Eukaryotic Cell IAI Online 2003
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Supplemental material
Right arrow Other Versions of this Article:
EC.00393-06v1
6/6/1006    most recent
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow Copyright Information
Right arrow Books from ASM Press
Right arrow MicrobeWorld
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Sato, N.
Right arrow Articles by Moriyama, T.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Sato, N.
Right arrow Articles by Moriyama, T.
Eukaryotic Cell, June 2007, p. 1006-1017, Vol. 6, No. 6
1535-9778/07/$08.00+0     doi:10.1128/EC.00393-06
Copyright © 2007, American Society for Microbiology. All Rights Reserved.

Genomic and Biochemical Analysis of Lipid Biosynthesis in the Unicellular Rhodophyte Cyanidioschyzon merolae: Lack of a Plastidic Desaturation Pathway Results in the Coupled Pathway of Galactolipid Synthesis{triangledown} ,{dagger}

Naoki Sato* and Takashi Moriyama

Department of Life Sciences, Graduate School of Arts and Sciences, The University of Tokyo, Komaba 3-8-1, Meguro-ku, Tokyo 153-8902, Japan

Received 8 December 2006/ Accepted 20 March 2007


   ABSTRACT
Top
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The acyl lipids making up the plastid membranes in plants and algae are highly enriched in polyunsaturated fatty acids and are synthesized by two distinct pathways, known as the prokaryotic and eukaryotic pathways, which are located within the plastids and the endoplasmic reticulum, respectively. Here we report the results of biochemical as well as genomic analyses of lipids and fatty acids in the unicellular rhodophyte Cyanidioschyzon merolae. All of the glycerolipids usually found in photosynthetic algae were found, such as mono- and digalactosyl diacylglycerol, sulfolipid, phosphatidylglycerol, phosphatidylcholine, phosphatidylethanolamine, and phosphatidylinositol. However, the fatty acid composition was extremely simple. Only palmitic, stearic, oleic, and linoleic acids were found as major acids. In addition, 3-trans-hexadecanoic acid was found as a very minor component in phosphatidylglycerol. Unlike the case for most other photosynthetic eukaryotes, polyenoic fatty acids having three or more double bonds were not detected. These results suggest that polyunsaturated fatty acids are not necessary for photosynthesis in eukaryotes. Genomic analysis suggested that C. merolae lacks acyl lipid desaturases of cyanobacterial origin as well as stearoyl acyl carrier protein desaturase, both of which are major desaturases in plants and green algae. The results of labeling experiments with radioactive acetate showed that the desaturation leading to linoleic acid synthesis occurs on phosphatidylcholine located outside the plastids. Monogalactosyl diacylglycerol is therefore synthesized by the coupled pathway, using plastid-derived palmitic acid and endoplasmic reticulum-derived linoleic acid. These results highlight essential differences in lipid biosynthetic pathways between the red algae and the green lineage, which includes plants and green algae.


   INTRODUCTION
Top
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The acyl lipids that make up the plastid membranes are highly enriched in polyunsaturated fatty acids in most plants and algae. In flowering plants, linoleic (18:2) and linolenic (18:3) acids are the most common unsaturated acids, whereas in many algae, highly unsaturated long-chain acids, such as arachidonic (20:4), eicosapentaenoic (20:5), and docosahexaenoic (22:6) acids, are found as major fatty acid components, depending on the species (4, 15, 17, 38, 50). (Fatty acids are expressed by a combination of the number of carbon atoms [X] and the number of double bonds [Y], such as X:Y. The positions of double bonds are specified in parentheses.) Unsaturated fatty acids, especially polyunsaturated ones, are important in maintaining membrane fluidity during the cold acclimation of photosynthesis (37). Linolenic acid was also found to be important in tolerance to high temperatures (32). The plastid lipids consist of two types of molecular species, namely, 1-C18-2-C16 species, or prokaryotic molecular species, and 1-C18-2-C18 species, or eukaryotic species. The relative abundance of these two types of molecular species varies with plants and algae. In flowering plants, the prokaryotic and eukaryotic types of lipid molecular species are synthesized by two distinct pathways, known as the prokaryotic and eukaryotic pathways, which are located within the plastids and the endoplasmic reticulum (ER), respectively (for a review, see reference 10). However, lipid biosynthesis in red algae, which constitute another major lineage of photosynthetic eukaryotes (11, 30, 42, 56), remains largely unknown.

Cyanidioschyzon merolae, a unicellular rhodophyte isolated from an Italian hot spring, has a very simple cell structure consisting of one mitochondrion, plastid, and microbody per cell (25). Its normal habitat is warm (up to 50°C) and acidic (pH 1.5 to 2.5) water containing sulfuric acid. The size of the nuclear genome is 16.5 Mbp (28). The cell proliferates by binary fission. These characteristics, as well as phylogenetic analyses (34), suggested that C. merolae is one of the most primitive red algae, probably diverged from near the root of the red lineage. The red lineage includes red algae, whereas the green lineage includes green algae and land plants (42). The single origin of plastids in the red and green lineages is believed to be highly probable (27, 42), and the single origin of plastid-harboring cells in these two lineages is gaining supporting evidence (30, 34). In addition, the chromists (brown algae, diatoms, cryptophytes, etc.) are believed to originate from secondary endosymbiosis by an ancestral red algal cell (11, 42, 56). C. merolae is therefore a good target of comparative biochemistry to reveal similarities and differences in the red and green lineages.

There is a short report on the total fatty acids of C. merolae (31). Among thermoacidophilic red algae, Cyanidium caldarium and C. merolae contained no detectable {alpha}-linolenic acid (18:3), while this acid was abundant in Pleurococcus sulfurarius (currently called Galdieria sulfuraria) at a low temperature. There was a report on {alpha}-18:3 in Cyanidium caldarium (23), but the current understanding is that several different algae were called Cyanidium in the past. No further analysis of the composition and biosynthesis of C. merolae has been attempted since the paper by Moretti and Nazzaro (31).

Apart from experimental analyses, one postgenomic study is to find all possible candidate enzymes of a metabolic pathway. For Arabidopsis thaliana, an attempt to make functional annotations for all proteins involved in lipid metabolism is in progress (7). For Chlamydomonas reinhardtii, draft sequence data were used to predict proteins that might be involved in lipid biosynthesis (36). The functional annotation of C. merolae has also been started (28). We have been trying to compare the genome contents of A. thaliana and C. merolae as well as cyanobacteria, and many proteins that are conserved in all of these photosynthetic organisms are being identified (45). Comparative genomics of these photosynthetic organisms are now conveniently analyzed through a web interface called the Gclust server (43; http://gclust.c.u-tokyo.ac.jp/). Based on such informatics, we are now able to predict probable proteins that are involved in acyl lipid metabolism in C. merolae.

In this report, we try to find distinct features of the red lineage with respect to lipid biosynthesis, using C. merolae as a model organism. We present results on the analysis of lipids and fatty acids in C. merolae. Next, lipid biosynthetic pathways of this alga are summarized based on the genomic data. The intracellular localization of some key enzymes was confirmed by green fluorescent protein (GFP) experiments. The origin of the minimal set of desaturases of this alga was established by comprehensive phylogenetic analysis. Finally, tracer experiments were conducted to validate the lipid biosynthetic pathway estimated by the genomic analysis. The coupled pathway for the synthesis of galactolipids is proposed, involving plastid-derived palmitic acid and extraplastidically synthesized linoleic acid. All of these data indicate distinct differences between the red and green lineages in lipid biosynthesis, in spite of their monophyletic origin.


   MATERIALS AND METHODS
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Growth of organism. Cells of Cyanidioschyzon merolae strain 10D (55) were grown under continuous white light (about 20 µE m–2 s–1) in the AA medium described by Allen (2), with aeration in 1% CO2, at 38°C or 25°C.

Analysis of lipids and fatty acids. All analytical methods used were essentially the same as those described in previous publications (44, 48). Briefly, lipids were extracted from the cells by the method of Bligh and Dyer (8) and were separated by two-dimensional thin-layer chromatography (TLC). Lipids were quantified by measuring the amounts of fatty acids, determined as their methyl esters, by gas chromatography. A fused silica capillary column (0.25-mm internal diameter by 50 m) coated with SS-10 (equivalent to Silar 10C; Sinwa Kako, Kyoto, Japan) was used. The following temperature program was used: 0.5 min at 180°C, a linear increase to 230°C at a rate of 3°C min–1, and then 10 min at 230°C. Under these conditions, most commonly occurring isomers of fatty acid methyl esters were clearly separated. Fatty acid methyl esters from total lipids of Adiantum capillus-veneris (44) and those from monogalactosyl diacylglycerol (MGDG) of Anabaena variabilis (46) were used as references. The positional distribution of fatty acids within individual classes of lipids (including phospholipids) was analyzed by specific hydrolysis of the C-1 acyl ester linkage with the lipase from Rhizopus delemar (16) or the C-2 acyl ester linkage with phospholipase A2.

Radiolabeling of lipids. C. merolae cells (25-ml culture) that had been grown at 38°C were incubated with [2-14C]acetate (2.0 MBq) at 38°C for 1 h in the light, with vigorous shaking, in a tightly closed 100-ml flask. Unlabeled acetate (3 mM) was then added, and the cells were harvested by centrifugation. They were washed once with fresh medium and then resuspended in fresh medium. The cells were allowed to grow under normal growth conditions for 20 h. Aliquots were withdrawn at 0, 2, 6, 10, and 20 h, and lipids were extracted. Lipids were separated by two-dimensional TLC, and then the lipid spots were detected with primuline under UV light. The analysis of radioactive lipids was performed essentially as described previously (39). Radioactivity was located by autoradiography. Radioactive lipid spots were scraped off, and the radioactivity was measured by liquid scintillation counting.

For detailed analysis, MGDG, digalactosyl diacylglycerol (DGDG), and phosphatidylcholine (PC) were recovered from the TLC plate. Lipid molecular species were analyzed by argentation TLC (48), and radioactivity was detected by autoradiography. A precoated silica gel plate (Merck) was impregnated with AgNO3 by immersing it in 5% AgNO3 in acetonitrile for 30 min and then was dried at 60°C for 30 min. The developing solvents were acetone-benzene-water (90:30:8, by volume) for MGDG and chloroform-methanol-water (60:30:5, by volume) for DGDG and PC. For fatty acid analysis of individual lipid classes, the isolated lipids were subjected to methanolysis. The resultant fatty acid methyl esters were analyzed by reversed-phase argentation TLC (26), a technique recently developed for the analysis of small amounts of radioactive fatty acids. An RP-18 HPTLC plate (5 by 10 cm; Merck) was used. The developing solvent was 10% AgNO3 in acetonitrile-1,4-dioxane-acetic acid (80:20:1, by volume). This TLC method clearly resolves 18:1 and 16:0, which comigrate in ordinary argentation TLC.

Incorporation of radioactive galactose was performed by incubating isolated plastids of C. merolae, which were prepared according to a published protocol (51), with modifications (T. Moriyama, K. Terasawa, M. Fujiwara, and N. Sato, unpublished data), with 37 kBq UDP-[U-14C]galactose (GE Healthcare/Amersham) in 400 µl plastid isolation medium at 38°C for 1 h. Lipids were extracted and separated by two-dimensional TLC. MGDG and DGDG were recovered, and the molecular species were analyzed by argentation TLC.

Genomic data and computational sequence analysis. Genomic data for C. merolae were generated and annotated by the Cyanidioschyzon Genome Project (28), in which genes related to lipid metabolism were estimated by BLAST2 (3) searches, with known genes as queries. The sequences of the seed genes were retrieved from GenomeNet (ftp://ftp.genome.ad.jp/). The Gclust database, recently made publicly accessible in the Gclust server (43), was also used to find phylogenetically conserved proteins. Sequence manipulation was performed with the SISEQ package, version 1.30 (40). The sequence alignment of desaturases was prepared by the Clustal X program (12) after trimming of poorly conserved N and C termini. Phylogenetic analysis was done with MEGA2 software, version 2.1 (24), PAUP software, version 4 beta 10 (Sinauer Associates, Sunderland, MA), and the MOLPHY package, version 2 beta 3 (1).

Targeting of GFP fusion proteins. DNA constructs consisting of the 35S promoter, a 5' part of the putative {Delta}12 desaturase gene, and the NOS terminator were made by successive PCRs. The sGFP plasmid (13) was used as the template for the 5' and the 3' parts, while the PCR fragment corresponding to various parts of the C. merolae CMK291C gene was obtained by using the C. merolae genome as a template. The following three constructs were made: Met1-GFP, Met2-GFP, and Met3-GFP, containing residues 1 to 531, 202 to 531, and 364 to 531, respectively (numbers refer to the nucleotide count beginning from the most upstream ATG). The 5' part includes the 35S promoter until the multiple cloning site, while the 3' part includes the NOS terminator sequence beginning from the multiple cloning site in the sGFP plasmid. The constructs were introduced into a scaly leaf of an onion bulb by particle bombardment using a PDS-1000/He particle delivery system (Bio-Rad). Rupture disks for 1,100 lb/in2 and tungsten particles with a diameter of 1.1 µm were used. After 24 h of continued growth, the epidermis was peeled and examined under a fluorescence microscope (Olympus model BX-60) with an IB filter cube.


   RESULTS
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Compositions of lipids and fatty acids. We started by analyzing the compositions of lipids and fatty acids in C. merolae cells (Table 1). Major lipids included common chloroplast lipids, such as MGDG, DGDG, and sulfoquinovosyl diacylglycerol (SQDG), as well as common phospholipids, namely, PG, phosphatidylcholine (PC), phosphatidylethanolamine (PE), and phosphatidylinositol (PI). The level of MGDG was higher at 25°C than at 38°C, whereas the level of PC was lower. Cardiolipin (diphosphatidylglycerol) and phosphatidylserine were not detected by the methods employed (see Discussion).


View this table:
[in this window]
[in a new window]

  		TABLE 1. Fatty acid composition of individual lipid classes at two different growth temperatures

 
Figure 1 shows representative results of capillary gas chromatographic analysis of fatty acid methyl esters. The composition of fatty acids in the total lipids (Fig. 1A) was strikingly simple. Palmitic (16:0; peak 1) and linoleic (18:2; peak 11) acids were the major acids. Stearic (18:0; peak 8) and oleic (18:1; peak 9) acids were also found as abundant components. PG contained the unusual fatty acid 3-trans-hexadecanoic acid (3-trans-16:1; peak 3), as do the PGs from all plants and algae. It is worth noting that no peak corresponding to either 18:3(9,12,15) (Fig. 1A, arrow) or 18:3(6,9,12) was detected in the analysis of fatty acids of C. merolae (Fig. 1A), even at a low temperature (Table 1). No peak corresponding to 18:3 isomers was detected in the analysis of any individual classes of lipids. This was also confirmed by direct methanolysis of the dried whole cells. Long-chain fatty acids, such as 20:4 and 22:6, were not detected.


Figure 1
View larger version (15K):
[in this window]
[in a new window]

  		FIG. 1. Gas chromatographic separation of fatty acid methyl esters. A capillary column (50 m long) coated with SS-10 was used. (A) Fatty acid methyl esters prepared from the total lipids of C. merolae cells grown at 25°C. (Inset) Enlargement (16-fold) to show the absence of 18:3(9,12,15) (arrow). (B) Fatty acid methyl esters prepared from the total lipids of a fern, Adiantum capillus-veneris. All peaks were identified previously (44) and served as a reference. The peaks before peak 1 were degradation products of pigments. (C) Fatty acid methyl esters prepared from the PG of C. merolae cells. (D) Reference fatty acid methyl esters prepared from MGDG of a cyanobacterium, Anabaena variabilis (46). Peaks: 1, 16:0; 2, 16:1(9); 3, 16:1(3-trans); 4, 16:2(9,12); 5, 17:0; 6, 17:1(9); 7, 16:3(7,10,13); 8, 18:0; 9, 18:1(9); 10, 18:1(11); 11, 18:2(9,12); 12, 18:3(6,9,12); 13, 18:3(9,12,15); and 14, 18:4(6,9,12,15).

 
Table 1 summarizes the fatty acid compositions of individual classes of lipids. Cells grown at 38°C and 25°C were analyzed. All classes of lipids contained 16:0 and 18:2 as the major acids, although 16:0 accumulated to about 70% in SQDG. 18:0 and 18:1 were also abundant in PE, PC, and PI. 16:1(9), which is commonly found in lipids of cyanobacteria (Fig. 1D) (22, 46) and algae (17), was detected only in PC, at a very low level. The level of 3-trans-16:1 in PG (Fig. 1C) was significantly lower in C. merolae (ca. 5%) than in land plants and algae analyzed to date (15 to 40%). Effects of growth temperature were noted for the fatty acid compositions of MGDG, PE, PC, and PI. In MGDG, the level of 18:2 was higher, while that of 16:0 was lower, at 25°C than at 38°C. For the three classes of phospholipids analyzed, the level of 18:2 was higher, while that of 18:1 was lower, at 25°C. The level of 18:0 was also lower in PE and PC. However, the growth temperature did not change the qualitative compositions of fatty acids.

Positional distribution of fatty acids. An analysis of the positional distribution of fatty acids in individual classes of lipids revealed marked differences between the lipids known as chloroplast lipids and other phospholipids (Table 2). In MGDG, DGDG, SQDG, and PG (chloroplast or plastid lipids), 16:0 was primarily bound to the C-2 position, whereas 18:2 was attached to the C-1 position. This is the prokaryotic type of distribution (1-C18-2-C16) found in the chloroplast lipids of plants and algae. In addition, a significant level of 18:2 was also found at the C-2 position in MGDG and DGDG. This points to the presence of the eukaryotic type of molecular species (1-C18-2-C18). SQDG contained 1-C16-2-C16 molecular species as well. The 3-trans-16:1 species was exclusively bound to the C-2 position of PG, as in the case of plants and algae. A totally different distribution was found in PE and PC. 16:0 and 18:0 were bound to the C-1 position, whereas the C-2 position was occupied mainly by 18:1 and 18:2. A low level of 18:2 also bound to the C-1 position. Therefore, PE and PC consisted mainly of the 1-saturated-2-unsaturated type of molecular species, as in many eukaryotes. These results suggest that the plastid lipids of C. merolae consist of prokaryotic and eukaryotic molecular species, as occurs in plants.


View this table:
[in this window]
[in a new window]

  		TABLE 2. Positional distribution of fatty acids in major classes of lipids

 
Genes involved in the synthesis of lipids and fatty acids. We then searched the genomic data of C. merolae to infer genes involved in the synthesis of fatty acids and lipids (Table 3). All common genes for the synthesis of long-chain saturated fatty acids were detected. Two types of acetyl-coenzyme A (acetyl-CoA) carboxylases, namely, a prokaryotic multisubunit enzyme of the plastid and a multifunctional cytoplasmic enzyme, were found. The plastid localization of the nuclear accC gene (CMS299C) product was confirmed by GFP experiments (Table 3; see Fig. S12 in the supplemental material). Two copies of the condensing enzymes, CMM286C and CML329C, were detected, but the former was confirmed as the plastid enzyme (see Fig. S12 in the supplemental material). Only three genes for putative desaturases were found. The gene for the stearoyl acyl carrier protein (ACP) desaturase, which is essential in producing oleate in flowering plants, was not detected. According to the Gclust database (data set CZ35, cluster 4024), putative genes for this enzyme were detected in the green alga Chlamydomonas reinhardtii (36) and the diatom Thalassiosira pseudonana (5).


View this table:
[in this window]
[in a new window]

  		TABLE 3. Genes for synthesis of lipids in the C. merolae genome

 
Genes encoding the enzymes involved in the synthesis of complex lipids were also identified by homology searches (Table 3). The eukaryotic MGDG synthase (encoded by CMI271C) (9) was found, and its intracellular localization was confirmed with a GFP fusion protein (Table 3; see Fig. S12 in the supplemental material). Curiously, a homolog of the Synechocystis enzyme Sll1377, which was recently found to be the enzyme involved in the synthesis of monoglucosyl diacylglycerol (GlcDG) (6), was also detected (CMT267C) (Gclust data set CZ20x0, cluster 2792). However, we could not detect a spot of putative GlcDG migrating slightly faster than MGDG in TLC after a short (10 min) pulse-labeling period (data not shown), which normally can detect a significant amount of GlcDG (equivalent to the amount of MGDG) in Anabaena (47). The function of CMT267C therefore still has to be determined.

Another point is the lack of a plant-type DGDG synthesis enzyme (DGD1/DGD2). No homolog of DGD1/2 (Gclust data set CZ35, cluster 5665) has been detected in cyanobacteria. Until now, a cyanobacterial DGDG synthesis enzyme has not been reported. This raises the possibility that C. merolae has a cyanobacterial DGDG synthesis enzyme (see Discussion).

Cardiolipin was not detected (see above), but genomic data suggested the presence of an enzyme for cardiolipin biosynthesis (CMN196C) (Gclust data set CZ20x0, cluster 4414). The family of cluster 4414 was originally annotated as a phosphatidylglycerophosphate synthase, but Katayama et al. (21) showed that the Arabidopsis homolog is involved in the synthesis of cardiolipin. To increase the sensitivity of detection of phospholipids, C. merolae cells were incubated with [32P]phosphate, and the lipids were analyzed by two-dimensional TLC (results not shown). However, the putative spot of cardiolipin was still obscure, and we cannot definitively confirm the presence of this lipid at a very low level.

The genomic analysis suggested that C. merolae has the capability of synthesizing some sterols (Table 3), although this is not the main topic of the present study. In Table 3, lanosterol synthase is listed because of its homology, but the exact specificity of the enzyme must be determined experimentally. Sterol methyltransferases with unknown specificity are also predicted (not listed in Table 3).

Phylogenetic analysis of desaturases. To obtain further information on the biosynthesis of fatty acids, a phylogenetic analysis of desaturases was performed (Fig. 2A). An uncompressed version of the identical tree is available upon request. The {Delta}9 desaturases are divergent from the {Delta}12 and {omega}3 desaturases, and all of these groups of desaturases, though still significantly homologous, diverged from each other before the separation of prokaryotes and eukaryotes (Fig. 2A). Other types of desaturases, such as {Delta}6 desaturases, were also included in the published large phylogenetic tree of desaturases (54), but they are too divergent to allow construction of a reliable tree. Among the acyl-CoA {Delta}9 desaturases, one group of enzymes have an extra cytochrome b5 domain (29, 33). One of the C. merolae enzymes (CMM045C) belongs to this type (20). Acyl lipid {Delta}9 desaturases are typically found in cyanobacteria (DesC), and homologs are also found in plants, which are known to function as {Delta}7 desaturases acting on MGDG (19). Cyanobacterial DesC and plant DesC-like proteins are sister groups (Fig. 2A). This suggests that the DesC-like enzymes in plants originated from the cyanobacterial endosymbiont. However, the second {Delta}9 desaturase (CMJ201C) in C. merolae is outside the DesC group and close to bacterial enzymes.


Figure 2
View larger version (37K):
[in this window]
[in a new window]

  		FIG. 2. Phylogenetic analysis of putative {Delta}12 desaturase in C. merolae (Cme). (A) Compressed phylogenetic tree of {Delta}9, {Delta}12, and {omega}3 desaturases. This tree was obtained by the neighbor-joining method, using MEGA 2 software. A full tree and sequence information are available from the authors upon request. Each number at the branch points indicates a bootstrap confidence level. (B) Detailed analysis of phylogenetic relationships of desaturases in cluster IV by neighbor-joining and maximum parsimony methods. (C) Best topology of cluster IV by the maximum likelihood method. (D) Alignment of the N-terminal regions of desaturases in cluster IV. Putative transit peptides for targeting to the ER and plastids are shown. The candidate initiation codons that were tested for the experiment shown in Fig. 3 are shown.

 
The phylogeny of the {Delta}12 (and {omega}3) desaturases is complicated (Fig. 2A). The cluster that diverges from the root includes bacterial and some marine cyanobacterial enzymes of unknown specificity (cluster I). Cluster II includes cyanobacterial DesA and plant FAD6 localized in the chloroplast. Another large cluster includes plant FAD2, which is localized in the ER (cluster IV). The {Delta}12 enzymes in marine cyanobacteria, C. merolae, and Phaeodactylum tricornutum (diatom) are related to the FAD2 group. The desaturases of nematodes (cluster III) diverge from both cluster II and cluster IV. Interestingly, {omega}3 desaturases (except the nematode enzyme) diverge from cluster IV of {Delta}12 desaturases. This is supported by a high bootstrap confidence level (89%). The cyanobacterial DesB protein was identified as the origin of the plant {omega}3 desaturases, including both chloroplast and ER isozymes.

These results show close relationships of cyanobacterial DesA and DesB with FAD6 and FAD7 in chloroplasts of plants and green algae (green lineage), respectively. Among cyanobacteria, marine species, such as Prochlorococcus marinus, have a cluster IV enzyme but no DesA or DesB. These enzymes were probably acquired by horizontal gene transfer, and this result should not be considered evidence that the C. merolae {Delta}12 desaturase originated from marine cyanobacteria. This is in clear contrast with {Delta}9 desaturases, for which both marine and freshwater species of cyanobacteria have orthologous DesC proteins.

These results suggest that the desaturases of C. merolae are unrelated to the enzymes of the cyanobacteria and the green lineage. Assuming the monophyletic origin of the red and green lineages (27, 28, 42), this indicates that the red alga does not keep cyanobacterial desaturases and retains the desaturases that existed before the cyanobacterial endosymbiosis. In addition, the {Delta}12 desaturases of the diatom P. tricornutum, PtFAD2 and PtFAD6, also clustered with the {Delta}12 desaturases of C. merolae and marine cyanobacteria (Fig. 2B and C). Neighbor joining (Fig. 2B) and maximum likelihood (Fig. 2C) as well as maximum parsimony (Fig. 2B) did not give a consistent relationship of these desaturases within this subcluster. PtFAD2 and PtFAD6 are known to be targeted to the ER and plastids, respectively (14).

Localization of {Delta}12 desaturase. The N terminus of PtFAD2 is similar to the N terminus of other ER-localized FAD2 proteins of plants and fungi (Fig. 2D), suggesting the presence of similar signal sequences. However, the {Delta}12 desaturase of C. merolae has an N-terminal extension, which is partially similar to the N-terminal extension of PtFAD6. Here we show the entire N-terminal sequence of the putative {Delta}12 desaturase (CMK291C). However, in the current version of annotation given in the Cyanidioschyzon website (http://merolae.biol.s.u-tokyo.ac.jp/), the CMK291C sequence begins from the second methionine. There is an in-frame upstream methionine codon, which could act as the initiation codon, and we used the entire sequence for the analysis in the present study. Intracellular localization of the putative {Delta}12 desaturase of C. merolae (CMK291C) was examined using GFP fusion constructs (Fig. 3). The polypeptide starting from Met1 was targeted to mitochondria, the polypeptide starting from Met2 was targeted to the ER, and the polypeptide beginning from Met3 showed no clear localization. It is interesting that the fluorescence of the Met2 construct is localized to the ER as well as to the membranes surrounding the nucleus (ER and nuclear envelope). It is therefore reasonable that the desaturase is translated from the second methionine, as described in the current database, and targeted to the ER.


Figure 3
View larger version (102K):
[in this window]
[in a new window]

  		FIG. 3. Targeting of putative {Delta}12 desaturase. There are three methionine codons that could act as the initiation codon in the putative {Delta}12 desaturase gene (CMK291C), as shown in Fig. 2D. The {Delta}12 sequences starting from the first, second, and third methionine codons were fused with the GFP gene and introduced into the onion epidermis by particle bombardment. Fluorescence of GFP and Nomarski differential interference images are shown. The control (GFP alone) is shown in Fig. S12 in the supplemental material. Tungsten particles are visible within the cells as small black patches.

 
Labeling of lipids. The computational analysis suggested that there is no desaturase in the plastid, but our lipid analysis showed that typical plastid lipids containing 18:2, such as MGDG, exist in this red alga. To solve this problem, we analyzed the flow of lipid synthesis within the cell by pulse labeling with radioactive acetate, followed by a chase period (Fig. 4). About 40% of added radiocarbon was incorporated into the total lipid fraction. The results of two-dimensional TLC indicated that PC and MGDG were the major labeled lipids. Interestingly, DGDG was not labeled efficiently after the 1-h labeling period, but it was densely labeled after the 20-h chase period. The radioactivity in PC decreased steadily during the chase period, while the radioactivity in MGDG increased. These results are consistent with the flow of carbon from PC to MGDG and then to DGDG, as documented for higher plants (10).


Figure 4
View larger version (16K):
[in this window]
[in a new window]

  		FIG. 4. Labeling of polar lipids with radioactive acetate. C. merolae cells were incubated with [14C]acetate for 1 h (A) and then chased for 20 h (B). Lipids were separated by two-dimensional TLC, and then the radioactivity of each lipid spot was counted (C). The radioactivity experiments were repeated three times, but representative results are shown throughout the paper for consistency of data.

 
Figure 5 shows the results of fatty acid analysis using a recently developed reversed-phase argentation TLC method. In the total lipid fraction, desaturation of fatty acids was clearly detected. In MGDG, however, only 16:0 and 18:2 were detected, with a decrease in 16:0 and an increase in 18:2. No possible intermediate, such as 18:0, 18:2, or 16:1, was detected as a labeled acid. For DGDG, labeling in the fatty acids was noted after the 6-h chase period. 16:0 was labeled first, and after the 20-h chase period, 18:2 was also labeled. In PC, various fatty acids were labeled, and the changes were explained by desaturation, except for the initial high level of labeling of 18:2, which might suggest rapid turnover of a small pool of PC-bound 18:2.


Figure 5
View larger version (15K):
[in this window]
[in a new window]

  		FIG. 5. Analysis of radioactivity in fatty acids in the total lipids (A), MGDG (B), DGDG (C), and PC (D). Each lipid class was isolated by two-dimensional TLC and recovered from the gel. Each isolated lipid was then subjected to methanolysis. Fatty acid methyl esters were extracted, separated by reversed-phase argentation TLC, and then quantified. (E) Autoradiogram of reversed-phase argentation TLC analysis of fatty acids in the 18:2/16:0 molecular species of MGDG before (lane 0) and after (lane 20) the chase. The original autoradiograms for panels A to D are shown in Fig. S10 in the supplemental material.

 
Lipid molecular species were also analyzed (Fig. 6). In MGDG, 18:2/16:0 was the major labeled molecular species from the beginning. At later times, 18:2/18:2 was also labeled. The rapid labeling of the 18:2/16:0 species of MGDG was not found in similar labeling experiments with plants (18), algae (17, 38, 50), or cyanobacteria (47) and seemed quite strange. This molecular species was recovered from the plate, and the fatty acids were analyzed (Fig. 5E). It was clear that the initially labeled 18:2/16:0-MGDG was labeled only in its 16:0 part, not in its 18:2 part. After the chase, both fatty acids were labeled. The delay in 18:2 labeling can be explained by the large size of the pool that provides 18:2. This suggests that MGDG is synthesized from 16:0 and 18:2 but that the two acids are supplied from different compartments, namely, the rapidly labeled 16:0 is supplied within the plastids, whereas the slowly labeled 18:2 is supplied from outside the plastids, possibly from the ER. Labeling of DGDG molecular species was similar to that for MGDG. In PC, various molecular species were labeled, and the radioactivity shifted from less unsaturated to more unsaturated species. This suggests that the desaturation of fatty acids occurs mainly on PC, as in other eukaryotes (10, 17).


Figure 6
View larger version (19K):
[in this window]
[in a new window]

  		FIG. 6. Analysis of radioactivity in various molecular species of MGDG (A), DGDG (B), and PC (C). Each lipid class was isolated by two-dimensional TLC. The molecular species were separated by argentation TLC and then quantified. The original autoradiograms for panels A to D are shown in Fig. S11 in the supplemental material.

 
The initial molecular species of galactolipid synthesis was also confirmed by incorporation of UDP-[U-14C]galactose, using isolated plastids (Fig. 7). 18:2/16:0 and 18:2/18:2 were the major labeled molecular species in both classes of lipids, although their exact proportions were not similar.


Figure 7
View larger version (66K):
[in this window]
[in a new window]

  		FIG. 7. Incorporation of radioactive galactose into various molecular species of MGDG (A) and DGDG (B) in isolated chloroplasts of C. merolae.

 

   DISCUSSION
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
Lack of trienoic acids. The results presented above indicate that C. merolae does not contain polyunsaturated acids with more than three double bonds. This is unusual for a photosynthetic eukaryote. Among the eukaryotes, Saccharomyces cerevisiae and some species of yeasts are the only organisms that possess only saturated and monounsaturated acids (15). Photosynthetic organisms in general have polyunsaturated acids such as 18:3, 20:4, or 22:6. The only known photosynthetic organisms that lack polyunsaturated acids are Synechococcus sp. strain PCC 6301 (formerly called Anacystis nidulans [49]) and its close relative PCC 7942 as well as some unicellular species of cyanobacteria (22). Polyunsaturated acids such as 18:3 are known to be important for two major reasons. First, 18:3 and 20:4 are precursors to physiologically active signaling compounds, such as jasmonate and prostaglandins in plants and animals, respectively. The lack of 18:3 implies that jasmonate might not act as a signal in this microalga. Second, 18:3 is required for maintaining photosynthetic activities at low and high temperatures. At low temperatures, this acid keeps the fluidity of membrane lipids to increase the tolerance to chilling and freezing (37). In addition, 18:3 is known to increase tolerance to high temperatures in plants (32). This does not mean that C. merolae does not have a tolerance to both cold and high temperatures. In fact, this alga can grow in a wide range of temperatures, from 25 to 50°C.

Pathway of lipid and fatty acid biosynthesis in C. merolae. C. merolae contains common glycerolipids that are usually found in plants and algae. Genomic analysis supports the hypothesis that this alga possesses a standard pathway of biosynthesis of these glycerolipids, except for DGDG (see Fig. S9 in the supplemental material). The biosynthesis of DGDG in plants is known to be catalyzed by the products of the DGD1/DGD2 genes. However, C. merolae does not have a homolog of these plant-type galactosyltransferases. A survey of the Gclust database (data set CZ20x0, cluster 2825) identified a putative glycosyltransferase (Ycf82) shared by cyanobacteria and the C. merolae plastid genome. The disruption of the Synechocystis homolog (slr1508) resulted in a lack of DGDG (I. Sakurai, N. Mizusawa, H. Wada, and N. Sato, unpublished data). This gene is also being analyzed in different laboratories, and we hope that this is the structural gene for the cyanobacterial and red algal DGDG synthesis enzyme. We also note that further studies are needed to obtain a conclusion about the presence of cardiolipin in C. merolae.

Biochemical analysis clearly indicated that C. merolae can synthesize saturated fatty acids and mono- and diunsaturated fatty acids. The gene involved in the synthesis of {Delta}3-trans-16:1, which is a typical fatty acid present at the C-2 position of PG in photosynthetic eukaryotes, is still unknown and is not a subject of the current discussion. The genomic analysis indicated that the pathway of fatty acid desaturation in this alga must be very different from that in flowering plants. Although fatty acid biosynthesis certainly occurs only in the plastids in C. merolae, as evidenced by the localization experiments using GFP (Table 3; see Fig. S12 in the supplemental material), the synthesis of oleate occurs in the ER and in the form of both acyl-CoA and acyl lipid (Table 3). This is fundamentally different from the {Delta}9 desaturation in flowering plants, in which oleate is produced only by the stearoyl ACP desaturase in the chloroplast, an enzyme that catalyzes desaturation by a different mechanism from that of acyl lipid desaturases (53). The genes encoding {Delta}7 acyl lipid desaturases have also been reported for plants, with some of their products acting as MGDG desaturases (19).

The second desaturation, namely, {Delta}12 desaturation, is catalyzed by the only {Delta}12 acyl lipid desaturase, which is likely to be localized in the ER (Fig. 3). However, MGDG, DGDG, and PG consist mainly of 1-(18:2)-2-(16:0) species, with small amounts of 1-(18:2)-2-(18:2) molecular species (Table 2). This indicates that the molecular species of these plastid lipids are synthesized by the coupled supply of 16:0 within the plastid and 18:2 from the outside. The role of the ER in acyl group desaturation was suggested by the results of tracer experiments with the cryptophyte Cryptomonas sp. (39), a descendant of secondary red algal endosymbiosis. In Cryptomonas, the radiolabel was initially incorporated into PC and then moved to MGDG. This and other results indicate that PC is the major substrate of desaturation in Cryptomonas. An essentially similar flow of carbon from PC to MGDG was found in C. merolae (Fig. 4).

Origins of desaturases. The desaturases in plants and algae have two different origins, namely, either the eukaryotic host or the cyanobacterial endosymbiont. The diversification of {Delta}9 and {Delta}12 desaturases had already occurred before the creation of eukaryotes, but some cyanobacterial enzymes were also transferred to plants and algae during endosymbiosis. It is clear that the {omega}3 desaturases were generated from a {Delta}12 desaturase, but the exact organism in which {omega}3 desaturase was created was not identified. Fig. 2A clearly shows that the cyanobacterial {omega}3 desaturase was not created within cyanobacteria. This is a question to be answered in the future.

The phylogenetic analysis (Fig. 2) suggested a cyanobacterial origin of {omega}3 desaturases of plants. Both plastidic (FAD7 and FAD8) and microsomal (FAD3) {omega}3 desaturases are monophyletic and originated from cyanobacterial DesB. Likewise, the plastidic {Delta}12 desaturase FAD6 in plants originated from cyanobacterial DesA. The {Delta}12 desaturase of C. merolae is a sister to eukaryotic {Delta}12 desaturases (FAD2) localized in the ER. Therefore, C. merolae completely lacks the desaturase genes of cyanobacterial origin, namely, desA, desB, FAD3, FAD6, FAD7, and FAD8. It is not yet clear if this is due to the selective loss of cyanobacterial enzymes in the lineage of primitive rhodophytes after endosymbiosis.

In cyanobacteria, the conversion of 18:0 to 18:1 occurs on acyl lipids by the action of DesC (Fig. 2). Neither DesC nor stearoyl ACP desaturase is present in C. merolae. Desaturation of 18:0 to 18:1 therefore proceeds by the two {Delta}9 desaturases, which have no direct relationship with {Delta}9 desaturases of plants or cyanobacteria (Fig. 2).

The similarity of the green lineage and the cyanobacteria with respect to desaturases is in clear contrast with the situation for the genomic machinery in the plastid (41, 42, 52). Various DNA-binding proteins of cyanobacterial origin are retained in the red lineage, whereas the genomic machinery acquired eukaryotic components in the green lineage. The genealogy of the enzymes of MGDG and DGDG synthesis supports the association of red algae and cyanobacteria. These observations shed light on the multiple or mosaic origins of lipid-related enzymes in the red algae.

Coupled pathway of MGDG synthesis. All of the results of tracer experiments clearly indicate that MGDG is synthesized from 16:0 and 18:2 within plastids. The DG moiety of MGDG is either 18:2/16:0 or 18:2/18:2 from the beginning (Fig. 7). 16:0 can be supplied within the plastids, whereas 18:2 cannot be supplied within the plastids because of the lack of desaturases within the plastids. It is most likely that PC in the ER is the site of desaturation, because all intermediate fatty acids were detected in the PC. A summary of galactolipid biosynthesis in C. merolae is illustrated in Fig. 8. Biosynthesis of MGDG therefore requires a supply of extraplastidic 18:2. This is the most interesting characteristic of C. merolae. We call this the coupled pathway, as opposed to the prokaryotic and eukaryotic pathways that were explained in the introduction. The coupled pathway of MGDG synthesis is a result of the total lack of desaturases of cyanobacterial origin and of stearoyl ACP desaturase.


Figure 8
View larger version (33K):
[in this window]
[in a new window]

  		FIG. 8. Comparison of pathways of galactolipid biosynthesis in C. merolae and flowering plants. In flowering plants, each of the prokaryotic (within the plastid) and eukaryotic (via the ER) pathways can produce unsaturated galactolipids, and the proportion of each pathway is different in different plants. In C. merolae, simultaneous functioning of both pathways is absolutely required for the synthesis of MGDG. The synthesis of MGDG is catalyzed by the plant-type enzyme. Although a homolog of cyanobacterial glucosyltransferase (Sll1377) was detected in the genome, there is no evidence for the production of GlcDG. The plant-type enzyme for the synthesis of DGDG does not exist in C. merolae. We suspect that another enzyme is used in cyanobacteria and C. merolae for the synthesis of DGDG.

 
In red algae other than those in the Cyanidiales (17, 35), C18 unsaturated fatty acids are not abundant. In Porphyra yezoensis, for example, C18 polyunsaturated acids amount to <1% of the galactolipids (4). This can be explained if, as in C. merolae, the synthesis of unsaturated fatty acids takes place mostly in the ER rather than in the plastids. The elongation of unsaturated fatty acids is also active in the ER, which results in vast production of C20 or C22 unsaturated fatty acids that characterize non-Cyanidiales red algal galactolipids. Although we still do not know if red algae in general lack stearoyl ACP desaturase, this enzyme could be an interesting switch that changes the metabolic flow of fatty acids in red algae.


   ACKNOWLEDGMENTS
 
This work was supported in part by grants-in-aid from JSPS (no. 13440234, 15370017, and 18017005), a grant-in-aid for priority areas from MEXT (Genome Biology), a grant-in-aid for creative scientific research (16GS0304), and a grant from the Takeda Science Foundation.


   FOOTNOTES
 
* Corresponding author. Mailing address: Department of Life Sciences, Graduate School of Arts and Sciences, University of Tokyo, 3-8-1 Komaba, Meguro-ku, Tokyo 153-8902, Japan. Phone: 81-3-54546631. Fax: 81-3-54546998. E-mail: naokisat{at}bio.c.u-tokyo.ac.jp Back

{triangledown} Published ahead of print on 6 April 2007. Back

{dagger} Supplemental material for this article may be found at http://ec.asm.org/. Back


   REFERENCES
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Adachi, J., and M. Hasegawa. 1992. Amino acid substitution of proteins coded for in mitochondrial DNA during mammalian evolution. Jpn. J. Genet. 67:187-197.[CrossRef][Medline]
  2. Allen, M. B. 1959. Studies with Cyanidium caldarium, an anomalously pigmented chlorophyte. Arch. Microbiol. 32:270-277.
  3. Altschul, S. F., T. L. Madden, A. A. Schäfer, J. Zhang, Z. Zhang, W. Miller, and D. J. Lipman. 1997. Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Res. 25:3389-3402.[Abstract/Free Full Text]
  4. Araki, S., T. Sakurai, T. Omata, A. Kawaguchi, and N. Murata. 1986. Lipid and fatty acid composition in the red alga Porphyra yezoensis. Jpn. J. Phycol. 34:94-100.
  5. Armbrust, E. V., J. A. Berges, C. Bowler, B. R. Green, D. Martinez, N. H. Putnam, S. Zhou, A. E. Allen, K. E. Apt, M. Bechner, M. A. Brzezinski, B. K. Chaal, A. Chiovitti, A. K. Davis, M. S. Demarest, J. C. Detter, T. Glavina, D. Goodstein, M. Z. Hadi, U. Hellsten, M. Hildebrand, B. D. Jenkins, J. Jurka, V. V. Kapitonov, N. Kroger, W. W. Lau, T. W. Lane, F. W. Larimer, J. C. Lippmeier, S. Lucas, M. Medina, A. Montsant, M. Obornik, M. S. Parker, B. Palenik, G. J. Pazour, P. M. Richardson, T. A. Rynearson, M. A. Saito, D. C. Schwartz, K. Thamatrakoln, K. Valentin, A. Vardi, F. P. Wilkerson, and D. S. Rokhsar. 2004. The genome of the diatom Thalassiosira pseudonana: ecology, evolution, and metabolism. Science 306:79-86.[Abstract/Free Full Text]
  6. Awai, K., T. Kakimoto, C. Awai, T. Kaneko, Y. Nakamura, K. Takamiya, H. Wada, and H. Ohta. 2006. Comparative genomic analysis revealed a gene for monoglucosyldiacylglycerol synthase, an enzyme for photosynthetic membrane lipid synthesis in cyanobacteria. Plant Physiol. 141:1120-1127.[Abstract/Free Full Text]
  7. Beisson, F., A. J. K. Koo, S. Ruuska, J. Schwender, M. Pollard, J. J. Thelen, T. Paddock, J. J. Salas, L. Savage, A. Milcamps, V. B. Mhaske, Y. Cho, and J. B. Ohlrogge. 2003. Arabidopsis genes involved in acyl lipid metabolism. A 2003 census of the candidates, a study of the distribution of expressed sequence tags in organs, and a web-based database. Plant Physiol. 132:681-697.[Abstract/Free Full Text]
  8. Bligh, E. G., and W. J. Dyer. 1959. A rapid method of total lipid extraction and purification. Can. J. Biochem. Physiol. 37:911-917.[Medline]
  9. Botte, C., C. Jeanneau, L. Snajdrova, O. Bastien, A. Imberty, C. Breton, and E. Maréchal. 2005. Molecular modeling and site-directed mutagenesis of plant chloroplast monogalactosyldiacylglycerol synthase reveal critical residues for activity. J. Biol. Chem. 280:34691-34701.[Abstract/Free Full Text]
  10. Browse, J., and C. Somerville. 1991. Glycerolipid synthesis: biochemistry and regulation. Annu. Rev. Plant Physiol. Plant Mol. Biol. 42:467-506.[CrossRef]
  11. Cavalier-Smith, T. 2003. Genomic reduction and evolution of novel genetic membranes and protein-targeting machinery in eukaryote-eukaryote chimaeras (meta-algae). Philos. Trans. R. Soc. Lond. B 358:109-134.[CrossRef][Medline]
  12. Chenna, R., H. Sugawara, T. Koik, R. Lopez, T. J. Gibson, D. G. Higgins, and J. D. Thompson. 2003. Multiple sequence alignment with the Clustal series of programs. Nucleic Acids Res. 31:3497-3500.[Abstract/Free Full Text]
  13. Chiu, W.-I., Y. Niwa, W. Zeng, T. Hirose, H. Kobayashi, and J. Sheen. 1996. Engineered GFP as vital reporter in plants. Curr. Biol. 6:325-330.[CrossRef][Medline]
  14. Domergue, F., P. Spiekermann, J. Lerchl, C. Beckmann, O. Kilian, P. G. Kroth, W. Boland, U. Zähringer, and E. Heinz. 2003. New insight into Phaeodactylum tricornutum fatty acid metabolism. Cloning and functional characterization of plastidial and microsomal delta12-fatty acid desaturases. Plant Physiol. 131:1648-1660.[Abstract/Free Full Text]
  15. Erwin, J. A. 1973. Comparative biochemistry of fatty acids in eukaryotic microorganisms, p. 41-143. In J. A. Erwin (ed.), Lipids and biomembranes of eukaryotic microorganisms. Academic Press, New York, NY.
  16. Fischer, W., E. Heinz, and M. Zeus. 1973. The suitability of lipase from Rhizopus arrhizus delemar for analysis of fatty acid distribution in dihexosyl diglycerides, phospholipids and plant sulfolipids. Hoppe-Seyler's Z. Physiol. Chem. 354:1115-1123.
  17. Harwood, J. L., and A. L. Jones. 1989. Lipid metabolism in algae. Adv. Bot. Res. 16:1-53.
  18. Heemskerk, J. W. M., H. Schmidt, U. Hammer, and E. Heinz. 1991. Biosynthesis and desaturation of prokaryotic galactolipids in leaves and isolated chloroplasts from spinach. Plant Physiol. 96:144-152.[Abstract/Free Full Text]
  19. Heilmann, I., S. Mekhedov, B. King, J. Browse, and J. Shanklin. 2004. Identification of the Arabidopsis palmitoyl-monogalactosyldiacylglycerol {delta}7-desaturase gene FAD5, and effects of plastidial retargeting of Arabidopsis desaturases on the fad5 mutant phenotype. Plant Physiol. 136:4237-4245.[Abstract/Free Full Text]
  20. Itoh, R., K. Toda, H. Takahashi, H. Takano, and T. Kuroiwa. 1998. Delta-9 fatty acid desaturase gene containing a carboxyl-terminal cytochrome b5 domain from the red alga Cyanidioschyzon merolae. Curr. Genet. 33:165-170.[CrossRef][Medline]
  21. Katayama, K., I. Sakurai, and H. Wada. 2004. Identification of an Arabidopsis thaliana gene for cardiolipin synthase located in mitochondria. FEBS Lett. 577:193-198.[CrossRef][Medline]
  22. Kenyon, C. N., R. Rippka, and R. Y. Stanier. 1972. Fatty acid composition and physiological properties of some filamentous blue-green algae. Arch. Mikrobiol. 83:216-236.[CrossRef][Medline]
  23. Kleinschmidt, M. G., and V. A. McMahon. 1970. Effect of growth temperature on the lipid composition of Cyanidium caldarium. II. Glycolipid and phospholipid components. Plant Physiol. 46:290-293.[Abstract/Free Full Text]
  24. Kumar, S., K. Tamura, I. B. Jakobsen, and M. Nei. 2001. MEGA2: molecular evolutionary genetics analysis software. Bioinformatics 17:1244-1245.[Abstract/Free Full Text]
  25. Kuroiwa, T. 1998. The primitive red algae Cyanidium caldarium and Cyanidioschyzon merolae as model system for investigating the dividing apparatus of mitochondria and plastids. BioEssays 20:344-354.[CrossRef]
  26. Marquardt, T. C., and R. F. Wilson. 1998. An improved reversed-phase thin-layer chromatography method for separation of fatty acid methyl esters. J. Am. Oil Chem. Soc. 75:1889-1892.[CrossRef]
  27. Martin, W., T. Rujan, E. Richly, A. Hansen, S. Cornelsen, T. Lins, D. Leister, B. Stoebe, M. Hasegawa, and D. Penny. 2002. Evolutionary analysis of Arabidopsis, cyanobacterial, and chloroplast genomes reveals plastid phylogeny and thousands of cyanobacterial genes in the nucleus. Proc. Natl. Acad. Sci. USA 99:12246-12251.[Abstract/Free Full Text]
  28. Matsuzaki, M., O. Misumi, T. Shin-i, S. Maruyama, M. Takahara, S. Miyagishima, T. Mori, K. Nishida, F. Yagisawa, K. Nishida, Y. Yoshida, Y. Nishimura, S. Nakao, T. Kobayashi, Y. Momoyama, T. Higashiyama, A. Minoda, M. Sano, H. Nomoto, K. Oishi, H. Hayashi, F. Ohta, S. Nishizaka, S. Haga, S. Miura, T. Morishita, Y. Kabeya, K. Terasawa, Y. Suzuki, Y. Ishii, S. Asakawa, H. Takano, N. Ohta, H. Kuroiwa, K. Tanaka, N. Shimizu, S. Sugano, N. Sato, H. Nozaki, N. Ogasawara, Y. Kohara, and T. Kuroiwa. 2004. Genome sequence of the ultrasmall unicellular red alga Cyanidioschyzon merolae 10D. Nature 428:653-657.[CrossRef][Medline]
  29. Mitchell, A. G., and C. E. Martin. 1995. A novel cytochrome b5-like domain is linked to the carboxyl terminus of the Saccharomyces cerevisiae {Delta}9 fatty acid desaturase. J. Biol. Chem. 270:29766-29772.[Abstract/Free Full Text]
  30. Moreira, D., H. Le Guyader, and H. Philippe. 2000. The origin of red algae and the evolution of chloroplasts. Nature 405:69-72.[CrossRef][Medline]
  31. Moretti, A., and R. Nazzaro. 1980. Fatty acids in thermoacidophilic algae. Delpinoa 21:3-11.
  32. Murakami, Y., M. Tsuyama, Y. Kobayashi, H. Kodama, and K. Iba. 2000. Trienoic fatty acids and plant tolerance of high temperature. Science 287:476-479.[Abstract/Free Full Text]
  33. Napier, J. A., O. Sayanova, P. Sperling, and E. Heinz. 1999. A growing family of cytochrome b5-domain fusion proteins. Trends Plant Sci. 4:2-4.[Medline]
  34. Nozaki, H., M. Matsuzaki, M. Takahara, O. Misumi, H. Kuroiwa, M. Hasegawa, T. Shin-i, Y. Kohara, N. Ogasawara, and T. Kuroiwa. 2003. Phylogeny of plastids based on cladistic analysis of gene loss inferred from complete plastid genome sequences. J. Mol. Evol. 56:485-497.[CrossRef][Medline]
  35. Nyberg, H., and K. Koskimies-Soininen. 1984. The glycolipid fatty acids of Porphyridium purpureum cultured in the presence of detergents. Phytochemistry 23:751-757.[CrossRef]
  36. Riekhof, W. R., B. B. Sears, and C. Benning. 2005. Annotation of genes involved in glycerolipid biosynthesis in Chlamydomonas reinhardtii: discovery of the betaine lipid synthase BTA1Cr. Eukaryot. Cell 4:242-252.[Abstract/Free Full Text]
  37. Routaboul, J.-M., S. F. Fischer, and J. Browse. 2000. Trienoic fatty acids are required to maintain chloroplast function at low temperatures. Plant Physiol. 124:1697-1705.[Abstract/Free Full Text]
  38. Sato, N. 1991. Lipids in Cryptomonas CR-1. I. Occurrence of betaine lipids. Plant Cell Physiol. 32:819-825.[Abstract/Free Full Text]
  39. Sato, N. 1991. Lipids in Cryptomonas CR-1. II. Biosynthesis of betaine lipids and galactolipids. Plant Cell Physiol. 32:845-851.[Abstract/Free Full Text]
  40. Sato, N. 2000. SISEQ: manipulation of multiple sequence and large database files for common platforms. Bioinformatics 16:180-181.[Abstract/Free Full Text]
  41. Sato, N. 2001. Was the evolution of plastid genetic machinery discontinuous? Trends Plant Sci. 6:151-156.[CrossRef][Medline]
  42. Sato, N. 2006. Origin and evolution of plastids: genomic view on the unification and diversity of plastids, p. 75-102. In R. R. Wise and J. K. Hoober (ed.), The structure and function of plastids. Springer, Berlin, Germany.
  43. Sato, N. 2006. Gclust server: phylogenetic profiling with pre-defined organism sets. Genome Inform. 17:P156.
  44. Sato, N., and M. Furuya. 1984. The composition of lipids and fatty acids determined at various stages of haploid and diploid generations in the fern Adiantum capillus-veneris. Physiol. Plant 62:139-147.[CrossRef]
  45. Sato, N., M. Ishikawa, M. Fujiwara, and K. Sonoike. 2005. Mass identification of chloroplast proteins of endosymbiont origin by phylogenetic profiling based on organism-optimized homologous protein groups. Genome Inform. 16:56-68.[Medline]
  46. Sato, N., and N. Murata. 1980. Temperature shift-induced responses in lipids in the blue-green alga, Anabaena variabilis. The central role of diacylmonogalactosylglycerol in thermo-adaptation. Biochim. Biophys. Acta 619:353-366.[Medline]
  47. Sato, N., and N. Murata. 1982. Lipid biosynthesis in the blue-green alga, Anabaena variabilis. II. Fatty acids and lipid molecular species. Biochim. Biophys. Acta 710:279-289.
  48. Sato, N., and N. Murata. 1988. Membrane lipids. Methods Enzymol. 167:251-259.
  49. Sato, N., N. Murata, Y. Miura, and N. Ueta. 1979. Effect of growth temperature on lipid and fatty acid compositions in the blue-green algae, Anabaena variabilis and Anacystis nidulans. Biochim. Biophys. Acta 572:19-28.[Medline]
  50. Sato, N., Y. Nemoto, and M. Furuya. 1988. Lipids of Chattonella antiqua (Raphidophyceae). Plant Physiol. Biochem. 26:93-98.
  51. Sato, N., K. Sekine, Y. Kabeya, S. Ehira, M. Onuma, and N. Ohta. 2004. Discontinuous evolution of plastid genomic machinery: radical replacement of major DNA-binding proteins. Endocytobiosis Cell Res. 15:286-293.
  52. Sato, N., K. Terasawa, K. Miyajima, and Y. Kabeya. 2003. Organization, developmental dynamics and evolution of the plastid nucleoid. Int. Rev. Cytol. 232:217-262.[Medline]
  53. Shanklin, J., and E. B. Cahoon. 1998. Desaturation and related modification of fatty acids. Annu. Rev. Plant Physiol. Plant Mol. Biol. 49:611-641.[CrossRef][Medline]
  54. Sperling, P., P. Ternes, T. K. Zank, and E. Heinz. 2003. The evolution of desaturases. Prostaglandins Leukot. Essent. Fatty Acids 68:73-95.[CrossRef][Medline]
  55. Toda, K., H. Takano, S. Miyagishima, H. Kuroiwa, and T. Kuroiwa. 1998. Characterization of a chloroplast isoform of serine acetyltransferase from the thermo-acidiphilic red alga Cyanidioschyzon merolae. Biochim. Biophys. Acta 1403:72-84.[Medline]
  56. Yoon, H. S., J. D. Hackett, G. Pinto, and D. Bhattacharya. 2002. A single origin of the peridinin- and fucoxanthin-containing plastids in dinoflagellates through tertiary endosymbiosis. Proc. Natl. Acad. Sci. USA 99:15507-15512.[Abstract/Free Full Text]

Eukaryotic Cell, June 2007, p. 1006-1017, Vol. 6, No. 6
1535-9778/07/$08.00+0     doi:10.1128/EC.00393-06
Copyright © 2007, American Society for Microbiology. All Rights Reserved.



This article has been cited by other articles:


This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Supplemental material
Right arrow Other Versions of this Article:
EC.00393-06v1
6/6/1006    most recent
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow Copyright Information
Right arrow Books from ASM Press
Right arrow MicrobeWorld
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Sato, N.
Right arrow Articles by Moriyama, T.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation