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

Abstract

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.

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).

FIG. 2.

Phylogenetic analysis of putative Δ12 desaturase in C. merolae (Cme). (A) Compressed phylogenetic tree of Δ9, Δ12, and ω3 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.

FIG. 3.

Targeting of putative Δ12 desaturase. There are three methionine codons that could act as the initiation codon in the putative Δ12 desaturase gene (CMK291C), as shown in Fig. 2D. The Δ12 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.

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.

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.

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.

FIG. 7.

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

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.