Xin et al., 2017). In our study, NoPDAT KD decreased TAG content by ∼32% (Figure 5D). Although these experiments were not strictly conducted under the same conditions and the gene KD efficiencies may vary, NoPDAT likely makes a comparable contribution to stress-induced TAG accumulation as each of the above-mentioned NoDGATs. However, if summing up the TAG decrease ratios caused by silencing each of the four acyltransferase genes in N. oceanica, the value exceeds 100%. In this context, these acyltransferases may have overlapping roles. The interactions between enzymes involved in TAG synthesis including DGAT and PDAT have been reported in mammals (Man et al., 2006; Jin et al., 2014), plants (Lee and Seo, 2019; Xu et al., 2019a), and algae (Xu et al., 2019b). In Arabidopsis, “oil-producing metabolons” were recently characterized, containing either DGAT1 or DGAT2 and PDAT, with accesses to separate substrate pools (Regmi et al., 2020). It would be important to evaluate whether similar metabolons might occur in a species with many oil-producing enzymes. In this study, we noticed little impact on the expression of genes coding for GPAT, LPATs, and DGATs in response to NoPDAT KD (Figure 8). Only a down-tuning of FA synthetic genes seemed to occur.

Altogether, NoPDAT is likely to operate as a parallel and independent oil-producing route, possibly dispensable to feed an actual form of carbon storage, and more important as an actor of membrane lipid turnover (Fan et al., 2014). The occurrence of such PDAT-mediated membrane lipid turnover in N. oceanica is further supported by the following evidence. First, as the culture time lasted, NoPDAT showed a steady increase in its transcript level (Figures 4A and 5A; Supplemental Figure S8b), accompanied by the decrease of many membrane lipids (Figure 7B). Second, following NoPDAT KD, membrane lipids particularly PG and PE exhibited a considerable increase regardless of the culture time points examined (Figure 7B).

NoPDAT location at the cER highlights the presence of both PG and PE in the outermost membrane of the plastid

Phosphatidylglycerolphosphate synthase (PGPS), responsible for PG production, has two possible forms: the prokaryotic form in the plastid of Archaeplastida, and the eukaryotic form in the mitochondrion (Luévano-Martínez, 2015). The situation in Heterokont has not been established, yet both types are encountered in N. oceanica based on sequence similarity search, suggesting the presence of PG biosynthesis in both the plastid and mitochondrion. Eventually, PG might also be exported to the endomembrane system via inter-organellar transport (Vance, 2015). In N. oceanica, the predominance of C16 acyls over C18 acyls in PG sn-2 position (Figure 7D) suggests that the bulk of PG is synthesized in the plastid. Considering the subcellular localization of NoPDAT at the cER (Figure 3), it could possibly access PG from both the ER (low abundance) and the plastid outer envelope. NoPDAT may not use PG species located at the thylakoid membrane or the chloroplast inner envelope per se, unless this lipid is translocated to the cER, which needs to be assumed to interpret the KD phenotype.

PE represents a major nonbilayer phospholipid widely distributed in the extra-chloroplastic membranes of Archaeplastida. PE biosynthesis involves mainly two metabolic pathways, that is, an ER-localized CDP-ethanolamine pathway (de novo biosynthetic pathway) and a mitochondrial PS decarboxylation pathway, with the former being the major one (Gibellini and Smith, 2010). In N. oceanica, PE is present predominantly in the form of molecular species with no less than four double bonds (Figure 7C; Li et al., 2014; Han et al., 2017; Meng et al., 2017). Based on our analysis of genes involved in the biosynthesis or conversion of PE, the produced LU-PE was mainly consumed by cER NoPDAT to form TAG and otherwise accumulated, and could not be converted to PE enriched in 20:3, 20:4, and 20:5. The cER thus appears as a major membrane lipid remodeling site, under nutrient stress, marked by a consumption of PE and PG molecular species.

LU-PE serves as an alternative carbon reservoir in N. oceanica under stress

Since LU-PE species are the acyl-donors for NoPDAT, KD of NoPDAT could result in LU-PE accumulation, which is boosted by elevated CO₂ supply (Figure 7; Supplemental Figure S8). Thus, LU-PE may serve as an alternative carbon reservoir for photosynthetically assimilated carbon. The large pool of LU-PE is produced in response to nutrient limitation when TAG biosynthesis is impaired or under high CO₂ conditions, which indicated a potential protection mechanism in cells under stress. The protective role of LU-PE is further supported as there’s no difference between the growth of WT and NoPDAT KD lines under static condition, despite lower TAG content in the latter. In contrast, KD of PDAT and knockout of pgd1 in C. reinhardtii decreased cellular TAG content and their growth rates in the mid-exponential growth phase and stationary phase under stress, respectively (Li et al., 2012; Yoon et al., 2012), supporting a role of TAG in protecting algal cells against oxidative damage. In NoPDAT KD lines, this protective role appears to be replaced by accumulation of LU-PE as an alternative electron and carbon sink against overreduction of photosynthetic electron transport chain and possibility production of reactive oxygen species under stress. Growth of N. oceanica strain MBIC10090 was lower at 2% and 5% CO₂ than under air condition (Supplemental Figure S10) and it appears that sensitivity to varying levels of CO₂ is strain/species specific for Nannochloropsis (Chiu et al., 2009; Wei et al., 2017b, 2019). However, the growth of PDAT–KD4 was similar between air and 2% CO₂ conditions, which indicates that LU-PE accumulation could alleviate the growth inhibition induced by excessive CO₂ supply. Furthermore, the presence of bigger cells in the PDAT KD lines (Supplemental Figure S14) might be beneficial to the carbon utilization under higher CO₂ conditions. Nevertheless, higher CO₂ resulted in lower contents of more-unsaturated MGDG and DGDG (Supplemental Figure S6), the main glycolipids in N. oceanica plastid (Mühlroth et al., 2017), which might contribute to the reduced growth at 5% CO₂.

Considering that PE enriched in 20:3, 20:4, and 20:5 showed no increase while LU-PE rose drastically (Figure 7C), the FA elongases and desaturases (Poliner et al., 2018) may not function on the acyl groups of those LU-PE species in N. oceanica (Figure 9). Due to the greatly different acyl-chain length and unsaturation of LU-PE, if the vastly increased LU-PE species are transported to the biologically functional two-layer membranes, the membrane structure and function would be changed. To avoid such a drastic change, LU-PE species may be stored on the monolayer of lipid droplets (LDs), which generally occurs in algal cytosol under stress to accommodate TAG (Wang et al., 2009; Vieler et al., 2012a). Recently, Lee et al. (2020) found that PE is essential for anchoring Delayed in TAG Hydrolysis 1, involved in the initial stage of LD degradation, to LDs in Chlamydomonas. More researches are to be carried out to identify the location and function of LU-PE in cells in the future.

Materials and methods

Algal strains

Nannochloropsis oceanica (MBIC10090) and P. tricornutum (CCMP2561) were obtained from the Marine Biotechnology Institute Culture Collection (Iwate, Japan) and the culture collection of the Provasoli-Guillard National Center for Culture of Marine Phytoplankton (Maine, USA), respectively. Algal cultures were grown axenically in artificial seawater enriched with f/2 nutrients (500 μM nitrate; Guillard, 1975) at 22°C under continuous illumination of 120 μmol photons m⁻² s⁻¹. Transformants were grown in medium with the presence of 1 (for N. oceanica) or 40 μg mL⁻¹ (for P. tricornutum) zeocin.

Identification of putative PDAT encoding gene in N. oceanica and phylogenetic analyses

The full-length coding regions of NoPDAT were obtained based on the annotation information in JGI N. oceanica CCMP1779 (Supplemental Figure S1) and further verified by 3′- and 5′-RACE. For phylogenetic analyses (Supplemental Figure S15 and Supplemental Datasets S3–S5), a final dataset (Supplemental Figure S16 and Supplemental Dataset S6) composed of 30 sequences was aligned, and a ML and a Bayesian phylogenetic inference were carried out. All details were in Supplemental Methods S1. Subcellular targeting predictions of NoPDAT were carried out using SignalP (https://services.healthtech.dtu.dk/service.php?SignalP-4.1), TargetP (https://services.healthtech.dtu.dk/service.php?TargetP-2.0), ChloroP (https://services.healthtech.dtu.dk/service.php?ChloroP-1.1), Mitoprot (https://ihg.gsf.de/ihg/mitoprot.html), and HECTAR (https://webtools.sb-roscoff.fr/).

NoPDAT expression in yeast, yeast microsome preparation, and in vitro PDAT activity assay

The ORF of NoPDAT was cloned into pYES2-CT (Invitrogen; V825120, Waltham, MA, USA) and the resulting pYES2-NoPDAT was introduced into the S. cerevisiae TAG-deficient strain H1246 for expression. Lipid extraction from yeast cells and TLC analysis were performed according to Liu et al. (2016a). BODIPY 493/503 (Invitrogen), a neutral lipid-specific fluorescent dye, was used for the staining of yeast cells.

The expression of NoPDAT protein in H1246 (Supplemental Figure S3), microsome preparation, and in vitro PDAT assay was conducted as described by Liu et al. (2017). A total of 10 polar lipids and 8 DAGs were used for the assay (details in Supplemental Methods S1).

GFP fusion and OE constructs

For the localization analysis of NoPDAT, two constructs (pPha-NoVCPp-NoPDAT-eGFP and pPhaT1-NoPDAT-eGFP) were generated to express C-terminal eGFP fusion proteins in N. oceanica and P. tricornutum cells, respectively. For the OE of NoPDAT, cholinephosphotransferase (CPT), ethanolaminephosphotransferase (EPT), and phosphatidylserine synthase (PSS) in N. oceanica, their ORFs were amplified using primer pairs listed in Supplemental Dataset S7, then ligated into the pPha-NoVCPp vector (Chen and Hu, 2019). Details of vector construction, and introduction of linearized vectors into WT, and observation of GFP positive cells were in Supplemental Methods S1.

RT-qPCR and western blot analysis

Total RNA extraction, cDNA synthesis, RT-qPCR analysis, protein preparation, and western blotting were performed as described in Supplemental Methods S1. Internal control genes were chosen according to Mühlroth et al. (2017) and primers used were shown in Supplemental Dataset S7.

Growth and Nile red staining

Growth was determined by counting cell numbers and the initial inoculation density was about 1 × 10⁶. Nitrate concentration in the culture was determined using spectrophotometry (Collos et al., 1999). The relative abundance of the algal neutral lipid was estimated by fluorescence spectroscopy with the fluorescent dye Nile red (Sigma-Aldrich, St. Louis, MO, USA; Ge et al., 2014).

Total lipid and TAG content analysis of N. oceanica

Total lipids of algal cells were extracted in a chloroform/methanol (1:1, v/v) solvent system (Bligh and Dyer, 1959) and were further separated by TLC on Silica gel plates 60 using hexane/diethyl ether/acetic acid (70:30:1, v/v/v) as developing solvent. TAGs were identified by staining with iodine vapor and recovered with methanol from the plates. Triolein (Sigma-Aldrich) was used as the standard. Total lipids and TAGs were transesterified, and FA methyl esters were analyzed by GC (Ge et al., 2014).

Construction of RNAi strains

For the silencing of NoPDAT, PSS, PEMT, PLMT, and PEAMT in N. oceanica, “long” and “short” fragments were amplified by primers (Supplemental Dataset S7) to obtain a hairpin construct (Wei et al., 2017b). After linearization with Sca I (NoPDAT, PEMT, PLMT, and PEAMT-RNAi vector) or Kpn I (PSS-RNAi vector), the hairpin construct driven by the native β-tubulin promoter was introduced into WT N. oceanica, and transformants were screened (Wei et al., 2017b).

Qualitative and quantitative analysis of TAG and polar lipids

TAGs recovered from TLC plates were dissolved in chloroform/methanol (1:1, v/v), dried and redissolved in isopropanol, and then were analyzed by a Shimadzu LC-20AD HPLC system (Hu et al., 2014).

Total lipid extracts and polar lipid standards (Avanti Polar Lipids) were subjected to TLC in a solvent system consisting of chloroform/methanol/acetic acid/distilled water (75:13:9:3 v/v/v/v). Each band on the plates was scraped off, respectively, and the lipid was extracted with methanol and dried, then redissolved in chloroform/methanol (1:1, v/v). The redissolved single lipids were identified using shotgun MS approach (Wei et al., 2018). Total lipids extracted from WT and NoPDAT KD strains were also analyzed using shotgun MS method directly.

Accession numbers

Sequence data of N. oceanica are from N. oceanica CCMP1779 v1.0 (JGI, https://mycocosm.jgi.doe.gov/Nanoce1779/Nanoce1779.home.html) with the Protein ID showing in Supplemental Dataset S7 if not indicated otherwise. For comparison, the version 2.0 annotation (https://mycocosm.jgi.doe.gov/Nanoce1779_2/Nanoce1779_2.home.html) was also used. The accession numbers used for the sequence alignment and phylogenetic analysis are in Supplemental Dataset S3.

Supplemental data

The following materials are available in the online version of this article.

Supplemental Dataset S1. Contents (nmol/mg dry biomass) of TAG species and the values of fold change shown in the heat map of Figure 6.

Supplemental Dataset S2. The values of fold change in transcript levels of genes encoding components involved in lipid metabolism.

Supplemental Dataset S3. The accession numbers used for the sequence alignment and phylogenetic analysis.

Supplemental Dataset S4. 146 PDAT sequences.

Supplemental Dataset S5. 80 PDAT sequences.

Supplemental Dataset S6. 30 PDAT sequences.

Supplemental Dataset S7. Primers used in this study for RT-qPCR, RNAi expression vector, OE vector, and GFP fusion vector construction.

Supplemental Figure S1. Gene structures of the N. oceanica PDAT annotated in JGI v1 (11464, 1202) and v2 (530913, 640709).

Supplemental Figure S2. Sequence alignment of the amino acids of PDAT.

Supplemental Figure S3. Expression of NoPDAT in S. cerevisiae H1246 analyzed by Western blot.

Supplemental Figure S4. Co-localization of ER and NoPDAT in N. oceanica.

Supplemental Figure S5. The original Western blot image of Figure 4B.

Supplemental Figure S6. Content of MGDG, DGDG, and PC species in WT and the NoPDAT KD line (PDAT-KD4) of N. oceanica.

Supplemental Figure S7. Effect of NoPDAT KD on DGTS of N. oceanica as determined by TLC analysis (the abundance of WT on Day 3 was set as 1).

Supplemental Figure S8. LU-PE accumulation and PDAT expression under different culture conditions.

Supplemental Figure S9. Nitrate concentration in the culture medium of WT and NoPDAT KD lines (PDAT-KD2 and PDAT-KD4) of N. oceanica under different CO₂ availability conditions.

Supplemental Figure S10. Growth curves of WT and NoPDAT KD lines (PDAT-KD2 and PDAT-KD4) of N. oceanica aerated with air, 2% CO₂ or 5% CO₂.

Supplemental Figure S11. Relative mRNA levels of the corresponding gene in CPT OE (CPT-OE1∼6), EPT OE (EPT-OE1∼5), PEMT KD (PEMT-KD2, 4–8), PLMT KD (PLMT-KD1∼6), PEAMT KD (PEAMT-KD3, 5), PSS KD (PSS-KD8, 11), and PSS OE (PSS-OE14, 15) lines in N. oceanica.

Supplemental Figure S12. Effect of CPT OE (CPT-OE1∼6), EPT OE (EPT-OE1∼5), PEMT KD (PEMT-KD2, 4–8), PLMT KD (PLMT-KD1∼6), PEAMT KD (PEAMT-KD3, 5), PSS KD (PSS-KD8, 11), and PSS OE (PSS-OE14, 15) on LU-PE accumulation in N. oceanica.

Supplemental Figure S13. In vitro assay of NoPDAT activity on PE, PC, and PG with various acyl compositions.

Supplemental Figure S14. Cell diameters of WT, NoPDAT KD lines (PDAT-KD2 and PDAT-KD4), and NoPDAT OE lines (PDAT-OE3 and PDAT-OE5) of N. oceanica under different CO₂ availability conditions.

Supplemental Figure S15. The tree with the highest log likelihood (-23816,03).

Supplemental Figure S16. The evolutionary history was inferred by using the ML method and Le_Gascuel_2008 model.

Supplemental Methods S1. Details associated with PDAT identification, phylogenetic analyses, polar lipid and DAG species, plasmid construction, GFP fluorescence analysis, RT-qPCR, and immunooblotting analysis.

Funding

This work was supported by the National Natural Science Foundation of China (41976119 and 91751117) and Strategic Priority Research Program of the Chinese Academy of Sciences (No. XDPB18). A.A., E.M., H.H., and Y.G. were supported by a CEA-CAS bilateral program. J.L. was supported by the National Key R&D Program of China (2018YFA0902500). A.A. and E.M. were supported by the French National Research Agency (GRAL Labex ANR-10-LABEX-04, and EUR CBS ANR-17-EURE-0003).

Conflict of interest statement. None declared.

 

H.H. conceived and designed the research. J.Y. and Y.P. performed the in vivo experiments. J.L. designed the in vitro experiments in yeast and performed them with M.L. and Y.P. A.A. contributed the phylogenetic analyses. H.H., J.L., J.Y., Y.P., and Y.G. analyzed the data. J.L. and H.H. drafted the manuscript. E.M., Y.L., J.L., A.A., and H.H. made critical revision. H.H. agrees to serve as the author responsible for contact and ensures communication.

The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (https://academic.oup.com/plphys/pages/general-instructions) is: Hanhua Hu (hanhuahu@ihb.ac.cn).