Membrane Lipid Oscillation: An Emerging System of Molecular Dynamics in the Plant Membrane

Membrane Lipid Oscillation: An Emerging System of Molecular Dynamics in the Plant Membrane Abstract Biological rhythm represents a major biological process of living organisms. However, rhythmic oscillation of membrane lipid content is poorly described in plants. The development of lipidomic technology has led to the illustration of precise molecular profiles of membrane lipids under various growth conditions. Compared with conventional lipid signaling, which produces unpredictable lipid changes in response to ever-changing environmental conditions, lipid oscillation generates a fairly predictable lipid profile, adding a new layer of biological function to the membrane system and possible cross-talk with the other chronobiological processes. This mini review covers recent studies elucidating membrane lipid oscillation in plants. Introduction Membrane lipids are indispensable components of biological membranes. Because of their fundamental role as a structural basis, membrane lipids are often considered rather inert metabolites. However, membrane lipid composition is established and maintained under the dynamic equilibrium of metabolic flux control. In plants, the polar glycerolipids, which have a hydrophilic head group and two hydrophobic acyl tails on a glycerol backbone, comprise the primary membrane lipid content. Because cellular membranes are composed of different classes of polar glycerolipids with distinct head and tail groups that are unevenly distributed among different organelles, the membrane lipid profile is a crucial factor that characterizes the physical property of organellar membranes. In addition, the membrane lipid composition is altered in response to environmental cues. For example, with cold stress, acyl groups of the lipids are polyunsaturated, so membrane fluidity is maintained under lower temperature (Wada et al. 1990). Indeed, Arabidopsis mutants defective in the unsaturation of acyl groups exhibit defective growth under low temperature (Somerville and Browse 1991, Miquel and Browse 1992, Browse et al. 1993). Moreover, during phosphate starvation, the phosphate-containing polar head group of membrane lipids is replaced by galactoses, producing galactolipids, which allow plants to obtain a scarce phosphate source yet maintain their membrane function, a process termed membrane lipid remodeling (Nakamura 2013). Furthermore, membrane lipids are the cellular reserve of lipid signals; on perception of external signals, different types of lipases produce a distinct type of lipid signal that triggers respective physiological responses (Hong et al. 2016). Although these responses are induced by unpredictable environmental stimuli, oscillatory signal inputs such as the diurnal cycle also affect the membrane lipidome profile. This oscillatory lipidome is highly distinct from conventional membrane lipid remodeling or signaling in that the future lipid profile is predictable based on past changes. As such, the underlying regulatory mechanism requires a high degree of fine metabolic regulation to establish and maintain the rhythmic profiles. In mammals, the functional connection between the circadian regulation of physiology and metabolism has been investigated, especially in the medical context (Adamovich et al. 2015). However, in plant lipid research, membrane lipid oscillation is an emerging issue whose physiological significance has been addressed only recently. Thus, this mini review aims to summarize recent advances in investigating rhythmic lipid changes in plants and proposes membrane lipid oscillation as a new model for discussing the physiology and regulatory mechanism of membrane lipid diversity. A Brief Overview of Membrane Lipid Metabolism Polar glycerolipids are primarily classified according to the structure of the polar head group. In plants, seven phospholipid classes, two galactolipid classes and one sulfolipid class are the major components of polar glycerolipids. Each polar glycerolipid class is subclassified according to the fatty acid species in the hydrophobic tails, which vary in number of carbon chains and number and position of unsaturated bonds. In plant cells, the composition of these polar glycerolipid classes differs among organelles. For example, chloroplasts are abundant in galactolipid monogalactosyldiacylglycerol (MGDG), which is essential for photosynthesis (Kobayashi et al. 2016). The combination of polar head group and fatty acid species results in a tremendous number of molecular species with distinct physical features. This membrane lipid diversity is a critical factor for characterizing the properties of the membrane system and the surrounding cellular environment. The biosynthesis of polar glycerolipids starts from the incorporation of fatty acids into glycerol 3-phosphate. The first reaction is catalyzed by glycerol 3-phosphate acyltransferase (GPAT) and the second by lysophosphatidate acyltransferase (LPAAT) (Fig. 1A). These two steps of reaction produce phosphatidic acid (PA), the structurally simplest phospholipid class. PA is a precursor for the synthesis of other polar glycerolipid classes and is also known as a lipid second messenger. To synthesize the other glycerolipid classes, PA is converted to diacylglycerol (DAG) or CDP-DAG (Fig. 1B). In the former case, the reaction is catalyzed by PA phosphatase (PAP) (Nakamura and Ohta 2010). The resulting DAG is further converted to phosphatidylcholine (PC) or phosphatidylethanolamine (PE) by aminoalcohol aminophosphotransferase (AAPT) (Liu et al. 2015), to MGDG or digalactosyldiacylglycerol (DGDG) by MGDG synthase (MGD) (Nakamura et al. 2010) and DGDG synthase (DGD) (Kalisch et al. 2016), respectively, or to sulfoquinovosyldiacylglycerol (SQDG) by SQD2 (Shimojima 2011). The AAPT-mediated reaction is assumed to occur in the endoplasmic reticulum, whereas the latter two reactions take place in the plastids. Phosphatidylserine (PS) is synthesized from PE by a base-exchanging activity catalyzed by PS synthase (PSS) (Yamaoka et al. 2011) and is converted back to PE by PS decarboxylase (PSD) (Nerlich et al. 2007). In the latter case, CDP-DAG is converted to phosphatidylinositol (PI) by PI synthase (PIS) (Löfke et al. 2008) or to phosphatidylglycerophosphate (PGP) by PGP synthase (PGPS) (Müller and Frentzen 2001). PGP is readily dephosphorylated by PGP phosphatase (PGPP) to produce phosphatidylglycerol (PG) (Lin et al. 2016, Zhou et al. 2017). In the mitochondrial inner envelope, PG is further converted to cardiolipin by cardiolipin synthase (Katayama et al. 2004, Nowicki et al. 2005), which is required for mitochondrial function (Pineau et al. 2013, Pan et al. 2014). Thus, glycerolipid metabolism consists of the following three steps: (i) incorporation of the fatty acid into the glycerol backbone; (ii) conversion of PA to DAG or CDP-DAG; and (iii) incorporation of the polar head group. Fig. 1 View largeDownload slide Polar glycerolipid metabolism in Arabidopsis. (A) Initial reactions to form phosphatidic acid (PA) from glycerol 3-phosphate (G3P) by incorporating acyl groups. Pink circles represent phosphate. Acyl groups are only schematically represented, and their molecular species are not shown in this illustration. (B) Subsequent reaction steps to produce different polar glycerolipid classes from PA. Green and orange arrows indicate plastidic and extraplastidic pathways, respectively. Compounds with gray shading are transitional precursors whose levels are usually very low. Acyl groups are only schematically represented, and their molecular species are not shown in this illustration. (C) Lipolytic pathways for phospholipids catalyzed by distinct types of phospholipases. Four different types of phospholipases cleave different positions of the phospholipid structure and give alternative reaction products. Broken arrows indicate the cleavage sites for each type of enzyme. Cho, choline; Etn, ethanolamine; FA, fatty acid; Gal, galactose; Gly, glycerol; Ins, inositol; PLA, phospholipase A; PLC, phospholipase C; PLD, phospholipase D; Ser, serine; Sq, sulfoquinovose. See the text and abbreviation list for the remaining abbreviations. Fig. 1 View largeDownload slide Polar glycerolipid metabolism in Arabidopsis. (A) Initial reactions to form phosphatidic acid (PA) from glycerol 3-phosphate (G3P) by incorporating acyl groups. Pink circles represent phosphate. Acyl groups are only schematically represented, and their molecular species are not shown in this illustration. (B) Subsequent reaction steps to produce different polar glycerolipid classes from PA. Green and orange arrows indicate plastidic and extraplastidic pathways, respectively. Compounds with gray shading are transitional precursors whose levels are usually very low. Acyl groups are only schematically represented, and their molecular species are not shown in this illustration. (C) Lipolytic pathways for phospholipids catalyzed by distinct types of phospholipases. Four different types of phospholipases cleave different positions of the phospholipid structure and give alternative reaction products. Broken arrows indicate the cleavage sites for each type of enzyme. Cho, choline; Etn, ethanolamine; FA, fatty acid; Gal, galactose; Gly, glycerol; Ins, inositol; PLA, phospholipase A; PLC, phospholipase C; PLD, phospholipase D; Ser, serine; Sq, sulfoquinovose. See the text and abbreviation list for the remaining abbreviations. The lipolytic pathways of polar glycerolipids are highly complex because of a number of lipases that hydrolyze different parts of the glycerolipid structure with distinct substrate specificity. Compared with galactolipids, hydrolysis of phospholipids is well investigated. At least four different types of phospholipases are known in Arabidopsis and other plant species. Phospholipase A1 (PLA1) and phospholipase A2 (PLA2) hydrolyze sn-1 and sn-2 positions of the acyl group to release a free fatty acid and a lysophospholipid (Chen et al. 2013). Phospholipase C (PLC) cleaves the ester bond between the glycerol backbone and the polar head group, giving a DAG and corresponding free polar head group (Munnik 2014, Nakamura 2014). Phospholipase D (PLD) hydrolyzes the ester bond between the phosphate group and the non-phosphorus part of the polar head group (Fig. 1C) (Wang et al. 2014). Oscillatory Profiles of Membrane Lipids An early study on the oscillatory profiles of membrane lipids investigated the fatty acid composition of total lipids as well as PC in spinach leaves (Browse et al. 1981). Under 10 h light and 14 h dark conditions, linolenic acid (18:3) content increased, with a concomitant decrease in oleic acid (18:1) content during the dark period. On entering the light period, 18:1 content was increased at the expense of 18:3 content. Moreover, the total amount of lipids decreased after the end of the dark period. Later, Arabidopsis leaves were analyzed for fatty acid profiles of total lipids as well as PC, PG and PE under long-day conditions (Ekman et al. 2007). During the light period, 18:1 content in total lipids was increased by 2-fold and contents of linoleic acid (18:2) and 18:3 were decreased. Under the dark period, 18:1 content was decreased and 18:2 and 18:3 contents were increased. This profile agreed with previous observations in spinach (Browse et al. 1981). Among PC, PE and PG, PC showed the most obvious diurnal fatty acid profile (Ekman et al. 2007). Diurnal fatty acid profiles of PC and PE were compared under short- and long-day conditions in Arabidopsis seedlings (Nakamura et al. 2014b). Seedlings were grown under short-day conditions for 2 weeks, then switched to long-day conditions or remained under short-day conditions for 1 d before time-course lipid profiling. The fatty acid composition of PE showed a rather stable profile and that of PC showed the profiles described previously; importantly, this profile clearly responded to different light/dark cycles (Nakamura et al. 2014b). Thus, the oscillatory profile of 18:1, 18:2 and 18:3 species is most probably diurnally controlled, and PC displays the clearest profile among the polar glycerolipid classes. Arabidopsis leaves under neutral-day (12 h light/12 h dark) conditions underwent a more comprehensive and sensitive analysis with lipidomics technology (Maatta et al. 2012): the content of most hexadecatrienoic acid (16:3) or 18:3-containing polar glycerolipid species as well as the total amount of PA and PS were increased during the dark period. In addition, levels of MGDG and DGDG were also significantly higher at the end of the dark period. Thus, minor polar glycerolipid classes as well as non-phosphorus glycerolipids also show diurnal profiles. Possible Mode of Regulation To establish and maintain the oscillatory profiles of membrane lipid content, a highly sophisticated regulatory mechanism is required at multiple levels (i.e. transcription, translation and post-translation). In early studies of spinach leaves, as mentioned in the previous section, fatty acid composition showed diurnal changes in the total polar glycerolipid and PC fractions (Browse et al. 1981). The biochemical and analytical studies suggested that the profile was established due to the flux balance between de novo fatty acid biosynthesis and polyunsaturation of acyl groups by fatty acid desaturases (FADs). In plants, fatty acids are synthesized in the chloroplasts via a circular reaction that elongates the chain length by two carbons. After the chain length reaches a certain range (C16 or C18 in most cases), the fatty acids are incorporated into the glycerol backbone by acyltransferase activity. Because plant FADs catalyzing polyunsaturation act on the acyl groups of glycerolipids, the polyunsaturation occurs after incorporation into the glycerol backbone. Importantly, acetyl-CoA carboxylase, which catalyzes the initial step of de novo fatty acid biosynthesis, is light dependent (Post-Beittenmiller et al. 1991), but such a requirement is not known for the FAD activity. Therefore, during the light period, de novo fatty acid biosynthesis has a stronger flux control over the polyunsaturation, so saturated fatty acids are the dominant content. In the dark, de novo fatty acid biosynthesis does not occur, but polyunsaturation continues, which increases the content of polyunsaturated fatty acids. This is a clear model to explain the profile of total polar glycerol lipids as well as PC (Browse et al. 1981). In addition, light affects the biosynthesis of some glycerolipid classes. For example, biosynthesis of MGDG, a primary and indispensable membrane lipid class in chloroplasts, is stimulated by light because MGDG synthase is activated by light (Yamaryo et al. 2003). Because MGDG is a major polar glycerolipid class and represents a pool of polyunsaturated fatty acids such as 18:3, light-regulated MGDG synthesis may be involved in the establishment of diurnal lipid profiles. Diurnal gene expression patterns of glycerolipid biosynthetic genes were investigated recently. In the publicly available transcriptome database accessed by a Web-based tool ‘Diurnal’ (http://diurnal.mocklerlab.org/), a number of glycerolipid biosynthetic genes showed significant diurnal profiles (Pearson’s correlation coefficient ≥0.8). These genes include LPAAT5/LPAT5, CDP-DIACYLGLYCEROL SYNTHASE 1 (CDS1), CDS4, DGD1, CHOLINE/ETHANOLAMINE KINASE3 (CEK3/CK5), CTP:PHOSPHORYLETHANOLAMINE CYTIDYLYLTRANSFERASE 1 (PECT1), AAPT2, PSD3, NON-SPECIFIC PHOSPHOLIPASE C6 (NPC6), PLDδ, DAG ACYLTRANSFERASE 1 (DGAT1) and PHOSPHOLIPID:DAG ACYLTRANSFERASE1 (PDAT1) (Nakamura et al. 2014a). Some of these genes have a crucial role in lipid metabolism. DGD1 is a major isoform of DGDG synthase, whose deletion significantly decreases DGDG content (Dörmann et al. 1999); PECT1 is a rate-limiting enzyme for PE biosynthesis, whose knockout abolishes PE and causes a lethal effect in plants (Mizoi et al. 2006); and DGAT1 and PDAT1 together account for triacylglycerol (TAG) biosynthesis, required for plant viability (Zhang et al. 2009). Although the content of these lipid classes under short- and long-day conditions has not been compared, some may show a diurnal profile because of their known function and high correlation with diurnal cycles in gene expression patterns. The database indicated that most of the genes, except NPC6 and PLDδ, showed circadian expression patterns. Indeed, the expression pattern of PECT1 was experimentally verified, showing a clear oscillatory profile that was identical between short- and long-day conditions (Nakamura et al. 2014a). A double mutant of the core clock system, CIRCADIAN CLOCK ASSOCIATED1 (CCA1) and LATE ELONGATED HYPOCOTYL (LHY), retained an oil body in young seedlings (Hsiao et al. 2014). This evidence suggests that the TAG metabolism involves regulation by a circadian clock. Physiological Role: A Case of Flowering Time Control A number of proteins bind a membrane lipid. In most cases, the ligand specificity is determined by the structure of both the polar head group and acyl chains, although the polar head group, whose hydrophilic nature allows for access by cytoplasmic proteins, is the primary determinant. However, in some cases, hydrophobic acyl chains may be critically involved in the function of protein–lipid binding. A recent example is the binding of FLOWERING LOCUS T (FT) protein to PC. FT is a component of florigen, a long-range mobile signal that moves from leaves to the shoot apex through the phloem to initiate flowering (Corbesier et al. 2007, Tamaki et al. 2007). With environmental cues that trigger flowering, FT is transcribed and translated in the leaf companion cells, from which the FT protein is exported to the sieve element (Andrés and Coupland 2012). In the shoot apex, FT interacts with FD protein and induces the expression of effector genes such as SUPPRESSOR OF OVEREXPRESSION OF CONSTANS1 (SOC1) and APETALA1 (AP1) (Abe et al. 2005, Wigge et al. 2005). Because the crystal structure of FT protein showed similarity to that of human PE-binding protein (Ahn et al. 2006), FT may also be a phospholipid-binding protein. Indeed, lipid–protein overlay and liposome co-precipitation assays revealed that FT bound to PC specifically (Nakamura et al. 2014b). A transgenic plant was created to alter relative PC levels by changing the expression level of PECT1. When PC levels were altered by using a leaf companion cell-specific promoter (SUC2 promoter), no effect on flowering time was observed. However, by using a shoot apex-specific promoter (FD promoter), it was demonstrated that the PC level in the shoot apex had a significant dose-dependent effect on flowering time. Transgenic plants with increased PC content in the shoot apex and, thus, early flowering showed increased expression of SOC1 and AP1. In addition, this effect was attenuated when FT was mutated, whereas simultaneous overexpression of FT and an increase in PC level had an additive effect on flowering time. Therefore, the early flowering effect by increasing the PC level in the shoot apex is due at least in part to FT activity, which suggests that PC binding activates FT protein function. Of note, FT prefers not to bind di18:3-PC, a PC species abundant during the dark period. PC containing 18:3 is synthesized by FAD3 (Arondel et al. 1992). To examine the effect of PC acyl species on flowering time, flowering time was assayed in a transgenic line overexpressing FAD3. Indeed, the overexpressor showed a late flowering phenotype, probably because the PC molecular species preferred by FT is at a low level (Nakamura et al. 2014b). This example demonstrates that diurnally controlled acyl profiles of membrane lipids affect protein–lipid binding to modulate protein function (Fig. 2A). Fig. 2 View largeDownload slide Membrane lipid oscillation. (A) Schematic representation of diurnal change of phosphatidylcholine (PC) molecular species and differential binding affinity for FLOWERING LOCUS T (FT) protein. During the day time, PC has more content of saturated acyl groups and binds well to FT protein. At night time, PC has three unsaturated bonds, which reduces the binding affinity of PC for FT. (B) Schematic representation of different membrane lipid dynamics. In lipid remodeling, lipid profiles are changed from the basal to altered status, but whether this profile changes in the future is unpredictable. In lipid signaling, a transient change follows rapid stimulation and attenuation of the metabolic reaction, with a unpredictable lipid profile. In lipid oscillation, lipid changes show periodical profiles, so future profiles are predictable. Fig. 2 View largeDownload slide Membrane lipid oscillation. (A) Schematic representation of diurnal change of phosphatidylcholine (PC) molecular species and differential binding affinity for FLOWERING LOCUS T (FT) protein. During the day time, PC has more content of saturated acyl groups and binds well to FT protein. At night time, PC has three unsaturated bonds, which reduces the binding affinity of PC for FT. (B) Schematic representation of different membrane lipid dynamics. In lipid remodeling, lipid profiles are changed from the basal to altered status, but whether this profile changes in the future is unpredictable. In lipid signaling, a transient change follows rapid stimulation and attenuation of the metabolic reaction, with a unpredictable lipid profile. In lipid oscillation, lipid changes show periodical profiles, so future profiles are predictable. Future Perspectives Membrane lipid oscillation is an emerging system of molecular dynamics in the plant membrane. It is highly distinct from conventional membrane lipid remodeling or lipid signaling: unlike these lipid changes, with lipid oscillation, the future lipid profile is predictable on the basis of past changes (Fig. 2B). Although oscillatory profiles of membrane lipid species have been reported (summarized above), their biological function and significance have not been investigated in plant science. As shown with FT–PC interaction, membrane lipid oscillation interacts with the protein function. This system provides an intriguing model for the field of lipid signaling. An important issue is to elucidate the underlying regulatory mechanism that requires a high degree of fine metabolic regulation to establish and maintain the rhythmic lipid profiles. Unlike conventional lipid signaling, which produces a transient stimulation and attenuation of a certain enzyme activity to control the amount of resulting signaling lipid, lipid oscillation requires a reversible change of metabolic reaction at a constant pace. This feature may require a highly sophisticated monitoring system. The effect of membrane lipid oscillation on the cellular environment is also an important issue from a cell biology and biophysical viewpoint. Given a striking change in fatty acid composition between day time and night time, the physical property of the membrane will differ considerably, which will affect the function of the membrane-interacting protein or any other molecular events occurring on the membranes. Whereas diurnal oscillation is dependent on light/dark cycles, circadian oscillation is created by the biological clock independent of light. Indeed, the effect of oscillating membrane lipids under the constant condition needs investigation because no report has been published on the circadian oscillation of membrane lipids. However, a number of lipid biosynthetic genes have a circadian profile of gene expression pattern (Nakamura et al. 2014a), so circadian profiles of membrane lipid species are anticipated. Moreover, investigating cross-talk between oscillating membrane lipid contents and other metabolism is worthwhile, given that metabolism is interconnected. For example, starch turnover is regulated by a diurnal cycle (Stitt and Zeeman 2012) for efficient carbon fixation after photosynthesis and its mobilization, and also for flowering because CONSTANS (CO), an upstream regulator of FT, is involved in photoperiodic modification of starch homeostasis to facilitate sugar mobilization to the shoot apex (Ortiz-Marchena et al. 2014). Moreover, diurnal oscillation can be found in the biosynthesis of anthocyanin (Pérez-García et al. 2015) and many other metabolites (Gibon et al. 2006). Since some of these metabolites are commercially important, a novel approach of metabolic engineering could be explored by modulating diurnal control of such metabolites. Furthermore, understanding the mechanism of such profiles will require examining an interaction with other chronobiological processes, such as regulation of lipid metabolism by a certain clock (or clock-regulated) gene. The concept of membrane lipid oscillation is still in its initial stages. A number of investigations are needed to delve into this emerging subject. Funding This work was supported by the Ministry of Science and Technology of Taiwan [MOST105-2628-B-001-006-MY3 to Y.N.] and the EMBO Young Investigator Program [to Y.N.]. Disclosures The author has no conflict of interest to declare. 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Google Scholar CrossRef Search ADS PubMed  Abbreviations Abbreviations AAPT aminoalcohol aminophosphotransferase AP1 APETALA1 CCA1 CIRCADIAN CLOCK ASSOCIATED1 CDP-DAG cytidine diphosphate diacylglycerol CDS CDP-DAG synthase CEK3 choline/ethanolamine kinase3 CO CONSTANS DAG diacylglycerol DGDG digalactosyldiacylglycerol DGD DGDG synthase FAD fatty acid desaturase FT FLOWERING LOCUS T GPAT glycerol 3-phosphate acyltransferase LPAAT lysophosphatidate acyltransferase LHY LATE ELONGATED HYPOCOTYL MGD MGDG synthase MGDG monogalactosyldiacylglycerol NPC6 non-specific phospholipase C6 PA phosphatidic acid PAP PA phosphatase PC phosphatidylcholine PDAT1 phospholipid:DAG acyltransferase1 PE phosphatidylethanolamine PECT1 CTP:phosphorylethanolamine cytidylyltransferase1 PG phosphatidylglycerol PGP phosphatidylglycerophosphate PGPS PGP synthase PGPP PGP phosphatase PI phosphatidylinositol PIS PI synthase PLA1 phospholipase A1 PLA2 phospholipase A2 PLC phospholipase C PLDδ phospholipase Dδ PS phosphatidylserine PSD PS decarboxylase PSS PS synthase SOC1 SUPPRESSOR OF OVEREXPRESSION OF CONSTANS1 SQDG sulfoquinovosyldiacylglycerol TAG triacylglycerol 16:3 hexadecatrienoic acid 18:1 oleic acid 18:2 linoleic acid 18:3 linolenic acid © The Author(s) 2018. Published by Oxford University Press on behalf of Japanese Society of Plant Physiologists. All rights reserved. For permissions, please email: journals.permissions@oup.com http://www.deepdyve.com/assets/images/DeepDyve-Logo-lg.png Plant and Cell Physiology Oxford University Press

Membrane Lipid Oscillation: An Emerging System of Molecular Dynamics in the Plant Membrane

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Abstract

Abstract Biological rhythm represents a major biological process of living organisms. However, rhythmic oscillation of membrane lipid content is poorly described in plants. The development of lipidomic technology has led to the illustration of precise molecular profiles of membrane lipids under various growth conditions. Compared with conventional lipid signaling, which produces unpredictable lipid changes in response to ever-changing environmental conditions, lipid oscillation generates a fairly predictable lipid profile, adding a new layer of biological function to the membrane system and possible cross-talk with the other chronobiological processes. This mini review covers recent studies elucidating membrane lipid oscillation in plants. Introduction Membrane lipids are indispensable components of biological membranes. Because of their fundamental role as a structural basis, membrane lipids are often considered rather inert metabolites. However, membrane lipid composition is established and maintained under the dynamic equilibrium of metabolic flux control. In plants, the polar glycerolipids, which have a hydrophilic head group and two hydrophobic acyl tails on a glycerol backbone, comprise the primary membrane lipid content. Because cellular membranes are composed of different classes of polar glycerolipids with distinct head and tail groups that are unevenly distributed among different organelles, the membrane lipid profile is a crucial factor that characterizes the physical property of organellar membranes. In addition, the membrane lipid composition is altered in response to environmental cues. For example, with cold stress, acyl groups of the lipids are polyunsaturated, so membrane fluidity is maintained under lower temperature (Wada et al. 1990). Indeed, Arabidopsis mutants defective in the unsaturation of acyl groups exhibit defective growth under low temperature (Somerville and Browse 1991, Miquel and Browse 1992, Browse et al. 1993). Moreover, during phosphate starvation, the phosphate-containing polar head group of membrane lipids is replaced by galactoses, producing galactolipids, which allow plants to obtain a scarce phosphate source yet maintain their membrane function, a process termed membrane lipid remodeling (Nakamura 2013). Furthermore, membrane lipids are the cellular reserve of lipid signals; on perception of external signals, different types of lipases produce a distinct type of lipid signal that triggers respective physiological responses (Hong et al. 2016). Although these responses are induced by unpredictable environmental stimuli, oscillatory signal inputs such as the diurnal cycle also affect the membrane lipidome profile. This oscillatory lipidome is highly distinct from conventional membrane lipid remodeling or signaling in that the future lipid profile is predictable based on past changes. As such, the underlying regulatory mechanism requires a high degree of fine metabolic regulation to establish and maintain the rhythmic profiles. In mammals, the functional connection between the circadian regulation of physiology and metabolism has been investigated, especially in the medical context (Adamovich et al. 2015). However, in plant lipid research, membrane lipid oscillation is an emerging issue whose physiological significance has been addressed only recently. Thus, this mini review aims to summarize recent advances in investigating rhythmic lipid changes in plants and proposes membrane lipid oscillation as a new model for discussing the physiology and regulatory mechanism of membrane lipid diversity. A Brief Overview of Membrane Lipid Metabolism Polar glycerolipids are primarily classified according to the structure of the polar head group. In plants, seven phospholipid classes, two galactolipid classes and one sulfolipid class are the major components of polar glycerolipids. Each polar glycerolipid class is subclassified according to the fatty acid species in the hydrophobic tails, which vary in number of carbon chains and number and position of unsaturated bonds. In plant cells, the composition of these polar glycerolipid classes differs among organelles. For example, chloroplasts are abundant in galactolipid monogalactosyldiacylglycerol (MGDG), which is essential for photosynthesis (Kobayashi et al. 2016). The combination of polar head group and fatty acid species results in a tremendous number of molecular species with distinct physical features. This membrane lipid diversity is a critical factor for characterizing the properties of the membrane system and the surrounding cellular environment. The biosynthesis of polar glycerolipids starts from the incorporation of fatty acids into glycerol 3-phosphate. The first reaction is catalyzed by glycerol 3-phosphate acyltransferase (GPAT) and the second by lysophosphatidate acyltransferase (LPAAT) (Fig. 1A). These two steps of reaction produce phosphatidic acid (PA), the structurally simplest phospholipid class. PA is a precursor for the synthesis of other polar glycerolipid classes and is also known as a lipid second messenger. To synthesize the other glycerolipid classes, PA is converted to diacylglycerol (DAG) or CDP-DAG (Fig. 1B). In the former case, the reaction is catalyzed by PA phosphatase (PAP) (Nakamura and Ohta 2010). The resulting DAG is further converted to phosphatidylcholine (PC) or phosphatidylethanolamine (PE) by aminoalcohol aminophosphotransferase (AAPT) (Liu et al. 2015), to MGDG or digalactosyldiacylglycerol (DGDG) by MGDG synthase (MGD) (Nakamura et al. 2010) and DGDG synthase (DGD) (Kalisch et al. 2016), respectively, or to sulfoquinovosyldiacylglycerol (SQDG) by SQD2 (Shimojima 2011). The AAPT-mediated reaction is assumed to occur in the endoplasmic reticulum, whereas the latter two reactions take place in the plastids. Phosphatidylserine (PS) is synthesized from PE by a base-exchanging activity catalyzed by PS synthase (PSS) (Yamaoka et al. 2011) and is converted back to PE by PS decarboxylase (PSD) (Nerlich et al. 2007). In the latter case, CDP-DAG is converted to phosphatidylinositol (PI) by PI synthase (PIS) (Löfke et al. 2008) or to phosphatidylglycerophosphate (PGP) by PGP synthase (PGPS) (Müller and Frentzen 2001). PGP is readily dephosphorylated by PGP phosphatase (PGPP) to produce phosphatidylglycerol (PG) (Lin et al. 2016, Zhou et al. 2017). In the mitochondrial inner envelope, PG is further converted to cardiolipin by cardiolipin synthase (Katayama et al. 2004, Nowicki et al. 2005), which is required for mitochondrial function (Pineau et al. 2013, Pan et al. 2014). Thus, glycerolipid metabolism consists of the following three steps: (i) incorporation of the fatty acid into the glycerol backbone; (ii) conversion of PA to DAG or CDP-DAG; and (iii) incorporation of the polar head group. Fig. 1 View largeDownload slide Polar glycerolipid metabolism in Arabidopsis. (A) Initial reactions to form phosphatidic acid (PA) from glycerol 3-phosphate (G3P) by incorporating acyl groups. Pink circles represent phosphate. Acyl groups are only schematically represented, and their molecular species are not shown in this illustration. (B) Subsequent reaction steps to produce different polar glycerolipid classes from PA. Green and orange arrows indicate plastidic and extraplastidic pathways, respectively. Compounds with gray shading are transitional precursors whose levels are usually very low. Acyl groups are only schematically represented, and their molecular species are not shown in this illustration. (C) Lipolytic pathways for phospholipids catalyzed by distinct types of phospholipases. Four different types of phospholipases cleave different positions of the phospholipid structure and give alternative reaction products. Broken arrows indicate the cleavage sites for each type of enzyme. Cho, choline; Etn, ethanolamine; FA, fatty acid; Gal, galactose; Gly, glycerol; Ins, inositol; PLA, phospholipase A; PLC, phospholipase C; PLD, phospholipase D; Ser, serine; Sq, sulfoquinovose. See the text and abbreviation list for the remaining abbreviations. Fig. 1 View largeDownload slide Polar glycerolipid metabolism in Arabidopsis. (A) Initial reactions to form phosphatidic acid (PA) from glycerol 3-phosphate (G3P) by incorporating acyl groups. Pink circles represent phosphate. Acyl groups are only schematically represented, and their molecular species are not shown in this illustration. (B) Subsequent reaction steps to produce different polar glycerolipid classes from PA. Green and orange arrows indicate plastidic and extraplastidic pathways, respectively. Compounds with gray shading are transitional precursors whose levels are usually very low. Acyl groups are only schematically represented, and their molecular species are not shown in this illustration. (C) Lipolytic pathways for phospholipids catalyzed by distinct types of phospholipases. Four different types of phospholipases cleave different positions of the phospholipid structure and give alternative reaction products. Broken arrows indicate the cleavage sites for each type of enzyme. Cho, choline; Etn, ethanolamine; FA, fatty acid; Gal, galactose; Gly, glycerol; Ins, inositol; PLA, phospholipase A; PLC, phospholipase C; PLD, phospholipase D; Ser, serine; Sq, sulfoquinovose. See the text and abbreviation list for the remaining abbreviations. The lipolytic pathways of polar glycerolipids are highly complex because of a number of lipases that hydrolyze different parts of the glycerolipid structure with distinct substrate specificity. Compared with galactolipids, hydrolysis of phospholipids is well investigated. At least four different types of phospholipases are known in Arabidopsis and other plant species. Phospholipase A1 (PLA1) and phospholipase A2 (PLA2) hydrolyze sn-1 and sn-2 positions of the acyl group to release a free fatty acid and a lysophospholipid (Chen et al. 2013). Phospholipase C (PLC) cleaves the ester bond between the glycerol backbone and the polar head group, giving a DAG and corresponding free polar head group (Munnik 2014, Nakamura 2014). Phospholipase D (PLD) hydrolyzes the ester bond between the phosphate group and the non-phosphorus part of the polar head group (Fig. 1C) (Wang et al. 2014). Oscillatory Profiles of Membrane Lipids An early study on the oscillatory profiles of membrane lipids investigated the fatty acid composition of total lipids as well as PC in spinach leaves (Browse et al. 1981). Under 10 h light and 14 h dark conditions, linolenic acid (18:3) content increased, with a concomitant decrease in oleic acid (18:1) content during the dark period. On entering the light period, 18:1 content was increased at the expense of 18:3 content. Moreover, the total amount of lipids decreased after the end of the dark period. Later, Arabidopsis leaves were analyzed for fatty acid profiles of total lipids as well as PC, PG and PE under long-day conditions (Ekman et al. 2007). During the light period, 18:1 content in total lipids was increased by 2-fold and contents of linoleic acid (18:2) and 18:3 were decreased. Under the dark period, 18:1 content was decreased and 18:2 and 18:3 contents were increased. This profile agreed with previous observations in spinach (Browse et al. 1981). Among PC, PE and PG, PC showed the most obvious diurnal fatty acid profile (Ekman et al. 2007). Diurnal fatty acid profiles of PC and PE were compared under short- and long-day conditions in Arabidopsis seedlings (Nakamura et al. 2014b). Seedlings were grown under short-day conditions for 2 weeks, then switched to long-day conditions or remained under short-day conditions for 1 d before time-course lipid profiling. The fatty acid composition of PE showed a rather stable profile and that of PC showed the profiles described previously; importantly, this profile clearly responded to different light/dark cycles (Nakamura et al. 2014b). Thus, the oscillatory profile of 18:1, 18:2 and 18:3 species is most probably diurnally controlled, and PC displays the clearest profile among the polar glycerolipid classes. Arabidopsis leaves under neutral-day (12 h light/12 h dark) conditions underwent a more comprehensive and sensitive analysis with lipidomics technology (Maatta et al. 2012): the content of most hexadecatrienoic acid (16:3) or 18:3-containing polar glycerolipid species as well as the total amount of PA and PS were increased during the dark period. In addition, levels of MGDG and DGDG were also significantly higher at the end of the dark period. Thus, minor polar glycerolipid classes as well as non-phosphorus glycerolipids also show diurnal profiles. Possible Mode of Regulation To establish and maintain the oscillatory profiles of membrane lipid content, a highly sophisticated regulatory mechanism is required at multiple levels (i.e. transcription, translation and post-translation). In early studies of spinach leaves, as mentioned in the previous section, fatty acid composition showed diurnal changes in the total polar glycerolipid and PC fractions (Browse et al. 1981). The biochemical and analytical studies suggested that the profile was established due to the flux balance between de novo fatty acid biosynthesis and polyunsaturation of acyl groups by fatty acid desaturases (FADs). In plants, fatty acids are synthesized in the chloroplasts via a circular reaction that elongates the chain length by two carbons. After the chain length reaches a certain range (C16 or C18 in most cases), the fatty acids are incorporated into the glycerol backbone by acyltransferase activity. Because plant FADs catalyzing polyunsaturation act on the acyl groups of glycerolipids, the polyunsaturation occurs after incorporation into the glycerol backbone. Importantly, acetyl-CoA carboxylase, which catalyzes the initial step of de novo fatty acid biosynthesis, is light dependent (Post-Beittenmiller et al. 1991), but such a requirement is not known for the FAD activity. Therefore, during the light period, de novo fatty acid biosynthesis has a stronger flux control over the polyunsaturation, so saturated fatty acids are the dominant content. In the dark, de novo fatty acid biosynthesis does not occur, but polyunsaturation continues, which increases the content of polyunsaturated fatty acids. This is a clear model to explain the profile of total polar glycerol lipids as well as PC (Browse et al. 1981). In addition, light affects the biosynthesis of some glycerolipid classes. For example, biosynthesis of MGDG, a primary and indispensable membrane lipid class in chloroplasts, is stimulated by light because MGDG synthase is activated by light (Yamaryo et al. 2003). Because MGDG is a major polar glycerolipid class and represents a pool of polyunsaturated fatty acids such as 18:3, light-regulated MGDG synthesis may be involved in the establishment of diurnal lipid profiles. Diurnal gene expression patterns of glycerolipid biosynthetic genes were investigated recently. In the publicly available transcriptome database accessed by a Web-based tool ‘Diurnal’ (http://diurnal.mocklerlab.org/), a number of glycerolipid biosynthetic genes showed significant diurnal profiles (Pearson’s correlation coefficient ≥0.8). These genes include LPAAT5/LPAT5, CDP-DIACYLGLYCEROL SYNTHASE 1 (CDS1), CDS4, DGD1, CHOLINE/ETHANOLAMINE KINASE3 (CEK3/CK5), CTP:PHOSPHORYLETHANOLAMINE CYTIDYLYLTRANSFERASE 1 (PECT1), AAPT2, PSD3, NON-SPECIFIC PHOSPHOLIPASE C6 (NPC6), PLDδ, DAG ACYLTRANSFERASE 1 (DGAT1) and PHOSPHOLIPID:DAG ACYLTRANSFERASE1 (PDAT1) (Nakamura et al. 2014a). Some of these genes have a crucial role in lipid metabolism. DGD1 is a major isoform of DGDG synthase, whose deletion significantly decreases DGDG content (Dörmann et al. 1999); PECT1 is a rate-limiting enzyme for PE biosynthesis, whose knockout abolishes PE and causes a lethal effect in plants (Mizoi et al. 2006); and DGAT1 and PDAT1 together account for triacylglycerol (TAG) biosynthesis, required for plant viability (Zhang et al. 2009). Although the content of these lipid classes under short- and long-day conditions has not been compared, some may show a diurnal profile because of their known function and high correlation with diurnal cycles in gene expression patterns. The database indicated that most of the genes, except NPC6 and PLDδ, showed circadian expression patterns. Indeed, the expression pattern of PECT1 was experimentally verified, showing a clear oscillatory profile that was identical between short- and long-day conditions (Nakamura et al. 2014a). A double mutant of the core clock system, CIRCADIAN CLOCK ASSOCIATED1 (CCA1) and LATE ELONGATED HYPOCOTYL (LHY), retained an oil body in young seedlings (Hsiao et al. 2014). This evidence suggests that the TAG metabolism involves regulation by a circadian clock. Physiological Role: A Case of Flowering Time Control A number of proteins bind a membrane lipid. In most cases, the ligand specificity is determined by the structure of both the polar head group and acyl chains, although the polar head group, whose hydrophilic nature allows for access by cytoplasmic proteins, is the primary determinant. However, in some cases, hydrophobic acyl chains may be critically involved in the function of protein–lipid binding. A recent example is the binding of FLOWERING LOCUS T (FT) protein to PC. FT is a component of florigen, a long-range mobile signal that moves from leaves to the shoot apex through the phloem to initiate flowering (Corbesier et al. 2007, Tamaki et al. 2007). With environmental cues that trigger flowering, FT is transcribed and translated in the leaf companion cells, from which the FT protein is exported to the sieve element (Andrés and Coupland 2012). In the shoot apex, FT interacts with FD protein and induces the expression of effector genes such as SUPPRESSOR OF OVEREXPRESSION OF CONSTANS1 (SOC1) and APETALA1 (AP1) (Abe et al. 2005, Wigge et al. 2005). Because the crystal structure of FT protein showed similarity to that of human PE-binding protein (Ahn et al. 2006), FT may also be a phospholipid-binding protein. Indeed, lipid–protein overlay and liposome co-precipitation assays revealed that FT bound to PC specifically (Nakamura et al. 2014b). A transgenic plant was created to alter relative PC levels by changing the expression level of PECT1. When PC levels were altered by using a leaf companion cell-specific promoter (SUC2 promoter), no effect on flowering time was observed. However, by using a shoot apex-specific promoter (FD promoter), it was demonstrated that the PC level in the shoot apex had a significant dose-dependent effect on flowering time. Transgenic plants with increased PC content in the shoot apex and, thus, early flowering showed increased expression of SOC1 and AP1. In addition, this effect was attenuated when FT was mutated, whereas simultaneous overexpression of FT and an increase in PC level had an additive effect on flowering time. Therefore, the early flowering effect by increasing the PC level in the shoot apex is due at least in part to FT activity, which suggests that PC binding activates FT protein function. Of note, FT prefers not to bind di18:3-PC, a PC species abundant during the dark period. PC containing 18:3 is synthesized by FAD3 (Arondel et al. 1992). To examine the effect of PC acyl species on flowering time, flowering time was assayed in a transgenic line overexpressing FAD3. Indeed, the overexpressor showed a late flowering phenotype, probably because the PC molecular species preferred by FT is at a low level (Nakamura et al. 2014b). This example demonstrates that diurnally controlled acyl profiles of membrane lipids affect protein–lipid binding to modulate protein function (Fig. 2A). Fig. 2 View largeDownload slide Membrane lipid oscillation. (A) Schematic representation of diurnal change of phosphatidylcholine (PC) molecular species and differential binding affinity for FLOWERING LOCUS T (FT) protein. During the day time, PC has more content of saturated acyl groups and binds well to FT protein. At night time, PC has three unsaturated bonds, which reduces the binding affinity of PC for FT. (B) Schematic representation of different membrane lipid dynamics. In lipid remodeling, lipid profiles are changed from the basal to altered status, but whether this profile changes in the future is unpredictable. In lipid signaling, a transient change follows rapid stimulation and attenuation of the metabolic reaction, with a unpredictable lipid profile. In lipid oscillation, lipid changes show periodical profiles, so future profiles are predictable. Fig. 2 View largeDownload slide Membrane lipid oscillation. (A) Schematic representation of diurnal change of phosphatidylcholine (PC) molecular species and differential binding affinity for FLOWERING LOCUS T (FT) protein. During the day time, PC has more content of saturated acyl groups and binds well to FT protein. At night time, PC has three unsaturated bonds, which reduces the binding affinity of PC for FT. (B) Schematic representation of different membrane lipid dynamics. In lipid remodeling, lipid profiles are changed from the basal to altered status, but whether this profile changes in the future is unpredictable. In lipid signaling, a transient change follows rapid stimulation and attenuation of the metabolic reaction, with a unpredictable lipid profile. In lipid oscillation, lipid changes show periodical profiles, so future profiles are predictable. Future Perspectives Membrane lipid oscillation is an emerging system of molecular dynamics in the plant membrane. It is highly distinct from conventional membrane lipid remodeling or lipid signaling: unlike these lipid changes, with lipid oscillation, the future lipid profile is predictable on the basis of past changes (Fig. 2B). Although oscillatory profiles of membrane lipid species have been reported (summarized above), their biological function and significance have not been investigated in plant science. As shown with FT–PC interaction, membrane lipid oscillation interacts with the protein function. This system provides an intriguing model for the field of lipid signaling. An important issue is to elucidate the underlying regulatory mechanism that requires a high degree of fine metabolic regulation to establish and maintain the rhythmic lipid profiles. Unlike conventional lipid signaling, which produces a transient stimulation and attenuation of a certain enzyme activity to control the amount of resulting signaling lipid, lipid oscillation requires a reversible change of metabolic reaction at a constant pace. This feature may require a highly sophisticated monitoring system. The effect of membrane lipid oscillation on the cellular environment is also an important issue from a cell biology and biophysical viewpoint. Given a striking change in fatty acid composition between day time and night time, the physical property of the membrane will differ considerably, which will affect the function of the membrane-interacting protein or any other molecular events occurring on the membranes. Whereas diurnal oscillation is dependent on light/dark cycles, circadian oscillation is created by the biological clock independent of light. Indeed, the effect of oscillating membrane lipids under the constant condition needs investigation because no report has been published on the circadian oscillation of membrane lipids. However, a number of lipid biosynthetic genes have a circadian profile of gene expression pattern (Nakamura et al. 2014a), so circadian profiles of membrane lipid species are anticipated. Moreover, investigating cross-talk between oscillating membrane lipid contents and other metabolism is worthwhile, given that metabolism is interconnected. For example, starch turnover is regulated by a diurnal cycle (Stitt and Zeeman 2012) for efficient carbon fixation after photosynthesis and its mobilization, and also for flowering because CONSTANS (CO), an upstream regulator of FT, is involved in photoperiodic modification of starch homeostasis to facilitate sugar mobilization to the shoot apex (Ortiz-Marchena et al. 2014). Moreover, diurnal oscillation can be found in the biosynthesis of anthocyanin (Pérez-García et al. 2015) and many other metabolites (Gibon et al. 2006). Since some of these metabolites are commercially important, a novel approach of metabolic engineering could be explored by modulating diurnal control of such metabolites. Furthermore, understanding the mechanism of such profiles will require examining an interaction with other chronobiological processes, such as regulation of lipid metabolism by a certain clock (or clock-regulated) gene. The concept of membrane lipid oscillation is still in its initial stages. A number of investigations are needed to delve into this emerging subject. Funding This work was supported by the Ministry of Science and Technology of Taiwan [MOST105-2628-B-001-006-MY3 to Y.N.] and the EMBO Young Investigator Program [to Y.N.]. Disclosures The author has no conflict of interest to declare. 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Google Scholar CrossRef Search ADS PubMed  Abbreviations Abbreviations AAPT aminoalcohol aminophosphotransferase AP1 APETALA1 CCA1 CIRCADIAN CLOCK ASSOCIATED1 CDP-DAG cytidine diphosphate diacylglycerol CDS CDP-DAG synthase CEK3 choline/ethanolamine kinase3 CO CONSTANS DAG diacylglycerol DGDG digalactosyldiacylglycerol DGD DGDG synthase FAD fatty acid desaturase FT FLOWERING LOCUS T GPAT glycerol 3-phosphate acyltransferase LPAAT lysophosphatidate acyltransferase LHY LATE ELONGATED HYPOCOTYL MGD MGDG synthase MGDG monogalactosyldiacylglycerol NPC6 non-specific phospholipase C6 PA phosphatidic acid PAP PA phosphatase PC phosphatidylcholine PDAT1 phospholipid:DAG acyltransferase1 PE phosphatidylethanolamine PECT1 CTP:phosphorylethanolamine cytidylyltransferase1 PG phosphatidylglycerol PGP phosphatidylglycerophosphate PGPS PGP synthase PGPP PGP phosphatase PI phosphatidylinositol PIS PI synthase PLA1 phospholipase A1 PLA2 phospholipase A2 PLC phospholipase C PLDδ phospholipase Dδ PS phosphatidylserine PSD PS decarboxylase PSS PS synthase SOC1 SUPPRESSOR OF OVEREXPRESSION OF CONSTANS1 SQDG sulfoquinovosyldiacylglycerol TAG triacylglycerol 16:3 hexadecatrienoic acid 18:1 oleic acid 18:2 linoleic acid 18:3 linolenic acid © The Author(s) 2018. Published by Oxford University Press on behalf of Japanese Society of Plant Physiologists. All rights reserved. For permissions, please email: journals.permissions@oup.com

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Plant and Cell PhysiologyOxford University Press

Published: Mar 1, 2018

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