Betaine Lipid Is Crucial for Adapting to Low Temperatureand Phosphate Deficiency in Nannochloropsis1[OPEN]

Hiroki Murakami, Takashi Nobusawa, Koichi Hori, Mie Shimojima, and Hiroyuki Ohta2

School of Life Science and Technology, Tokyo Institute of Technology, Yokohama 226-8501, Japan

ORCID IDs: 0000-0002-0530-181X (K.H.); 0000-0002-1068-3242 (H.O.).

Diacylglyceryl-N,N,N-trimethylhomo-Ser (DGTS) is a nonphosphorous, polar glycerolipid that is regarded as analogous to thephosphatidylcholine in bacteria, fungi, algae, and basal land plants. In some species of algae, including the stramenopilemicroalga Nannochloropsis oceanica, DGTS contains an abundance of eicosapentaenoic acid (EPA), which is relatively scarce inphosphatidylcholine, implying that DGTS has a unique physiological role. In this study, we addressed the role of DGTS inN. oceanica. We identified two DGTS biosynthetic enzymes that have distinct domain configurations compared to previouslyidentified DGTS synthases. Mutants lacking DGTS showed growth retardation under phosphate starvation, demonstrating apivotal role for DGTS in the adaptation to this condition. Under normal conditions, DGTS deficiency led to an increase in therelative amount of monogalactosyldiacylglycerol, a major plastid membrane lipid with high EPA content, whereas excessiveproduction of DGTS induced by gene overexpression led to a decrease in monogalactosyldiacylglycerol. Meanwhile, lipidanalysis of partial phospholipid-deficient mutants revealed a role for phosphatidylcholine and phosphatidylethanolamine inEPA biosynthesis. These results suggest that DGTS and monogalactosyldiacylglycerol may constitute the two major pools ofEPA in extraplastidic and plastidic membranes, partially competing to acquire EPA or its precursors derived fromphospholipids. The mutant lacking DGTS also displayed impaired growth and a lower proportion of EPA in extraplastidiccompartments at low temperatures. Our results indicate that DGTS is involved in the adaptation to low temperatures through amechanism that is distinct from the DGTS-dependent adaptation to phosphate starvation in N. oceanica.

Betaine lipids (BLs) are a family of glycerolipidscharacterized by a nonphosphorous, zwitterionic,polar head group and an ether bond connecting thehead group with a diacylglycerol (DAG) backbone.To date, three types of BL have been identifiedin various organisms: diacylglyceryl-N,N,N-trime-thylhomo-Ser (DGTS), diacylglyceryl-hydroxymethyl-N,N,N-trimethyl-b-Ala, and diacylglyceryl-carboxyhydrox-ymethylcholine (Dembitsky, 1996; Cañavate et al., 2016).Although information about the occurrence of the lattertwo BLs is relatively scarce and has only been described inseveral taxonomic groups of algae, DGTS is widely dis-tributed across multiple kingdoms such as Bacteria, Pro-tozoa, Chromista, Fungi, and Plantae (Sato and Furuya,1985; Kato et al., 1996; Künzler and Eichenberger, 1997;

Rozentsvet et al., 2000; Shemi et al., 2016; Cañavate et al.,2016, 2017). The amount of BL in cellular membranes canvary both within a species and among species. In general,limited to no BLs are produced by various species underoptimal culture conditions, whereas BLs accumulate inresponse to phosphate (Pi) starvation (Benning et al., 1995;Geiger et al., 1999; Khozin-Goldberg and Cohen, 2006;Van Mooy et al., 2009; Geske et al., 2013; Riekhof et al.,2014; Abida et al., 2015; Senik et al., 2015; Shemi et al.,2016). Because Pi starvation also triggers phospholipiddegradation, it has been proposed that the increase in BLsmay complement the reduction in phospholipids, therebyreallocating Pi use frommembrane lipid synthesis to othermetabolic pathways. One remarkable case involves thegreen alga Chlamydomonas reinhardtii, which intrinsicallylacks phosphatidylcholine (PC), a major phospholipidclass in most eukaryotes, and instead produces a sub-stantial amount of DGTS regardless of Pi concentration(Giroud et al., 1988). In organisms likeC. reinhardtii, DGTSseems to replace PC constitutively rather than faculta-tively. The complementary association between BLs andphospholipids has long been accepted as an adaptivestrategy to cope with periodic Pi deficiency in the naturalenvironment. In certain algae species, however, BLs seemto have a distinct role aside from substituting for phos-pholipids since the fatty acid profile of BLs is dissimilar tothat of any of the phospholipids, since it contains higherunsaturated groups (Iwai et al., 2015; Cañavate et al., 2016;Dolch et al., 2017; Nobusawa et al., 2017). Moreover, thedegree of change in cellular BLs and PC contents upon

1 This work was supported by CREST (Core Research forEvolutional Science and Technology) and OPERA (Program on OpenInnovation Platform with Enterprises, Research Institute and Acade-mia) of the Japan Science and Technology Agency.

2 Address correspondence to ohta.h.ab@m.titech.ac.jp.The author responsible for distribution of materials integral to the

findings presented in this article in accordance with the policy de-scribed in the Instructions for Authors (www.plantphysiol.org) is:Hiroyuki Ohta (ohta.h.ab@m.titech.ac.jp).

H.M. and H.O. conceived the original research plans; H.M. per-formed the experiments and in silico analysis; T.N. provided techni-cal assistance to H.M.; H.M., T.N., and H.O. analyzed the data; H.M.wrote the manuscript with contributions from all authors.

[OPEN] Articles can be viewed without a subscription.www.plantphysiol.org/cgi/doi/10.1104/pp.17.01573

Plant Physiology�, May 2018, Vol. 177, pp. 181–193, www.plantphysiol.org � 2018 American Society of Plant Biologists. All Rights Reserved. 181

https://plantphysiol.orgDownloaded on February 12, 2021. - Published by Copyright (c) 2020 American Society of Plant Biologists. All rights reserved.

fluctuations in external Pi availability is quite dif-ferent among species (Cañavate et al., 2017). Thesespecies-dependent differences in BLs fatty acid pro-files and susceptibility to low-Pi stress imply meta-bolic diversity among algae, as well as versatility inBLs function and requirement under Pi-starvedconditions, yet we lack a comprehensive under-standing of BLs functions (Cañavate et al., 2016).

DGTS is the sole BLs for which the biosyntheticpathway, precursors, and enzymes have been iden-tified. DGTS biosynthesis is achieved through twomain steps: attachment of a C4 homo-Ser moiety to aDAG backbone followed by the consecutive additionof three methyl groups to its amino group (Mooreet al., 2001; Hofmann and Eichenberger, 1996; Fig.1A). In prokaryotes, these first- and second-step re-actions are catalyzed by BtaA and BtaB, respectively,whereas in eukaryotes, betaine lipid synthase1 (BTA1), which contains both BtaA- and BtaB-likedomains, carries out the entire process of DGTS bio-synthesis (Klug and Benning, 2001; Riekhof et al.,2005; Fig. 1, A and B). The gene encoding BTA1 wasinitially identified in C. reinhardtii, and subsequentwork identified putative BTA1 orthologs in severalDGTS-producing fungi (Riekhof et al., 2005, 2014;Senik et al., 2015).

Nannochloropsis is a genus of microalgae that be-longs to the stramenopiles (also referred to as hetero-konta) and possesses a secondary plastid derived froma red alga. Nannochloropsis species are capable of ac-cumulating a large amounts of neutral lipids (i.e. tri-acylglycerol) and eicosapentaenoic acid (or 20:5,number of carbons:number of double bonds) andthus have receivedmuch interest as a biological sourcefor these compounds (Mühlroth et al., 2013; Slocombe

et al., 2015; Ma et al., 2016). These Nannochloropsisspecies traits could be applied to the production ofspecialty fatty acids that are not synthesized by nativestrains but are valuable for industrial or pharmaceu-tical use. To exploit the ability of Nannochloropsis spe-cies to generate specialty lipids, a great deal of efforthas been devoted toward omics studies such as ge-nome sequencing, as well as transcriptomic and lip-idomic analyses (Radakovits et al., 2012; Vieler et al.,2012; Wang et al., 2014; Li et al., 2014; Poliner et al.,2015; Alboresi et al., 2016). Moreover, the develop-ment of molecular biology tools that enable the gen-eration of Nannochloropsis transformants is increasing(Kilian et al., 2011; Wang et al., 2016; Wei et al., 2017).The coordination of omics data and functional analy-ses of key genes related to lipid biosynthesis wouldyield important information on algal lipid metabolismthat is expected to form the basis for future metabolicengineering techniques (Dolch et al., 2017; Nobusawaet al., 2017; Zienkiewicz et al., 2017).

Our previous study revealed that Pi starvation in-creases the DGTS content of the Nannochloropsis speciesNannochloropsis oceanica (strain NIES-2145; Iwai et al.,2015). Additionally, several recent reports demonstratedthat DGTS is highly enriched in 20:5 fatty acids, whichare minor components of PC in Nannochloropsis (Iwaiet al., 2015; Cañavate et al., 2016; Dolch et al., 2017;Nobusawa et al., 2017). To investigate the physiologicalrole of DGTS in N. oceanica, we characterized two DGTSbiosynthetic enzymes and generated DGTS-lacking mu-tants, DGTS-overproducing lines, and phospholipid-reducing mutants. Our results suggest that DGTS is re-quired not only for normal cell proliferation underPi-starved conditions, but also for adaptation to lowtemperatures.

Figure 1. Biosynthetic pathway for DGTS. A,DGTS is synthesized from DAG via two step re-actions. DGHS, Diacylglycerylhomo-Ser. B, Bio-synthetic enzymes for DGTS with their catalyticdomains. RsBtaA and the C-terminal domain ofCrBTA1 catalyze the first step, and RsBtaB and theN-terminal domain of CrBTA1 catalyze the sec-ond step. Domains of 3-amino-3-carboxypropyl-transferase and N-methyltransferase predictedwith Pfam 30.0 are indicated by red and blueboxes, respectively. The start and end point of thedomains are indicated above each construct, andthe total number of amino acid residues (AA) isshown to the right. Rs, Rhodoobacter sphaer-oides; Cr, Chlamydomonas reinhardtii.

182 Plant Physiol. Vol. 177, 2018

Murakami et al.

https://plantphysiol.orgDownloaded on February 12, 2021. - Published by Copyright (c) 2020 American Society of Plant Biologists. All rights reserved.

RESULTS

BTA1 Homologs in N. oceanica, Other Algae, andLand Plants

First, we searched our RNA sequencing data set(Nobusawa et al., 2017) for DGTS biosynthetic en-zymes in N. oceanica strain NIES-2145. Two proteinshad high amino acid sequence similarity to C. rein-hardtii BTA1 (Riekhof et al., 2005) and were designatedBTA1L (764 amino acid residues) and BTA1S (261 res-idues; Fig. 1B). The domain configurations of theseproteins were distinct from that of CrBTA1. BTA1Lhas a reverse domain order, with a BtaA-like domainat the N terminus and a BtaB-like domain at theC terminus, and BTA1S contains only a domain ho-mologous to the N terminus of CrBTA1 and thus issimilar to the bacterial BtaB (Fig. 1B). Previous workidentified binding sites for the substrate S-adenosyl-Met in both the BtaA-like and BtaB-like domains ofCrBTA1 and shows that these binding sites are nec-essary for enzyme activity (Riekhof et al., 2005). Wetherefore investigated whether the Val-Asp motif inthe substrate binding site is present in N. oceanicaBTA1L and BTA1S and found that BTA1L has thismotif in both the N- and C-terminal domains, whereasthe Val in the motif is replaced with Leu in BTA1S(Supplemental Fig. S1, A and B). Given that BtaB fromRhodoobacter sphaeroides, which can synthesize DGTStogether with BtaA during Pi deficiency, encodes Leuat this locus (Supplemental Fig. S1B), BTA1S may alsoplay a role in DGTS biosynthesis. Thus, N. oceanicaharbors two proteins, one of which has a complete setof putative catalytic domains that could possibly me-diate DGTS biosynthesis, whereas the other only has aBtaB-like N-methyltransferase domain (Fig. 1B).Having observed a domain-reversed type of BTA1

in N. oceanica, we sought BTA1L homologs in otheralgae and land plants using available databases. Wefound BTA1L-like proteins in prasinophyte green algae,chlorarachniophyte algae, and several species ofChromista and Alveolata; the latter two are cladespossessing a red alga-derived secondary plastid(Supplemental Fig. S1C). On the other hand, putativeorthologs of CrBTA1 were detected in viridiplantae (aclade including green algae and land plants) except forprasinophyceae and seed plants (Supplemental Fig.S1C). Phylogenetic analysis of the BTA1L-like andCrBTA1-like proteins revealed that these algal andplant putative BTA1 proteins could be separated intotwo subgroups (types A and B) according to their do-main arrangements (Supplemental Fig. S2). In addition,we found BTA1S-like proteins in some species ofstreptophyta, such as Klebsormidium nitens (formerlyidentified as Klebsormidium flaccidum; Hori et al., 2014;Ohtaka et al., 2017), Marchantia polymorpha, Physcomi-trella patens, and Selaginella moellendorffii, all of whichhave type A BTA1, but not in the majority of the stra-menopile algae, which have type B BTA1 proteins(Supplemental Fig. S1C).

Pi Starvation Upregulates BTA1L and BTA1S Expressionand Enhances DGTS Accumulation

Our previous study revealed that the DGTS level in-creases in N. oceanica in response to Pi starvation (Iwaiet al., 2015). Under Pi-starved conditions in Gram-negative bacteria and fungi, DGTS biosynthesis istightly controlled by a transcription factor (Geiger et al.,1999; Riekhof et al., 2014). To determine whether Pistarvation-induced DGTS accumulation in N. oceanica isalso transcriptionally regulated, we quantified expres-sion of BTA1L and BTA1S in N. oceanica grown underPi-depleted, Pi-replete, or other stress conditions (nitrogenstarvation and low temperature). First, we confirmed thatthe Pi depletion conditions indeed induced DGTS accu-mulation (Fig. 2A; Supplemental Fig. S3). As we describebelow in detail, cold treatment also increased the relativeproportion of DGTS, albeit to a lower extent than Pistarvation (Fig. 2A). Reverse transcription quantitativePCR (RT-qPCR) revealed that BTA1L and BTA1S weresignificantly upregulated by a factor of 2 during Pi star-vation compared with normal Pi-replete conditions (Fig.2, B andC). This result indicated that the increase inDGTSlevel observed in Pi-starved N. oceanica is also transcrip-tionally regulated. Because we could not identify an ap-parent ortholog of Pho4p, a known transcriptionalactivator of BTA1 in fungi, the factor(s) responsible forinduction of these genes remains unknown.

Figure 2. Changes in membrane lipid composition relative expressionlevel of BTA1L and BTA1S genes under stress. A, Membrane lipidcomposition of N. oceanica wild-type strain grown under normal,nitrogen-starved, Pi-starved, or 15°C conditions for 3 d. B andC, Relativeexpression of BTA1L (B) and BTA1S (C) under the four conditions. Ex-pression was normalized to that of the gene encoding NADH dehydro-genase subunit 11. Data represent the means 6 SD of three biologicallyindependent experiments. PG, Phosphatidylglycerol; PI, phosphatidyl-inositol; SQDG, sulfoquinovosyldiacylglycerol.

Plant Physiol. Vol. 177, 2018 183

Dual Roles for a Betaine Lipid in Nannochloropsis

https://plantphysiol.orgDownloaded on February 12, 2021. - Published by Copyright (c) 2020 American Society of Plant Biologists. All rights reserved.

BTA1L and BTA1S Are DGTS Synthases

To obtain evidence that BTA1L and BTA1S arebona fide DGTS biosynthetic enzymes, we generatedN. oceanica knockout mutants of BTA1L and BTA1S byhomologous recombination and assessed their capacityto synthesize DGTS (Supplemental Fig. S4). One-dimensional thin-layer chromatography (TLC) wasused for isolating DGTS from the total lipid fraction.With our TLC solvent system,DGTS separated into threespots probably because of differences in the content of20:5 fatty acids (Supplemental Fig. S5). These three spotswere absent in the total lipid fraction of the bta1l mu-tants, demonstrating that BTA1L is essential for DGTSbiosynthesis (Figure 3). Conversely, the bta1s mutantsdid not show any defect in DGTS content even whengrown under Pi-deficient conditions, which has the ef-fect of up-regulating BTA1S expression and cellularDGTS levels (Supplemental Fig. S6). This result sug-gested that BTA1S either has no relevance to DGTS bi-osynthesis or has a similar function to that of theC-terminal domain of BTA1L, which compensates forthe absence of BTA1S. To determine whether BTA1Sis actually involved in DGTS biosynthesis in vivo,we constructed a mutant lacking only the C-terminaldomain of BTA1L by incorporating a C-terminaldomain-truncated BTA1L (BTA1LDC) into bta1l andsubsequently examining the ability of the complemented

mutant to synthesize DGTS (Supplemental Fig. S7).Because we could not knockout BTA1S from bta1l;BTA1LDC, BTA1LDC was also introduced into thebta1s;bta1l double mutant as a negative control. bta1s;bta1l was generated by BTA1L knockout of the bta1smutant background. As a positive control, the bta1s;bta1l mutant was complemented with full-lengthBTA1L. As shown in Figure 3, DGTS was detected inbta1l;BTA1LDC and bta1s;bta1l;BTA1L but not in bta1s;bta1l and bta1s;bta1l;BTA1LDC. To exclude the possi-bility that inadequate induction of BTA1LDC in bta1s;bta1l;BTA1LDC caused the lack of DGTS production,we compared the gene expression level of trans-formed BTA1LDC or BTA1L between bta1l;BTA1LDC,bta1s;bta1l;BTA1LDC, and bta1s;bta1l;BTA1L. Therewas no apparent reduction in the BTA1LDC transcriptlevel in bta1s;bta1l;BTA1LDC (Supplemental Fig. S8).These data indicated that either BTA1S or theC-terminal domain of BTA1L is necessary for DGTSbiosynthesis; thus, BTA1S can function in DGTS syn-thesis in N. oceanica.

Under Pi Starvation, Proliferation of bta1l Mutants IsRetarded and the MGDG Level Is Elevated

Next, we cultured the bta1l, bta1s, and bta1s;bta1lmutants in either Pi-sufficient or Pi-deficient mediumto examine whether the lack of DGTS affects cell pro-liferation and the membrane lipid profile. As shown inFigure 4, A and B, the proliferation of bta1l cells wascomparable to that of the empty-vector (EV) controlstrain during exponential growth under Pi-sufficientconditions, whereas proliferation was impaired underPi-deficient conditions. This result demonstrated theimportance of DGTS for proper growth under Pistarvation. Unexpectedly, the cell density of bta1s;bta1lwas reduced under normal conditions (Fig. 4A). Wethen analyzed the membrane lipid composition ofbta1l and bta1s;bta1l grown under Pi-replete condi-tions. DGTS was not detected in bta1l or bta1s;bta1l(Fig. 4C), as was shown in Figure 3. Mutant bta1lexhibited an increase in the relative proportion ofmonogalactosyldiacylglycerol (MGDG; a major lipidof the chloroplast membrane) and phosphatidyletha-nolamine (PE), whereas bta1s;bta1l had an increasedproportion of digalactosyldiacylglycerol (DGDG), PC,and PE compared with EV (Fig. 4C). Fatty acid com-position of each lipid class was also altered in thesemutants (Supplemental Fig. S9): in bta1s;bta1l, theproportion of 20:5 decreased in MGDG but increasedin DGDG (Supplemental Fig. S10, A and B). Both bta1land bta1s;bta1l had more 18:1, 18:2, and 20:4 but less16:1 and 16:2 in PC, whereas they had more 20:4 butless 16:0, 16:1, 16:2, and 18:2 in PE (SupplementalFig. S9, E and F). These observations indicated thatdefects in the DGTS biosynthetic pathway affected theprofiles of both plastidic and extraplastidic membranelipids, even under conditions that did not impair cellproliferation.

Figure 3. DGTS synthesis in bta1l and the BTA1L- or BTA1LDC-com-plemented mutant as assessed with TLC. Total lipid of EV, bta1l, bta1l;BTA1LDC, bta1s;bta1l, bta1s;bta1l;BTA1LDC, and bta1s;bta1l;BTA1Lcultivated under Pi-starved conditions for 4 d were separated by one-dimensional TLC using solvent system chloroform/methanol/aceticacid/water (170:30:15:3, v/v/v/v). The asterisk denotes spots of DGTS.

184 Plant Physiol. Vol. 177, 2018

Murakami et al.

https://plantphysiol.orgDownloaded on February 12, 2021. - Published by Copyright (c) 2020 American Society of Plant Biologists. All rights reserved.

Overexpression of BTA1L Leads to an Increase in DGTSand a Decrease in MGDG

We overexpressed BTA1L in N. oceanica to over-produce DGTS under normal culture conditions andanalyzed the effect this had on membrane lipid

composition (Supplemental Fig. S10). BTA1L wasexpressed under the control of the violaxanthin-chlorophyll a binding protein 1 (VCP1) promoter, whichfacilitates potent expression under normal growthconditions (Kilian et al., 2011). We introduced the geneconstruct into N. oceanica and obtained an over-expression line (OEBTA1L) that had ;10-fold higherexpression than the control strain (Supplemental Fig.S10). Consistent with this, the proportion of DGTS inthe polar lipid fraction increased by;40% inOEBTA1Lcompared with EV (Fig. 5A). Conversely, OEBTA1Lhad a smaller proportion of MGDG and phosphati-dylglycerol but exhibited no significant change in theproportion of PC (Fig. 4A; Klug and Benning, 2001;Riekhof et al., 2014). The distribution of 20:5 within themajor classes of membrane lipid was also altered inOEBTA1L: this ratio was higher in DGTS but lower inMGDG (Fig. 5E). Because DGTS is enriched in 20:5(Supplemental Fig. S9H), we initially expected that theoverproduction of DGTS led to an overall increase in20:5 content. However, the total 20:5 content was notgreater in OEBTA1L (EV, 19.46 1.78 mg L–1; OEBTA1L,20.1 6 0.979 mg L–1; n = 3), probably owing to the re-duction in the relative amount of 20:5 in DGTS (Fig. 5B).Thus, although DGTS has an abundance of 20:5, en-hancing DGTS biosynthesis alone may not be useful forincreasing the 20:5 content in N. oceanica.

The Proportion of 20:5 in Galactolipids Is SignificantlyReduced in cct1 and pect1 Mutants

In Nannochloropsis, de novo fatty acid synthesis isconsidered to occur inside chloroplasts, whereas bio-synthetic processes of 20:5, namely, desaturation andelongation of long-chain fatty acids, proceed at the en-doplasmic reticulum (ER) membrane (Guschina andHarwood, 2006; Dolch et al., 2017). ER-localized PC andPEhave been proposed to contribute to 20:5 biosynthesisbecause these two phospholipids harbor mainly desa-turated fatty acids of lengths C18 (PC) and C20 (PE;Schneider and Roessler, 1994; Kaye et al., 2015). To gainfurther insight into the metabolic function of membranelipids, we aimed to generate mutants that lack either PCor PE. To this end, we targeted CTP:phosphocholinecytidylyltransferase 1 (CCT1) and CTP:phosphoethano-lamine cytidylyltransferase 1 (PECT1), which catalyzethe respective rate-limiting step for PC and PE biosyn-thesis (Inatsugi et al., 2009; Mizoi et al., 2006). For N.oceanica sp NIES-2145, CCT1 and PECT1 were eachpredicted to be encoded by single genes based on a se-quence similarity search, and a knockout strain wasproduced for each gene (cct1 and pect1; SupplementalFig. S11). As shown in Figure 6A, under normal cultureconditions, we observed a 44% reduction of PC in cct1and a 43% reduction of PE in pect1, suggesting redun-dant function for these enzymes that precluded completedepletion of PC or PE in each single mutant. Under thesame conditions, the cct1 mutant had a higher propor-tion of DGTS (+24%) and phosphatidylglycerol (+14%)

Figure 4. Phenotypes of mutants bta1l, bta1s, and bta1s;bta1l. A and B,Growth curve for EV, bta1l, bta1s, and bta1s;bta1l under Pi-sufficient (A)and Pi-deficient (B) conditions. Each cell culture was diluted with me-dium to an initial density of 107cells mL–1. Data represent themean6 SD

of four (A) and three (B) biologically independent experiments. C,Comparison of relative polar lipid composition. The EV line (control)and mutant lines were cultivated under normal conditions for 3 d andthen harvested for lipid analysis. Data represent the mean6 SD of threebiologically independent experiments. Statistical significance was de-termined with Dunnett’s test (compared with EV): #, P , 0.1;*, P , 0.05; **, P , 0.01. SQDG, Sulfoquinovosyldiacylglycerol; PG,phosphatidylglycerol; PI, phosphatidylinositol; N.D., not detected.

Plant Physiol. Vol. 177, 2018 185

Dual Roles for a Betaine Lipid in Nannochloropsis

https://plantphysiol.orgDownloaded on February 12, 2021. - Published by Copyright (c) 2020 American Society of Plant Biologists. All rights reserved.

compared with the EV control, whereas the membranelipid profile was not significantly altered in pect1 (Fig.6A). Moreover, cct1 had smaller relative amount of 14:0,16:1, and 20:5 and a greater amount of 20:4 in PC,whereas pect1 had a significantly smaller proportion of16:1 in PE (Fig. 6, B and C). In DGTS, cct1 had a greaterproportion of 14:0, 16:0, 18:2, and 20:4 at the expense of20:5 (Fig. 6F). Importantly, the proportion of 20:5 inMGDG and DGDG was reduced by ; 10% in bothcct1 and pect1, indicating the importance of PC andPE in producing the 20:5 that is incorporated into thechloroplast membrane (Fig. 6, B and C).

Impaired Growth of bta1l at Low Temperatures

Growth at low temperatures has been shown to fa-cilitate the accumulation of 20:5 in Nannochloropsis (Huand Gao, 2006; Hoffmann et al., 2010), yet how polarlipid composition changes has not been described in

detail. We therefore cultured the N. oceanica wild-typestrain at 15°C and assessed its lipid composition. Theproportions of DGDG, phosphatidylglycerol, andDGTS increased by up to 42, 17, and 66%, respectively,at the expense of MGDG (Fig. 2A, Supplemental Fig.S3). Unlike Pi starvation, the low temperature causedan increase in DGTS content without stimulating theexpression of BTA1L and BTA1S (Fig. 2, B and C). Next,the bta1l, cct1, and pect1mutants were exposed to a lowtemperature (15°C) to clarify which lipid is importantfor adaptation to low temperature and how membranelipid composition changes when either of DGTS, PC, orPE is deficient. The results revealed that only bta1l had asignificantly lower cell density (Fig. 7A). We then ana-lyzed the lipid composition of these mutants and foundthat bta1l had noDGTS but had an increased proportionof MGDG, PC, and PE, compared with the EV control(Fig. 7B). With regards to fatty acid composition of eachlipid class in bta1l, we observed a marked increase of

Figure 5. Membrane lipid analysis of theBTA1L overexpression line. A, Comparisonof membrane lipid composition between EVandOEBTA1L. B to D, Fatty acid profiles forDGTS (B), MGDG (C), and DGDG (D). Datarepresent the mean 6 SD of three biologi-cally independent experiments. Statisticalsignificance was determined with Tukey’stest and is indicated by letters at the top. E,Distribution of 20:5 within the eight classesof membrane lipid after 2 d of culture. Datarepresent the mean 6 SD of three biologi-cally independent experiments. Statisticalsignificance was determined with the two-tailed Student’s t test: *, P, 0.05; **, P, 0.01.SQDG, Sulfoquinovosyldiacylglycerol; PG,phosphatidylglycerol; PI, phosphatidylinositol.

186 Plant Physiol. Vol. 177, 2018

Murakami et al.

https://plantphysiol.orgDownloaded on February 12, 2021. - Published by Copyright (c) 2020 American Society of Plant Biologists. All rights reserved.

20:5 in PC and PE (Fig. 7, C and D). Nevertheless, theproportion of 20:5 in the bta1l extraplastidic membraneswas significantly smaller comparedwith that measuredfor EV (Fig. 7F). We excluded phosphatidylglycerolfrom this evaluation because it is known to exist in bothplastidic and extraplastidic compartments (Babiychuket al., 2003). These bta1l phenotypes indicated thatDGTS is required for maintaining both optimal growth

and a large amount of 20:5 in extraplastidic membranesat low temperatures. The cct1 mutant produced lessthan half of the PC and had a larger proportion ofDGTS, sulfoquinovosyldiacylglycerol, and PE than theEV control (Fig. 7B). Unexpectedly, the amount of PE inpect1was comparable to that in the EV control (Fig. 7B),possibly owing to activation of CCT1 at low tempera-ture, as reported in Arabidopsis (Arabidopsis thaliana;

Figure 6. Membrane lipid composition ofmutants cct1 and pect1 grown under normalconditions. A, Membrane lipid compositioncompared with EV (control). EV and mutantlines were cultured for 3 d under normalconditions before lipid analysis. B to F, Fattyacid profile of PC (B), PE (C), MGDG (D),DGDG (E), and DGTS (F) in EV, cct1, andpect1. Data represent the mean6 SD of fourbiologically independent experiments. Sta-tistical significance was determined withDunnett’s test (compared with EV): #, P ,0.1; *, P , 0.05; **, P , 0.01; ***, P ,0.001. SQDG, Sulfoquinovosyldiacylgly-cerol; PG, phosphatidylglycerol; PI, phos-phatidylinositol.

Plant Physiol. Vol. 177, 2018 187

Dual Roles for a Betaine Lipid in Nannochloropsis

https://plantphysiol.orgDownloaded on February 12, 2021. - Published by Copyright (c) 2020 American Society of Plant Biologists. All rights reserved.

Inatsugi et al., 2009). Consistent with this result, pect1did not show any significant difference in the propor-tion of other membrane lipid or fatty acid profiles ofeach lipid class compared with EV (Fig. 7, B–E).

DISCUSSION

Although genes for DGTS biosynthesis are widelyfound inmany algae and basal land plants (SupplementalFigs. S1 and S2), previous studies on these DGTS-relatedgenes have been limited in some species such asC. reinhardtii and R. sphaeroides (Klug and Benning,2001; Riekhof et al., 2005). In this work, we investi-gated the functions of the DGTS biosynthetic enzymes

and the DGTS product via identification of twoBTA1 homologs in the oleaginous alga N. oceanica.BTA1L has a reversed domain structure comparedwiththat of a previously reported BTA1 (Riekhof et al., 2005,2014; Senik et al., 2015), with putative orthologs inseveral species of secondary algae and prasinophytealgae (Fig. 1B; Supplemental Figs. S1C and S2). TheBTA1L-like proteins form a clade that is phylogeneti-cally distinct from the conventional type BTA1proteins that are mainly distributed in viridiplantae(Supplemental Fig. S2). These observations indicate theexistence of two types of bifunctional BTA1-encodinggenes that have been inherited through different line-ages in algae and plants (types A and B; seeSupplemental Fig. S2). We also discovered that BTA1S

Figure 7. Phenotypes of themutants bta1l, cct1, and pect1 grown at low temperature. A, Number of cells for EVand the indicatedmutants after 3 d of culture. B to E, Comparison of relative polar lipid composition (B) and fatty acid profile of PC (C), PE (D), andDGTS (E) in EV, bta1l, cct1, and pect1. F and G, Molar ratio of 20:5 in extraplastidic (F) and plastidic (G) membrane lipids. Datarepresent the mean 6 SD of three biologically independent experiments. Statistical significance was determined with Dunnett’stest (compared with EV): *, P , 0.05; **, P , 0.01; ***, P , 0.001. SQDG, Sulfoquinovosyldiacylglycerol; PG, phosphatidyl-glycerol; PI, phosphatidylinositol.

188 Plant Physiol. Vol. 177, 2018

Murakami et al.

https://plantphysiol.orgDownloaded on February 12, 2021. - Published by Copyright (c) 2020 American Society of Plant Biologists. All rights reserved.

is a homolog of the bacterial BtaB enzyme, as deter-mined by its involvement in DGTS biosynthesis in N.oceanica (Figs. 1B and 3; Supplemental Fig. S7). BTA1S-like proteins were also found in streptophyta(Supplemental Fig. S1C). Because these distant orga-nisms harbor a common set of genes, the bifunctionalBTA1-encoding gene and the gene encoding a BtaB-likeN-methyltransferase, BTA1S and its homologs mayplay specific roles in DGTS biosynthesis and/or otherunknown processes that could not be accomplished bythe bifunctional BTA1 alone. Consistent with thisspeculation, bta1s;bta1l had retarded growth comparedwith bta1l under normal conditions (Fig. 4A), eventhough DGTS was equally deficient in both mutants(Fig. 3). We initially proposed that BTA1S may be im-plicated in betaine (Gly betaine) synthesis becauseBTA1 is a BL synthase. Supplementation with Gly be-taine, however, could not complement the growth de-ficiency of bta1s;bta1l. The catalytic reaction mediatedby BTA1S may be reminiscent of the biosyntheticpathways of PC, which comprise triple methylation of PEor phosphoethanolamine (Sato et al., 2016; Hirashimaet al., 2017). However, because the PC content was notreduced in bta1s;bta1l (Fig. 4C), it is unlikely that BTA1Scontributes to PC formation. Although this studycould not reveal BTA1S function, future analyses willelucidate the complete functions of NannochloropsisBTA1 homologs.We generated DGTS-lacking mutants to obtain new

insight into the physiological role of DGTS in N. oce-anica. The growth of bta1l was not impaired undernormal conditions, whereas it was strongly arrestedunder Pi starvation and also at low temperatures (Figs.4A, 4B, and 7A). Given that the wild-type strain accu-mulated a larger amount of DGTS in response to Pistarvation and low temperature, DGTS appears to havean essential role in the adaptation to these stresses (Fig.2A). As many previous studies have described, DGTSwould be expected to be crucial for adaptation toPi-starved conditions as an alternative polar lipid tophospholipids (Van Mooy et al., 2009; Riekhof et al.,2014; Cañavate et al., 2017). On the other hand, wepropose two possibilities that explain why DGTS isrequired for optimal cell proliferation at low tempera-tures. First, DGTS may be important as a store for 20:5in extraplastidic membranes. Polyunsaturated fattyacids that are esterified to membrane lipids contributeto the maintenance of membrane fluidity, especially atlow temperature (Murata and Los, 1997). Recently,Zorin et al. (2017) suggested that arachidonic acid(20:4)-containingMGDGmight contribute to sustainingchloroplast membrane fluidity at low temperatures inthe green algae Lobosphaera incisa. Our results indicatethat, at low temperature, DGTS is naturally enriched in20:5 and that DGTS deficiency leads to a smaller pro-portion of 20:5 in extraplastidic compartments com-pared with the EV control (Supplemental Figs. S3 andS9; Fig. 7F). Moreover, no other lipid class located inextraplastidic membranes contains as much 20:5 asDGTS. We also analyzed fatty acid composition of

triacylglycerol in the EV control and the bta1 mutantand found very low levels of 20:5 with no large differ-ence between these two lines. Therefore, we haveomitted the idea that the neutral lipid has a great in-fluence on themetabolism related to 20:5. InN. oceanica,DGTS may have a role in retaining 20:5; therefore, itsrelative deficiency affects the amount of 20:5, and hencethe functionality, of the plasma membrane and otherextraplastidic membranes, leading to reduced prolifer-ation at 15°C. Second, the polar head group of DGTS,which is relatively large and thus may help maintainmembrane-bilayer homeostasis even at low tempera-ture, may be important for adapting to temperaturechanges.

By characterizing compositional changes in mem-brane lipids in mutants lacking DGTS, mutants over-producing DGTS, and phospholipid-reducing mutants,we attempted to elucidate the mechanism of lipidmetabolism in N. oceanica. Lipid analyses in bta1l andOEBTA1L suggested a negative correlation between theamount of DGTS and MGDG (Figs. 4C and 5A). Thistrend was also reported in Chlorella minutissima (Haighet al., 1996): DGTS and MGDG levels fluctuated peri-odically with a reciprocal relationship, leading to thespeculation that DGTS might be a donor of 20:5 forchloroplast MGDG, another major pool of 20:5. Like-wise, a suite of experiments using chemical inhibitorsand low-Pi stress suggested that DGTS provides 20:5for the sn-1 positions of galactolipids in the eustigma-tophyte Monodus subterraneus (Khozin-Goldberg et al.,2002; Khozin-Goldberg and Cohen, 2006). However,our results suggest that DGTS is not crucial for deliv-ering 20:5 from the ER to chloroplast membranes be-cause bta1l did not show a reduction in 20:5 ofgalactolipids, nor galactolipids themselves, and con-versely OEBTA1L had a reduced level of MGDG (Figs.4C and 5A; Supplemental Fig. S9, A and B). DGTS andMGDG may rather compete for the two major 20:5destinations in extraplastidic and plastidic membranes,for which fatty acids are produced via the catabolism ofphospholipids (Fig. 8). This hypothesis would also ex-plain the mechanism by which DGTS levels increasedduring low temperatures compared with normal con-ditions in the wild type; since MGDG levels droppedunder low temperature in the EV, and the DGTS-lacking mutant had more MGDG at low temperaturesthan the EV, the accumulation of DGTS may be causedby the repression ofMGDG synthesis. In this scenario, itis not activation of the DGTS synthetic enzymes butoversupplementation of substrates that would lead toenhanced DGTS levels. Alternatively, because thesubcellular compartment in which DGTS and MGDGexists has yet to be defined in secondary symbiotic al-gae, it may be that DGTS and MGDG localize to thesame place, e.g. the chloroplast envelope membrane,and act complementarily to each other, as do DGTS andPC in extraplastidic membranes. Notably, in second-ary symbiotic algae, the chloroplast is enclosed bythree or four envelopes, and the outermost envelope iscontinuous with the ER membrane (Murakami and

Plant Physiol. Vol. 177, 2018 189

Dual Roles for a Betaine Lipid in Nannochloropsis

https://plantphysiol.orgDownloaded on February 12, 2021. - Published by Copyright (c) 2020 American Society of Plant Biologists. All rights reserved.

Hashimoto, 2009). This fact may underlie intermixedmembrane lipid composition in the chloroplast enve-lope derived from these two organelles, and a moresophisticated fractionation method must be developedto unequivocally determine the membrane lipid profileat the subcellular level.

In the case of secondary algae including N. oceanica,their genomes generally lack any TRIGALACTOSYLDIA-CYLGLYCEROL1-5 homologs, whichmainlymediate theeukaryotic pathway in Arabidopsis (Li et al., 2016; Horiet al., 2016). This means that the principle for the DAGsupplying pathway to galactolipids synthesis in higherplants would not be directly applied to their metabolism.However, elongation of 16:0 and desaturation of theelongated very-long-chain fatty acids, specifically forma-tion of 20:5, mainly occurs in the ER not in plastids inNannochloropsis (Schneider andRoessler, 1994; Dolch et al.,2017), suggesting that extensive lipid transport from theER to the plastid must take place. This lipid transfer routeis discerned from the eukaryotic pathway and is rathertermed the omega pathway, while its detail is unresolved(Petroutsos et al., 2014). Our phenotypic analyses of cct1and pect1 suggested that 20:5 are mainly producedthrough PC and PE and then transferred to plastid gal-actolipids, without being mediated by DGTS, and toDGTS and other lipids.We observed a lower occupancy of20:5 in the galactolipids of cct1 and pect1 under normalconditions (Fig. 6, D and E). In cct1, DGTS also had a de-creased proportion of 20:5, although this may be partlyattributable to the relative increase in 20:5-free DGTS(Fig. 6F). These results suggest that PC and PE are im-portant for producing the 20:5 used for the formation ofthe chloroplast membrane, and these roles cannot be fullycompensated by excess DGTS. Because there was only amoderate reduction in 20:5 in cct1 and pect1 under normalconditions, the possibility remains that DGTS partly sup-ports the metabolic roles of these phospholipids. We notethat, at low temperatures, cct1 did not have a decreasedlevel of plastidic 20:5 even though the amount of PC wasless than half (Fig. 7B). This result suggests the possibility

that DGTS metabolically compensated for the PC defi-ciency or that even a small amount of PCwas sufficient forsynthesizing a comparable amount of 20:5 in thewild-typestrain, at least under these conditions.

Despite the widespread distribution of BLs amongphytoplankton, understanding their defined roles incellular physiology has been limited mostly to thecondition of Pi starvation. This study reveals two dis-tinct roles for DGTS inN. oceanica, namely, that DGTS isnot only important as a surrogate lipid of phospho-lipids during Pi starvation but also required for adap-tation to low temperatures. Furthermore, our resultsprovide information about the mechanism of mem-brane lipid metabolism in N. oceanica. Although themicroalgal lineage exhibits vast metabolic diversity, wespeculate that the characteristics of DGTS may be ap-plicable to other organisms, especially species that aretaxonomically close to Nannochloropsis or algae thatinhabit low Pi and/or low-temperature environments.Future detailed lipid analyses of algae and plants willhelp to clarify the diversity of roles played by BLs inalgal homeostasis.

MATERIALS AND METHODS

Nannochloropsis oceanica Strains and Culture Conditions

The N. oceanica strain NIES-2145 was obtained from the National Institutefor Environmental Studies. For the “normal” condition, cells were cultured in a50-mL volume at 25°C in F2Nmedium (Kilian et al., 2011) with 36 g L–1 artificialseawater (Wako Pure Chemical Industries) in a NEG test tube (Nichiden-RikaGlass) with continuous bubbling of 2% CO2 and constant light (40 mmol pho-tons m–2 s–1). Before each experiment, the cells were precultured for 3 d undernormal conditions starting from a density of 107 cells mL–1. Cell density wasmeasured using a bacteria counter (Sunlead Glass) with an optical microscope.To induce nitrogen or Pi starvation, cells were washed twice with nitrogen- orphosphorus-free F2N medium (cells were collected at 1,500g for 7 min andresuspended in the respective medium) as described by Iwai et al. (2015). Agrowth chamber (LH-80LED-DT; Nippon Medical & Chemical Instruments)was used for low-temperature experiments.

Phylogenetic Analysis

Amino acid sequences of putative BTA1 homologs from algae and plantswere retrieved from available databases and were aligned usingMAFFT v7.245(Katoh and Standley, 2013). Regions of low sequence conservation were elim-inated with the Gblocks server version 0.91b (Talavera and Castresana, 2007).The resultant alignment was subjected to phylogenetic analysis with MEGA6.06 (Tamura et al., 2013) using the maximum-likelihood method with theamino acid substitution model of LG model + Gamma (eight categories).

Plasmid Construction and Nuclear Transformation

The NT7 gene cassette (Kilian et al., 2011), which consists of the VCP2promoter (ProVCP2), the Streptoalloteichus hindustanus bleomycin resistancegene (Sh ble), and the VCP1 terminator (terVCP1), was constructed between thePstI and KpnI sites of the pUC18 vector (reverse complement direction) as de-scribed by Nobusawa et al. (2017). Sh ble was replaced with the Streptomyceshygroscopicus aminoglycoside phosphotransferase gene (Aph7) or Streptomycesrimosus aminoglycoside 3-phosphotransferase gene (Aph8) to confer hygromycinB or paromomycin resistance, respectively. For making gene knockout con-structs,;1 kb of the 59 and 39 flanking genomic regions of the relevant genewereamplified by PCRusingN. oceanica genomic DNAas a template andwere clonedinto the PstI and KpnI sites of vector pUC18/NT7, respectively. The sequence ofthe flanking region was obtained from genome information of N. oceanicaIMET1 (Wang et al., 2014). For gene overexpression and complementation,

Figure 8. Schematic model for the DGTS biosynthesis in Nanno-chloropsis. We proposed that 20:5-containing DGTS and 20:5-freeDGTS are formed by different pathways with different physiologicalrelevance. The 20:5-enriched DAG derived from phospholipids woulddiverge to use for DGTS orMGDGbiosynthesis catalyzed byBTA1L andBTA1S or MGD1, respectively. Blue represents enzymes and blackrepresents metabolites.G3P, Glycerol 3-phosphate; MGD1, MGDGsynthase 1.

190 Plant Physiol. Vol. 177, 2018

Murakami et al.

https://plantphysiol.orgDownloaded on February 12, 2021. - Published by Copyright (c) 2020 American Society of Plant Biologists. All rights reserved.

the promoter sequence (;1 kb) of VCP1 (for overexpression) or lipid dropletsurface protein gene (for complementation) with a NotI site were cloned be-tween Sh ble (or Aph7 or Aph8) and terVCP1 of pUC18/NT7 by inverse PCR.BTA1L (2,295 bp) and BTA1LDC (1,335 bp with a termination codon) codingsequences were PCR amplified using a cDNA template and were clonedinto the NotI site (ProVCP2:sh ble:ProVCP1:gene:terVCP1). Information aboutprimers used for plasmid construction is provided in Supplemental Table S1.Linearized DNA fragments amplified from each construct and M13 for-ward (59-GTAAAACGACGGCCAGT-39) and M13 reverse (59-CAGGAAA-CAGCTATGAC-39) primers were used for nuclear transformation. Genetictransformation experiments were conducted essentially according to Kilianet al. (2011). Cells at the exponential phase of growth were centrifuged (4,900g,7 min) and washed twice with 375 mM sorbitol (Wako Pure Chemical Indus-tries). The pellet was resuspended in a small amount of sorbitol, and 100 mL ofthe resultant cell suspension and 3 to 5 mg linearized DNA were added to a2-mm cuvette (Bio-Rad) and then employed for electroporation performedwitha Gene Pulser II (Bio-Rad). The electroporation conditions were 2.2 kV, 50 mF,and 600 Ohm. Electroporated cells were immediately transferred to a culturetube with 5 mL F2N medium and then were shaken for 48 h under continuouslight. The cells were collected by centrifugation and resuspended in 5 mLmelted top agar (f/2 medium with 0.4% Bacto agar [Difco] and 18 g L–1 sea-water). Cell suspensions were sowed on f/2 agar plates (f/2 mediumwith 0.8%agar and 18 g L–1 seawater) containing 2 mg mL–1 zeocin (Thermo FisherScientific), 350 mg mL–1 hygromycin B (Wako Pure Chemical Industries), or150 mg mL–1 paromomycin (Sigma-Aldrich), depending on the antibiotic resistancegene used for transformation. After single colonies became visible, each line wasseeded into a 96-well plate filled with F2Nmedium. Genomic DNAwas isolated asdescribed by Nobusawa et al. (2017) and used for PCR-based screening with theprimers shown in Supplemental Table S1.

RNA Preparation and RT-qPCR

Total RNAwas preparedusing the TRI reagent (Sigma-Aldrich). First-strandcDNA was synthesized using Superscript II reverse transcriptase (Invitrogen).PCR products were used to confirm gene knockouts and for quantification ofrelative mRNA levels. Semiquantitative PCR was carried out with the primersdescribed in Supplemental Table S1. For quantitative PCR, cDNA was ampli-fied with SYBR Premix Ex Taq II (Takara), and the signal was detected with theThermal Cycler Dice Real-Time System (Takara). Delta Ct values were calcu-lated based on the Ct value for the gene encoding NADH dehydrogenasesubunit 11. RT-qPCR was carried out with the primers shown in SupplementalTable S1.

Lipid Analyses

Total cellular lipids were extracted according to Bligh and Dyer (1959). Polarlipids were separated by two-dimensional TLC using TLC silica plates (Merck),the solvent system chloroform/methanol/7 N ammonia water (120:80:8, v/v/v)as the first dimension, and chloroform/methanol/acetic acid/water(170:30:15:3, v/v/v/v) as the second dimension. To visualize each lipid classspot, each developed TLC plate was sprayed with 0.01% (w/v) primuline in80% (v/v) acetone and then irradiated with black light (352 nm). Each lipid spotwas scraped from the TLC plate, and 100 mL of 1 mM pentadecanoic acid wasadded as an internal standard. Each suspension was then incubated with 5%(v/v) HCl methanol solution (Supelco) at 85°C for 1 h for conversion into thecorresponding fatty acid methyl ester. The methyl esters were then extractedwith hexane and quantified via gas chromatography with a flame ionizationdetector (Shimadzu) with a HR-SS-10 capillary column (Shinwa ChemicalIndustries).

Accession Numbers

Sequence data of Nannochloropsis protein identified from this article can befound in the DNA Databank of Japan (http://www.ddbj.nig.ac.jp) with thefollowing accession numbers: BTA1S (LC375791), BTA1L (LC375792), PECT1(LC375793), and CCT1 (LC375794). The accession numbers used for the phy-logenetic analysis were as follows: Selaginella moellendorffii (XP_002980648),Physcomitrella patens (XP_001757434), Marchantia polymorpha (OAE21146),Klebsormidium nitens (GAQ92949), Coccomyxa subellipsoidea (XP_005649296),Chlorella variabilis NC64A (XP_005844132), Volvox carteri (XP_002955110),Chlamydomonas reinhardtii (XP_001700879),Micromonas pusilla (XP_003063768),

Ostreococcus lucimarinus (XP_001421536), Bigelowiella natans (JGI Protein ID89413), Emiliania huxleyi (XP_005763111), Thalassiosira oceanica (EJK50635),Fragilariopsis cylindrus (OEU16585), Phaeodactylum tricornutum (XP_002176772),Guillardia theta (XP_005822595), and Vitrella brassicaformis (CEM35387).

Supplemental Data

The following supplemental materials are available.

Supplemental Figure S1. Predicted substrate-binding sites for BTA1L andBTA1S and putative BTA1 homologs found in algae and plants.

Supplemental Figure S2. Phylogenetic tree for the putative BTA1 homologsfound in plants and algae.

Supplemental Figure S3. Fatty acid composition of each lipid class of thewild type grown under normal, -N, -Pi, and low-temperature conditions.

Supplemental Figure S4. Generation of knockout mutants of BTA1L andBTA1S via homologous recombination.

Supplemental Figure S5. Fatty acid composition of three TLC spots corre-sponding to molecular variants of DGTS.

Supplemental Figure S6. Membrane lipid composition of bta1s mutantsgrown under Pi-sufficient or -deficient conditions.

Supplemental Figure S7. Complementation with full length andC-terminally truncated BTA1L in mutants bta1l and bta1s;bta1l.

Supplemental Figure S8. Comparison of expression level of N-terminaldomain of BTA1L.

Supplemental Figure S9. Relative fatty acid composition of each lipid classof bta1l and bta1s;bta1l.

Supplemental Figure S10. Relative expression of BTA1L in the overexpres-sion mutant.

Supplemental Figure S11. Generation of mutants cct1 and pect1.

Supplemental Table S1. List of sequences of primers used in this study.

ACKNOWLEDGMENTS

N. oceanica strain NIES-2145 was obtained from the National Institute forEnvironmental Studies in Japan.

Received November 2, 2017; accepted February 21, 2018; published March 19,2018.

LITERATURE CITED

Abida H, Dolch LJ, Meï C, Villanova V, Conte M, Block MA, Finazzi G,Bastien O, Tirichine L, Bowler C, et al (2015) Membrane glycerolipidremodeling triggered by nitrogen and phosphorus starvation in Phaeo-dactylum tricornutum. Plant Physiol 167: 118–136

Alboresi A, Perin G, Vitulo N, Diretto G, Block M, Jouhet J, Meneghesso A,Valle G, Giuliano G, Maréchal E, Morosinotto T (2016) Light remodelslipid biosynthesis in Nannochloropsis gaditana by modulating carbon parti-tioning between organelles. Plant Physiol 171: 2468–2482

Babiychuk E, Müller F, Eubel H, Braun HP, Frentzen M, Kushnir S (2003)Arabidopsis phosphatidylglycerophosphate synthase 1 is essential forchloroplast differentiation, but is dispensable for mitochondrial func-tion. Plant J 33: 899–909

Benning C, Huang ZH, Gage DA (1995) Accumulation of a novel glyco-lipid and a betaine lipid in cells of Rhodobacter sphaeroides grown underphosphate limitation. Arch Biochem Biophys 317: 103–111

Bligh EG, Dyer WJ (1959) A rapid method of total lipid extraction andpurification. Can J Biochem Physiol 37: 911–917

Cañavate JP, Armada I, Ríos JL, Hachero-Cruzado I (2016) Exploring oc-currence and molecular diversity of betaine lipids across taxonomy ofmarine microalgae. Phytochemistry 124: 68–78

Cañavate JP, Armada I, Hachero-Cruzado I (2017) Interspecific variabilityin phosphorus-induced lipid remodelling among marine eukaryoticphytoplankton. New Phytol 213: 700–713

Plant Physiol. Vol. 177, 2018 191

Dual Roles for a Betaine Lipid in Nannochloropsis

https://plantphysiol.orgDownloaded on February 12, 2021. - Published by Copyright (c) 2020 American Society of Plant Biologists. All rights reserved.

Dembitsky VM (1996) Betaine ether-linked glycerolipids: chemistry andbiology. Prog Lipid Res 35: 1–51

Dolch LJ, Rak C, Perin G, Tourcier G, Broughton R, Leterrier M,Morosinotto T,Tellier F, Faure JD, Falconet D, et al (2017) A palmitic acid elongase affectseicosapentaenoic acid and plastidial monogalactosyldiacylglycerol levels inNannochloropsis. Plant Physiol 173: 742–759

Geiger O, Röhrs V, Weissenmayer B, Finan TM, Thomas-Oates JE (1999)The regulator gene phoB mediates phosphate stress-controlled synthesisof the membrane lipid diacylglyceryl-N,N,N-trimethylhomoserine inRhizobium (Sinorhizobium) meliloti. Mol Microbiol 32: 63–73

Geske T, Vom Dorp K, Dörmann P, Hölzl G (2013) Accumulation ofglycolipids and other non-phosphorous lipids in Agrobacterium tumefa-ciens grown under phosphate deprivation. Glycobiology 23: 69–80

Giroud C, Gerber A, Eichenberger W (1988) Lipids of Chlamydomonasreinhardtii. Analysis of molecular species and intracellular site(s) of bi-osynthesis. Plant Cell Physiol 29: 587–595

Guschina IA, Harwood JL (2006) Lipids and lipid metabolism in eukary-otic algae. Prog Lipid Res 45: 160–186

Haigh WG, Yoder TF, Ericson L, Pratum T, Winget RR (1996) The char-acterisation and cyclic production of a highly unsaturated homoserinelipid in Chlorella minutissima. Biochim Biophys Acta 1299: 183–190

Hirashima T, Toyoshima M, Moriyama T, Nakamura Y, Sato N (2017)Characterization of phosphoethanolamine-N-methyltransferases ingreen algae. Biochem Biophys Res Commun 488: 141–146

Hofmann M, Eichenberger W (1996) Biosynthesis of diacylglyceryl-N,N,N-trimethylhomoserine in Rhodobacter sphaeroides and evidence for lipid-linked N methylation. J Bacteriol 178: 6140–6144

Hoffmann M, Marxen K, Schulz R, Vanselow KH (2010) TFA and EPAproductivities of Nannochloropsis salina influenced by temperature andnitrate stimuli in turbidostatic controlled experiments. Mar Drugs 8:2526–2545

Hori K, Maruyama F, Fujisawa T, Togashi T, Yamamoto N, Seo M, Sato S,Yamada T, Mori H, Tajima N, et al (2014) Klebsormidium flaccidum ge-nome reveals primary factors for plant terrestrial adaptation. NatCommun 5: 3978

Hori K, Nobusawa T, Watanabe T, Madoka Y, Suzuki H, Shibata D,Shimojima M, Ohta H (2016) Tangled evolutionary processes withcommonality and diversity in plastidial glycolipid synthesis in photo-synthetic organisms. Biochim Biophys Acta 1861: 1294–1308

Hu H, Gao K (2006) Response of growth and fatty acid compositions ofNannochloropsis sp. to environmental factors under elevated CO2 con-centration. Biotechnol Lett 28: 987–992

Inatsugi R, Kawai H, Yamaoka Y, Yu Y, Sekiguchi A, Nakamura M,Nishida I (2009) Isozyme-specific modes of activation of CTP:phos-phorylcholine cytidylyltransferase in Arabidopsis thaliana at low tem-perature. Plant Cell Physiol 50: 1727–1735

Iwai M, Hori K, Sasaki-Sekimoto Y, Shimojima M, Ohta H (2015) Ma-nipulation of oil synthesis in Nannochloropsis strain NIES-2145 with aphosphorus starvation-inducible promoter from Chlamydomonas rein-hardtii. Front Microbiol 6: 912

Katoh K, Standley DM (2013) MAFFT multiple sequence alignment soft-ware version 7: improvements in performance and usability. Mol BiolEvol 30: 772–780

Kato M, Sakai M, Adachi K, Ikemoto H, Sano H (1996) Distribution ofbetaine lipids in marine algae. Phytochemistry 42: 1341–1345

Kaye Y, GrundmanO, Leu S, ZarkaA, Zorin B, Didi-Cohen S, Khozin-Goldberg I,Boussiba S (2015) Metabolic engineering toward enhanced LC-PUFA biosyn-thesis in Nannochloropsis oceanica: Overexpression of endogenous D12 desaturasedriven by stress-inducible promoter leads to enhanced deposition of polyunsat-urated fatty acids in TAG. Algal Res 11: 387–398

Khozin-Goldberg I, Didi-Cohen S, Cohen Z (2002) Biosynthesis of eico-sapentaenoic acid (EPA) in the freshwater eustigmatophyte Monodussubterraneus (eustigmatophyceae). J Phycol 38: 745–756

Khozin-Goldberg I, Cohen Z (2006) The effect of phosphate starvation onthe lipid and fatty acid composition of the fresh water eustigmatophyteMonodus subterraneus. Phytochemistry 67: 696–701

Kilian O, Benemann CSE, Niyogi KK, Vick B (2011) High-efficiency ho-mologous recombination in the oil-producing alga Nannochloropsis sp.Proc Natl Acad Sci USA 108: 21265–21269

Klug RM, Benning C (2001) Two enzymes of diacylglyceryl-O-49-(N,N,N,-trimethyl)homoserine biosynthesis are encoded by btaA and btaB in thepurple bacterium Rhodobacter sphaeroides. Proc Natl Acad Sci USA 98:5910–5915

Künzler K, Eichenberger W (1997) Betaine lipids and zwitterionic phos-pholipids in plants and fungi. Phytochemistry 46: 883–892

Li J, Han D, Wang D, Ning K, Jia J, Wei L, Jing X, Huang S, Chen J, Li Y,Hu Q, Xu J (2014) Choreography of transcriptomes and lipidomes ofNannochloropsis reveals the mechanisms of oil synthesis in microalgae.Plant Cell 26: 1645–1665

Li N, Xu C, Li-Beisson Y, Philippar K (2016) Fatty acid and lipid transportin plant cells. Trends Plant Sci 21: 145–158

Ma XN, Chen TP, Yang B, Liu J, Chen F (2016) Lipid production fromNannochloropsis. Mar Drugs 14: 61

Mizoi J, Nakamura M, Nishida I (2006) Defects in CTP:PHOSPHOR-YLETHANOLAMINE CYTIDYLYLTRANSFERASE affect embryonicand postembryonic development in Arabidopsis. Plant Cell 18: 3370–3385

Moore TS, Du Z, Chen Z (2001) Membrane lipid biosynthesis in Chlamy-domonas reinhardtii. In vitro biosynthesis of diacylglyceryl-trimethylhomoserine. Plant Physiol 125: 423–429

Mühlroth A, Li K, Røkke G, Winge P, Olsen Y, Hohmann-Marriott MF,Vadstein O, Bones AM (2013) Pathways of lipid metabolism in marinealgae, co-expression network, bottlenecks and candidate genes for en-hanced production of EPA and DHA in species of Chromista. Mar Drugs11: 4662–4697

Murakami R, Hashimoto H (2009) Unusual nuclear division in Nanno-chloropsis oculata (Eustigmatophyceae, Heterokonta) which may ensurefaithful transmission of secondary plastids. Protist 160: 41–49

Murata N, Los DA (1997) Membrane fluidity and temperature perception.Plant Physiol 115: 875–879

Nobusawa T, Hori K, Mori H, Kurokawa K, Ohta H (2017) Differentlylocalized lysophosphatidic acid acyltransferases crucial for triacylglyc-erol biosynthesis in the oleaginous alga Nannochloropsis. Plant J 90:547–559

Ohtaka K, Hori K, Kanno Y, Seo M, Ohta H (2017) Primitive auxin re-sponse without TIR1 and Aux/IAA in the charophyte alga Klebsormi-dium nitens. Plant Physiol 174: 1621–1632

Petroutsos D, Amiar S, Abida H, Dolch LJ, Bastien O, Rébeillé F, Jouhet J,Falconet D, Block MA, McFadden GI, et al (2014) Evolution of gal-actoglycerolipid biosynthetic pathways–from cyanobacteria to primaryplastids and from primary to secondary plastids. Prog Lipid Res 54: 68–85

Poliner E, Panchy N, Newton L, Wu G, Lapinsky A, Bullard B, Zienkiewicz A,Benning C, Shiu SH, Farré EM (2015) Transcriptional coordination ofphysiological responses in Nannochloropsis oceanica CCMP1779 underlight/dark cycles. Plant J 83: 1097–1113

Radakovits R, Jinkerson RE, Fuerstenberg SI, Tae H, Settlage RE, Boore JL,Posewitz MC (2012) Draft genome sequence and genetic transforma-tion of the oleaginous alga Nannochloropis gaditana. Nat Commun 3:686

Riekhof WR, Sears BB, Benning C (2005) Annotation of genes involved inglycerolipid biosynthesis in Chlamydomonas reinhardtii: discovery of thebetaine lipid synthase BTA1Cr. Eukaryot Cell 4: 242–252

Riekhof WR, Naik S, Bertrand H, Benning C, Voelker DR (2014) Phosphatestarvation in fungi induces the replacement of phosphatidylcholine withthe phosphorus-free betaine lipid diacylglyceryl-N,N,N-trimethylhomo-serine. Eukaryot Cell 13: 749–757

Rozentsvet OA, Dembitsky VM, Saksonov SV (2000) Occurrence of di-acylglyceryltrimethylhomoserines and major phospholipids in someplants. Phytochemistry 54: 401–407

Sato N, Furuya M (1985) Distribution of diacylglyceryl-trimethylhomoserine and phosphatidylcholine in non-vascular greenplants. Plant Sci 38: 81–85

Sato N, Mori N, Hirashima T, Moriyama T (2016) Diverse pathways ofphosphatidylcholine biosynthesis in algae as estimated by labelingstudies and genomic sequence analysis. Plant J 87: 281–292

Schneider JC, Roessler P (1994) Radiolabeling studies of lipids and fattyacids in Nannochloropsis (eustigmatophyceae), an oleaginous marinealga. J Phycol 30: 594–598

Senik SV, Maloshenok LG, Kotlova ER, Shavarda AL, Moiseenko KV,Bruskin SA, Koroleva OV, Psurtseva NV (2015) Diacylglyceryl-trimethylhomoserine content and gene expression changes triggered byphosphate deprivation in the mycelium of the basidiomycete Flammulinavelutipes. Phytochemistry 117: 34–42

Shemi A, Schatz D, Fredricks HF, Van Mooy BA, Porat Z, Vardi A (2016)Phosphorus starvation induces membrane remodeling and recycling inEmiliania huxleyi. New Phytol 211: 886–898

192 Plant Physiol. Vol. 177, 2018

Murakami et al.

https://plantphysiol.orgDownloaded on February 12, 2021. - Published by Copyright (c) 2020 American Society of Plant Biologists. All rights reserved.

Slocombe SP, Zhang Q, Ross M, Anderson A, Thomas NJ, Lapresa Á,Rad-Menéndez C, Campbell CN, Black KD, Stanley MS, Day JG(2015) Unlocking nature’s treasure-chest: screening for oleaginous algae.Sci Rep 5: 9844

Talavera G, Castresana J (2007) Improvement of phylogenies after re-moving divergent and ambiguously aligned blocks from protein se-quence alignments. Syst Biol 56: 564–577

Tamura K, Stecher G, Peterson D, Filipski A, Kumar S (2013) MEGA6: MolecularEvolutionary Genetics Analysis version 6.0. Mol Biol Evol 30: 2725–2729

Van Mooy BA, Fredricks HF, Pedler BE, Dyhrman ST, Karl DM, KoblízekM, Lomas MW, Mincer TJ, Moore LR, Moutin T, Rappé MS, Webb EA(2009) Phytoplankton in the ocean use non-phosphorus lipids in responseto phosphorus scarcity. Nature 458: 69–72

Vieler A,WuG, Tsai CH, Bullard B, CornishAJ, Harvey C, Reca IB, Thornburg C,Achawanantakun R, Buehl CJ, et al (2012) Genome, functional gene an-notation, and nuclear transformation of the heterokont oleaginous algaNannochloropsis oceanica CCMP1779. PLoS Genet 8: e1003064

Wang D, Ning K, Li J, Hu J, Han D, Wang H, Zeng X, Jing X, Zhou Q,Su X, et al (2014) Nannochloropsis genomes reveal evolution of micro-algal oleaginous traits. PLoS Genet 10: e1004094

Wang Q, Lu Y, Xin Y, Wei L, Huang S, Xu J (2016) Genome editing ofmodel oleaginous microalgae Nannochloropsis spp. by CRISPR/Cas9.Plant J 88: 1071–1081

Wei L, Xin Y, Wang Q, Yang J, Hu H, Xu J (2017) RNAi-based targetedgene knockdown in the model oleaginous microalgae Nannochloropsisoceanica. Plant J 89: 1236–1250

Zienkiewicz K, Zienkiewicz A, Poliner E, Du ZY, Vollheyde K, Herrfurth C,Marmon S, Farré EM, Feussner I, Benning C (2017) Nannochloropsis, a richsource of diacylglycerol acyltransferases for engineering of triacylglycerolcontent in different hosts. Biotechnol Biofuels 10: 8

Zorin B, Pal-Nath D, Lukyanov A, Smolskaya S, Kolusheva S, Didi-Cohen S,Boussiba S, Cohen Z, Khozin-Goldberg I, Solovchenko A (2017) Arachidonicacid is important for efficient use of light by the microalga Lobosphaera incisaunder chilling stress. Biochim Biophys Acta 1862: 853–868

Plant Physiol. Vol. 177, 2018 193

Dual Roles for a Betaine Lipid in Nannochloropsis

https://plantphysiol.orgDownloaded on February 12, 2021. - Published by Copyright (c) 2020 American Society of Plant Biologists. All rights reserved.