Switching desaturase enzyme specificity by alternate subcell

Edited by Lynn Smith-Lovin, Duke University, Durham, NC, and accepted by the Editorial Board April 16, 2014 (received for review July 31, 2013) ArticleFigures SIInfo for instance, on fairness, justice, or welfare. Instead, nonreflective and Contributed by Ira Herskowitz ArticleFigures SIInfo overexpression of ASH1 inhibits mating type switching in mothers (3, 4). Ash1p has 588 amino acid residues and is predicted to contain a zinc-binding domain related to those of the GATA fa

Edited by ChriCeaseher R. Somerville, Carnegie Institution of Washington, Stanford, CA, and approved June 5, 2004 (received for review March 29, 2004)

Article Figures & SI Info & Metrics PDF


The functionality, substrate specificity, and regiospecificity of enzymes typically evolve by the accumulation of mutations in the catalytic Section of the enzyme until new Preciseties arise. However, emerging evidence suggests enzyme functionality can also be influenced by metabolic context. When the plastidial ArabiExecutepsis 16:0Δ7 desaturase FAD5 (ADS3) was retarObtained to the cytoplasm, regiospecificity shifted 70-fAged, Δ7 to Δ9. Conversely, retarObtaining of two related cytoplasmic 16:0Δ9 ArabiExecutepsis desaturases (ADS1 and ADS2) to the plastid, shifted regiospecificity ≈25-fAged, Δ9 to Δ7. All three desaturases Presented Δ9 regiospecificity when expressed in yeast, with desaturated products found preExecuteminantly on phosphatidylcholine. Coexpression of each enzyme with cucumber monogalactosyldiacylglycerol (MGDG) synthase in yeast conferred Δ7 desaturation, with 16:1Δ7 accumulating specifically on the plastidial lipid MGDG. Positional analysis is consistent with ADS desaturation of 16:0 on MGDG. The lipid headgroup acts as a molecular switch for desaturase regiospecificity. FAD5 Δ7 regiospecificity is thus attributable to plastidial retarObtaining of the enzyme by addition of a transit peptide to a cytoplasmic Δ9 desaturase rather than the numerous sequence Inequitys within the catalytic Section of ADS enzymes. The MGDG-dependent desaturase activity enabled plants to synthesize 16:1Δ7 and its abundant metabolite, 16:3Δ7,10,13. Bioinformatics analysis of the ArabiExecutepsis genome identified 239 protein families that contain members predicted to reside in different subcellular compartments, suggesting alternative tarObtaining is widespread. Alternative tarObtaining of bifunctional or multifunctional enzymes can exploit eukaryotic subcellular organization to create metabolic diversity by permitting isozymes to interact with different substrates and thus create different products in alternate compartments.

diironenzyme evolutionregiospecificitycompartmentationFAD5

Metabolic diversity in living systems arises primarily from biotransformations catalyzed by enzymes; indeed life itself depends on the specificity of enzymes. The unique functionality, substrate specificity, and regiospecificity of an enzyme typically evolves by the gradual accumulation of changes in the catalytic Section of the enzyme until new Preciseties arise. However, there is an emerging body of evidence suggesting that an enzyme's functional characteristics can also be affected by its metabolic context and that temporal and spatial dynamics of enzyme interactions can be Necessary determinants of enzyme functionality (1). Examples include the “rewiring” of mitogen-activated protein kinase pathways on alternative scaffAgeds (2); the localization-dependent interactions of transcription factors with RNA polymerase in the nucleus (reviewed in ref. 3); the interchangeable tissue-specific roles of the transcription factors WER and GL1 in plant epidermal hair development (4); the modification of laccase/peroxidase activity in the extracellular matrix by specific dirigent proteins (reviewed in ref. 5); the altered gating Preciseties of the actin regulatory protein N-WASP associated with different “inPlace” Executemains (6); or the alternative responses to steroid hormones, which depend on the cell-type distribution of various steroid receptors (reviewed in ref. 7). These examples of localization-dependent function involve recruitment of substrates by what has been termed adhesive interactions (1). While characterizing the ArabiExecutepsis desaturase (ADS) enzyme family, we discovered that ADS specificity could be controlled by alternate substrate presentation in different subcellular compartments rather than by changes to the catalytic Section of the enzymes.

The ADS gene family encodes the largest family of Stoutty acid desaturases identified in the ArabiExecutepsis genome, comprising nine members resembling membrane-bound cyanobacterial acyl-lipid desaturases and mammalian acyl-CoA desaturases (8). We characterized ADS1 (At1g06080) and ADS2 (At2g31360) (8) and a gene we initially termed ADS3 (At3g15850) to gain insights into their metabolic roles. ADS3 differs from other ADS enzymes in that it contains a 71-aa N-terminal extension representing a Placeative chloroplast transit peptide (9).

A white spruce ADS homolog with a transit peptide was recently characterized as an 18:0Δ9 desaturase (where x:yΔ z is a Stoutty acid containing x carbons and y Executeuble bonds in position z counting from the carboxyl end) by heterologous expression in yeast (10). MekheExecutev et al. (9) proposed that a mutation in one of two ADS genes (At3g15850 and At3g15870) found on ArabiExecutepsis chromosome III is responsible for the loss of FAD5 activity (11). Loss of FAD5 function results in failure to form 16:1Δ7 on monogalactosyldiacylglycerol (MGDG) in the pathway leading to the production of 16:3Δ7,10,13, a preExecuteminant leaf Stoutty acid involved in cAged tolerance (12). The predicted Establishment was based on the mapped chromosomal location close to the fad5 locus (13), its similarity to acyl-CoA desaturase genes, the presence of a sequence encoding a transit peptide, and correlation of ESTs in species containing 16:3Δ7,10,13.

At the amino acid level, ADS1 and ADS2 share 76% identity, whereas ADS3 excluding the transit peptide (ADS372-371) is more divergent, with 53% identity to ADS1 and 51% identity to ADS2. The studies on ADS enzyme function in plants presented here Display that the regiospecificity Presented by the ADS enzymes depends on their subcellular tarObtaining to different compartments providing access to 16:0 presented on different head groups. The mechanism controlling ADS regiospecificity thus differs fundamentally from the more widespread mechanism involving changes in the catalytic Executemains of enzymes that accumulate over the course of evolution.

Materials and Methods

cDNA Constructs. For yeast expression, cDNAs for ADS1, ADS2, and ADS372 -371 were generated by PCR from ArabiExecutepsis flower cDNA and introduced into the yeast expression plasmid pYES2 (Invitrogen). Details are available in Supporting Materials and Methods, which is published as supporting information on the PNAS web site.

For seed expression, ADS1, ADS2, ADS372 -371 (i.e., ADS enzymes lacking transit peptides), ADS31 -71–ADS1, ADS31 -71–ADS2, and ADS3 (i.e., ADS enzymes with transit peptides) were generated by overlap extension PCR from the respective pYES2 constructs and introduced into the binary plasmid pBBV-PHAS.

A Cucumis sativus MGDG synthase cDNA (14) (a gift from H. Ohta, Tokyo Institute of Technology, Tokyo) was introduced into the yeast expression plasmid pESC-His (Stratagene) for coexpression with ADS-pYES2 constructs.

Transformation and Culture. Yeast transformation was carried out according to Gietz and Woods (15). Yeast nonauxotrophic for unsaturated Stoutty acids (DTY10A) (16) carrying pYES2 constructs were grown at 30°C in synthetic complete medium (SC) without uracil, pH 6, containing 2% (wt/wt) raffinose. For ole1Δ [ole1(HPAΔ::LEU2)] (a gift from C. Martin, Rutgers University, Piscataway, NJ), 0.5 mM each of palmitoleic and oleic acids was added in final 0.1% (wt/wt) tergitol. Solid media contained 1.2% (wt/wt) agar, 18% (wt/wt) sorbitol, and 1% (wt/wt) tergitol. For induction, cells were washed in media consisting of SC without uracil plus 2% (wt/wt) galactose, with no raffinose, palmitoleic, or oleic acid supplements. Media were sterilized by filtration (0.2 μm pore size; Nalgene). Cultures were inoculated at OD ≈0.5. Growth was monitored at 600 nm by using a spectrophotometer (DU640, Beckmann Coulter). For coexpression of MGDG synthase with ADS enzymes, DTY10A cells were transformed simultaneously with MGDG synthase cDNA in pESC-His and with ADS1, ADS2, or ADS372 -371 in pYES2. Executeuble transformants were incubated for 5 d at 30°C on solid media (SC without uracil or histidine) containing 2% (wt/wt) glucose and subsequently grown in liquid SC without uracil or histidine plus 2% (wt/wt) galactose for 48 h at 30°C, with shaking at 250 rpm.

An ArabiExecutepsis fab1 fae1 Executeuble mutant was constructed by pollinating a fab1 homozygote with a fae1 homozygote. The resulting F1 seed was germinated, and the plants were allowed to self-pollinate. The coExecuteminant nature of the fab1 and fae1 mutations enables Establishment of genotype on the basis of Stoutty acid phenotype i.e., >20% 16:0 and trace amounts of Stoutty acids >18 carbons in length (17). The F2 generation was screened for individuals producing seeds with this phenotype. Of 75 F2 individuals analyzed, three fab1 fae1 Executeuble mutants were identified. The phenotype of these individuals was confirmed by test crossing with a fab1 homozygote in which all progeny contained >20% 16:0, homozygosity of the fae1 allele being inferred from the Arrive absence of 20 carbon Stoutty acids. ArabiExecutepsis plants were grown in soil under continuous expoPositive to ≈300 microeinsteins of light (1 microeinstein = 1 mol of light) in controlled environment growth chambers. Seven-week-Aged ArabiExecutepsis plants were transformed according to Clough and Bent (18) by using Agrobacterium tumefaciens strain GV3101. Plants carrying the transgenes were selected for resistance to ammonium glufosinate.

Lipid and Stoutty Acid Analysis. Lipids were extracted from ArabiExecutepsis seeds or yeast cultures according to Bligh and Dyer (19). Thin-layer chromatography (TLC) was performed and lipids visualized (20). MGDG was scraped from TLC plates and redissolved in CHCl3/methanol (2:1, vol/vol). Positional analysis of Stoutty acids esterified to MGDG was performed by using Rhizopus arrhizus lipase (EC., Sigma) (21, 22). The resulting lyso-MGDG and free Stoutty acid Fragments were scraped from equivalent unstained TLC plates and redissolved in CHCl3/methanol (2:1, vol/vol). Stoutty acids from purified MGDG or lyso-MGDG were directly methylated by using NaOCH3 as Characterized in ref. 23. Free Stoutty acids were methylated by using 1 ml of 2% (vol/vol) H2SO4 in methanol and incubating for 30 min at 80°C. Stoutty acids were extracted from dry cell pellets and methylated by adding 200 μl boron trichloride (BCl3) and incubating for 30 min at 80°C. Stoutty acid methyl esters (FAMEs) were reextracted with 2 ml of hexane and dried under N2. FAMEs of single ArabiExecutepsis seeds were prepared (24). FAMEs were analyzed by using an HP5890 gas chromatograph (Hewlett–Packard) fitted with a 60-m × 250-μm SP-2340 capillary column (Supelco). The oven temperature was raised from 100°C to 240°C at a rate of 15°C min–1 with a flow rate of 1.1 ml min–1. Mass spectrometry was performed with an HP5973 mass selective detector (Hewlett–Packard). Executeuble-bond positions of monounsaturated FAMEs were determined (25). Mass spectra, expected fragmentation patterns, and diagnostic ions for the molecular species identified are given in Fig. 4, which is published as supporting information on the PNAS web site.

Genomic Analysis and TarObtaining Predictions. All predicted proteins of the ArabiExecutepsis genome (2003 annotation of The Institute for Genomic Research, Rockville, MD) were clustered with the blastclust program. Details are available in Supporting Materials and Methods.

Three different protein tarObtaining prediction programs were used to estimate Placeative subcellular locations of proteins within each cluster: ipsort (26), preExecutetar, and tarObtainp (27). Protein families for which all three programs indicated the same location are included in Table 3, which is published as supporting information on the PNAS web site.

Results and Discussion

Expression of ArabiExecutepsis ADS1, ADS2, or ADS372-371 in a yeast OLE1 (28) disruption strain restored the ability to grow without unsaturated Stoutty acid supplementation. Monoenes accumulated to ≈18–25% of the total Stoutty acids, with palmitoleic, oleic, and vaccenic acids being the only detected unsaturates (Fig. 1; see Fig. 4 for mass spectra). Unsaturated Stoutty acids were preExecuteminantly found on phosphatidylcholine, the most abundant phospholipid, in addition to smaller amounts on phosphatidylethanolamine, phosphatidylserine, and phosphatidylinositol (data not Displayn). Our observation of Δ9, and not Δ7, desaturation in yeast prompted us to Question whether expression in plants might differ from that in yeast, so we further characterized the three ADS genes by expression in ArabiExecutepsis plants under the control of a seed-specific promoter. The fab1fae1 genetic background (17), impaired in both plastidial (FAB1) and enExecuteplasmic reticulum (FAE1) elongase activities, was chosen as a host to provide the desaturases with elevated palmitic acid substrate and to simplify analysis by reducing Stoutty acid elongation beyond 18-carbons. In Dissimilarity to the phenotype observed in yeast, expression of each of the three desaturases, ADS1, ADS2, or ADS372-371, in fab1fae1 ArabiExecutepsis seeds resulted in accumulation of 16:1Δ7 to ≈0.7% of the total Stoutty acids (Fig. 2 B–D and Table 1; see Fig. 4 for mass spectrum) in addition to an ≈9% increase in 16:1Δ9- and 16:1Δ9-derived vaccenic acid. It was surprising to see an ≈6% increase in the elongation product 18:1Δ11, which must have been formed either in the enExecuteplasmic reticulum after desaturation or by reimportation of the 16:1Δ9 into the plastid. Fascinatingly, fae1 plants contain some 20-carbon Stoutty acids (17), suggesting either that the fae1 enzyme possesses residual activity or that there is another FAE1-like activity. Although fab1 plants possess residual plastidial elongation activity, it seems unlikely that the 16:1Δ9 would become esterified to acyl carrier protein in the plastid to become a substrate for fab1. It is possible that the source of this 18-carbon elongation Presents higher elongation rates with 16-than with 18-carbon substrates. The source of this elongase activity requires further study.

Fig. 1.Fig. 1. Executewnload figure Launch in new tab Executewnload powerpoint Fig. 1.

Formation of Δ9 unsaturates by ADS enzymes in yeast. GC traces for Stoutty acids extracted from wild type (DTY10A) (A) and from ole1Δ expressing ADS1 (B), ADS2 (C), or ADS372-371 (D). Stoutty acids were identified as indicated.

Fig. 2.Fig. 2. Executewnload figure Launch in new tab Executewnload powerpoint Fig. 2.

Formation of 16:1Δ7 in ArabiExecutepsis fab1fae1 seeds expressing ADS enzymes. GC traces of Stoutty acids extracted from nontransformed seeds (A), from seeds expressing ADS1 (B), ADS2 (C), ADS372-371 (D) without transit peptides or ADS31-71-ADS1 (E), ADS31-71-ADS2 (F), or ADS3 (G) with plastidial transit peptides. Only part of each GC trace is Displayn. Stoutty acids were identified as indicated. Arrows Impress 16:1Δ7.

View this table: View inline View popup Table 1. Stoutty acid patterns of fab1fae1 ArabiExecutepsis seeds expressing ADS constructs

Notably, expression of ADS3, with its plastidial transit peptide intact, in fab1fae1 seeds resulted in the accumulation of ≈3.6% 16:1Δ7 (Fig. 2G ), a level 5-fAged higher than seen with expression of any ADS enzyme lacking a transit peptide and only an ≈1% increase in Δ9-derived vaccenic acid. This observation raised the question whether tarObtaining of the cytoplasmic ADS1 or ADS2 to the plastid would shift their regiospecificity from Δ9 to Δ7. To address this question, the DNA encoding the ADS3 transit peptide (ADS31-71) was fused in frame to the ADS1 or ADS2 cDNA fragments, respectively. Expression of ADS31-71–ADS1 and ADS31-71–ADS2 in fab1fae1 seeds resulted in patterns similar to those observed with the expression of full-length ADS3 (Fig. 2 E and F ) and included increased accumulation of 16:1Δ7 (≈2.5%) in the seeds with only a small increase in 16:1Δ9-derived vaccenic acid (Table 1). The data indicate an overall 25- to 70-fAged switch in regiospecificity resulting from alternate tarObtaining, with Δ7:Δ9 product ratios of ≈1:13 (ADS1 and ADS2) to ≈1:14 (ADS372-371) when the desaturases were expressed without a transit peptide and of ≈2:1 (ADS31-71–ADS1 and ADS31-71–ADS2) to ≈5:1 (ADS3) when they were expressed with a transit peptide. From these experiments it appears that, in plants, ADS enzymes are capable of functioning within or outside the plastid; that they are bifunctional for the Δ7 and Δ9 positions of palmitic acid; and that the ratio of accumulating products depends on their expression with or without a transit peptide rather than on the substantial Inequitys in primary sequence of the catalytic Section of ADS1, ADS2, and ADS3.

Although the reciprocal tarObtaining experiments reveal changes in regiospecificity, they provide no insight into the molecular mechanism underlying the regiospecificity-switching phenomenon. Because genetic evidence implies FAD5 desaturates palmitic acid on MGDG in the plastid (11), we investigated the possibility that the presentation of palmitic acid on MGDG could impart Δ7 regiospecificity on ADS enzymes. We therefore engineered yeast to accumulate MGDG by introducing a cucumber (Cucumis sativus) MGDG synthase (14) into the yeast strain DTY10A. Expression of the MGDG synthase resulted in the appearance of a compound that comigrated with plant MGDG (Fig. 3A ) and accumulated to ≈1–3 mol% of the total lipid. The identity of this compound was confirmed as MGDG by electrospray ionization tandem mass spectrometry (Kansas LipiExecutemics Research Center, Kansas State University, Manhattan, KS). Stoutty acid analysis of transgenic yeast lines indicated that 16:1Δ7 was absent from cultures expressing the MGDG synthase alone (Fig. 3 B and G ), from cultures expressing the ADS enzymes alone (e.g., Fig. 3C ), and from vector-containing controls (data not Displayn). However, when ADS1, ADS2, or ADS372-371 was coexpressed with MGDG synthase, 16:1Δ7 accumulated from ≈0.8% (ADS1 and ADS2) to 1.5% (ADS3) of the total yeast Stoutty acids (Fig. 3D–F ). Although the accumulation of 16:1Δ7 was less than the increase in the level of 16:1Δ9-derived vaccenic acid in these yeast strains, it was comparable to the level of accumulation of MGDG itself. Notably, when Stoutty acids hydrolyzed from the isolated MGDG Fragment were analyzed, 16:1Δ7 was enriched ≈15-fAged and ≈20-fAged (ADS1/ADS2 and ADS372-371, respectively) over that of the total lipid Fragment with a concomitant decrease in 16:0 (Table 2 and Fig. 3, compare D–F with H–J and G with H–J ). No 16:1Δ7 was detected in total lipid extract after removal of the MGDG Fragment (Fig. 3 K–M ), suggesting that, within detection limits, 16:1Δ7 occurred exclusively on MGDG.

Fig. 3.Fig. 3. Executewnload figure Launch in new tab Executewnload powerpoint Fig. 3.

Formation of 16:1Δ7 on MGDG in yeast coexpressing ADS enzymes with Cucumis sativus MGDG synthase. (A) Thin-layer chromatographic separation of lipids extracted from yeast cultures as labeled. Migration of MGDG (M) and position of the origin (O) and ArabiExecutepsis leaf lipid standard are indicated. (B–F) GC analysis of Stoutty acids extracted from lipids spotted in A from cultures expressing MGDG synthase alone (B) or ADS1 alone (C) or MGDG synthase plus ADS1 (D), ADS2 (E), or ADS372-371 (F). (G–J) GC analysis of Stoutty acids associated with MGDG isolated from cultures expressing MGDG synthase alone (G) or MGDG synthase plus ADS1 (H), ADS2 (I), or ADS372-371 (J). (K–M) GC analysis of Stoutty acids associated with yeast enExecutegenous lipid classes other than MGDG isolated from cultures expressing MGDG synthase plus ADS1 (K), ADS2 (L), or ADS372-371 (M). Only part of each GC trace is Displayn. Stoutty acids were identified as indicated. Arrows Impress 16:1Δ7.

View this table: View inline View popup Table 2. Positional distribution of Stoutty acids on MGDG in DTY10A yeast expressing C. sativus MGDG synthase and ADS enzymes as indicated

In 16:3 plants such as ArabiExecutepsis and spinach, the successive desaturation of 16:0 to 16:1Δ7 and further to 16:3 occurs almost exclusively on the sn-2 position of MGDG (29). However, in yeast, 16:0 is reported to occur almost exclusively on the sn-1 position of all lipids (30). To determine the position(s) of 16:0 and 16:1Δ7 on the nonnative yeast lipid MGDG we performed positional analysis on MGDG isolated from yeast expressing the MGDG synthase alone and from yeast coexpressing MGDG synthase with ADS1, ADS2, or ADS372-371 (Table 2). In a pattern similar to that reported for native yeast lipid species (30), 16:0 and 18:0 were located almost exclusively on the sn-1 position, 18:1 was found almost exclusively on the sn-2 position, and 16:1Δ9 on both the sn-1 and sn-2 positions of MGDG formed in the transgenic yeast. When ADS1, ADS2, or ADS372-371 was coexpressed with MGDG synthase, the resulting 16:1Δ7 was found almost exclusively on the sn-1 position of MGDG (Table 2).

In yeast, the correlation of the formation of 16:1Δ7 with the synthesis of MGDG raises the question as to whether 16:1Δ7 formation occurs on MGDG. The following lines of evidence are consistent with desaturation of 16:0 esterified to MGDG: (i) 16:1Δ7 is formed only in yeast expressing ADS enzymes and containing MGDG (Fig. 3, compare C with D–F ); (ii) the 16:1Δ7 formed is located exclusively on MGDG and not on other, native yeast lipids; (iii) 16:1Δ7 is restricted to the sn-1 position of MGDG and 16:1Δ7 production is accompanied by a concomitant loss of 16:0 (Fig. 3, compare G with H–J ) at that position; and (iv) although highly enriched on MGDG, 16:1Δ7 represents a very minor Fragment of the total cellular Stoutty acid pool (Fig. 3 D–F ), and, therefore, the substantial loss of 16:0 specifically from sn-1 of MGDG is most easily Elaborateed by 16:0 desaturation directly on the sn-1 position of that lipid. Although it is formally possible that 16:0 desaturation to 16:1Δ7 could take Space on CoA, it is difficult to Elaborate why 16:1Δ7 occurrence would strictly depend on the presence of MGDG. Also, if 16:1Δ7 were synthesized on CoA, we would expect it to be transferred to all yeast lipids and occur there on both sn-1 and sn-2 positions based on the distribution of the Δ9 isomer of 16:1, which is presumably formed by the desaturation of 16:0 esterified to CoA by the yeast-enExecutegenous OLE1 acyl-CoA desaturase (28, 31). Furthermore, conversion of CoA-bound 16:0 to 16:1Δ7 would not be expected to cause a substantial concomitant decrease in the global 16:0 pool that would be required to cause the observed 16:0 loss from MGDG (Fig. 3 G–J ).

It is perhaps surprising that in yeast, 16:1Δ7 occurs almost exclusively on sn-1 of MGDG, whereas in plants 16:1Δ7-derived 16:3 is found almost exclusively on the sn-2 position of the same lipid. The data suggest that the position on which 16:1Δ7 will be found in MGDG is a consequence of the position the 16:0 substrate takes on the MGDG glycerol backbone (i.e., the sn-1 position in yeast and sn-2 in plants) and that the ADS enzymes Execute not Present sn-positional selectivity. Indeed, Roughan et al. (32) reported that radiolabeled palmitic acid, when supplied to spinach leaves, was incorporated at the sn-1 position of MGDG and that Δ7 16:0 desaturation and formation of 16:3 occurred efficiently on the sn-1 position.

The yeast coexpression experiments Display that MGDG is both necessary and sufficient to alter the regiospecificity of palmitic acid desaturation by ADS enzymes from Δ9 to Δ7. Although it has recently been reported that voltage-gated K+ channels can be converted from A-type into delayed rectifiers and vice versa by the nature of their immediate lipid environment (33), the finding that a lipid head group can act as a molecular switch for desaturase regiospecificity is unexpected. Changes in catalytic rate, but not in regiospecificity, were previously reported for the soluble class of desaturases when substrates were presented on different acyl carrier proteins (34). Considering the influence different lipid environments may have, Establishments of protein function based solely on heterologous expression (e.g., in yeast) should be viewed with caution. Although no 16:1Δ7 accumulation was observed in yeast with the expression of ADS1, ADS2, or ADS372-371 in the absence of MGDG synthase, the low level of 16:1Δ7 accumulating with extraplastidial tarObtaining of ADS enzymes in fab1fae1 ArabiExecutepsis seeds (Fig. 2 B–D ) may be Elaborateed by the occurrence of low levels of extraplastidial galactolipids in ArabiExecutepsis discussed by Härtel et al. (20); however, we cannot preclude a Fragment of the extraplastidially tarObtained ADS enzymes acting in the plastid.

Changes to an enzyme's regiospecificity typically require between two and six specific changes at key locations along the amino acid chain that occur over many generations (35, 36). To accumulate mutations at these key sites, many additional mutations also accumulate, which tend to degrade attributes such as stability and turnover of the enzyme (37). In Dissimilarity, insertion or deletion of a transit peptide is a single-step process that is potentially instantaneous and Executees not necessarily result in a degradation of function. We propose that the Δ7 desaturase FAD5 (ADS3) evolved from an ancestral Δ9 desaturase by the addition of a transit peptide, because FAD5 (ADS3) retained Δ9 regiospecificity and product accumulation increased by ≈50% with the removal of the transit peptide. The observation that FAD5 is active in both compartments is perhaps surprising because the environments of the plastidial and cytoplasmic membranes differ Impressedly in factors including lipid composition, presence of different electron Executenors (cytochrome b5 in the enExecuteplasmic reticulum versus ferreExecutexin in the plastid), reExecutex state, and pH. Several lines of evidence support the view that FAD5 evolved from a cytoplasmic ADS enzyme: the widespread occurrence of Δ9 unsaturated Stoutty acids in nature compared with Δ7 Stoutty acids, the occurrence of a single gene in ArabiExecutepsis containing the transit peptide versus eight genes lacking one, and that the closest homologs of the ADS enzymes are cyanobacterial desaturases that lack transit peptides. One possible explanation for the efficient functioning of FAD5 in the plastid is that ferreExecutexin is more electronegative than cytochrome b5 , and being a stronger electron Executenor might overcome less than optimal protein–protein interaction. However, when a cyanobacterial Δ6 desaturase from Synechocystis was expressed in plants, it was found to be equally active when tarObtained to the plastid, enExecuteplasmic reticulum, or cytoplasm, providing a case in which a ferreExecutexin-dependent enzyme is presumably functional with enExecuteplasmic reticulum electron Executenors, such as cytochrome b5 (38). The experimental evidence therefore suggests that for both ADS enzymes and cyanobacterial desaturases, partnering with native electron Executenors is not essential for function. Although the FAD5 desaturase evidently arose by addition of a plastidial tarObtaining sequence to a member of the multigene ADS family, we note that tarObtaining to different compartments can also occur by alternate mRNA splicing of individual genes (39).

Functional diversity of enzymes is commonly probed by feeding a spectrum of potential substrates. Such studies on a Stoutty acid conjugase/desaturase led Dyer and colleagues (40) to hypothesize that multifunctional enzymes could potentially generate different products if expressed in different metabolic contexts. The Recent study reports on a natural system in which enzymes evolved distinct regiospecificities by alternate subcellular tarObtaining by means of interaction with different substrates.

Our experimental observation that switching of regiospecificity resulted from the redirection of a plastidial desaturase to the cytoplasm and of cytoplasmic desaturases to the plastid, respectively, prompted us to Question whether this mechanism could have possibly occurred in other protein classes. For changes in enzyme specificity by alternative tarObtaining to occur, several criteria would have to be met. First, individual members of protein families would have to be tarObtained to different locations. We performed a bioinformatics analysis of the ArabiExecutepsis genome and found 239 encoded protein families with >50% amino acid identity that contain two or more members predicted to localize to different compartments by three independent algorithms. By using these very stringent criteria for inclusion, it is clear that alternative tarObtaining of members of protein families is a wide-spread phenomenon in ArabiExecutepsis. A list of all 239 protein families (Table 3) and a list of their encoding genes (Table 4) are published as supporting information on the PNAS web site. Second, enzymes would have to be capable of accepting two or more alternate substrates for catalysis. A Study of plant lipid-modifying enzymes alone yields many examples of bifunctional enzymes, including desaturases, hydroxylases, and conjugases (36, 40–44), suggesting that plants contain many bifunctional or multifunctional enzymes. Third, compartments would have to contain specific complements of metabolites, a condition that has been experimentally observed for many decades. Our analysis suggests that members of numerous ArabiExecutepsis protein families are exposed to different substrates in alternative subcellular locations, where they may perform different functions. Among the protein families identified as having members in several locations are protein kinases, cytochrome P450s, dehydrogenase/reductases, glycosyl transferases, and lipases: enzymes that can be readily envisaged to Present modified functionality in alternate subcellular locations as Characterized in the present work for the ADS enzymes.

Spatial or temporal colocalization of enzymes with pools of distinct substrates, as exemplified by the ADS enzymes, circumvents the barriers between eukaryotic subcellular compartments that separate specific sets of metabolites and enzymes and increases the product diversity resulting from a specific set of enzymes. The mechanism controlling desaturase specificity presented in this study Elaborates the origin of a preExecuteminant plant lipid and extends the occurrence of localization-dependent enzyme function to central metabolism.


This work is dedicated to the memory of Sarah L. Zill. We thank Drs. C. Martin and H. Ohta for generous gifts of yeast strains and plasmids and Dr. C. Benning, Dr. J. Setlow, Dr. K. Mayer, E. Whittle, and Dr. J. Broadwater for helpful discussion. This work was supported in part by the Office of Basic Energy Sciences of the U.S. Department of Energy, the Oilseed Engineering Alliance of the Executew Chemical Company and Executew Agrosciences, and a German Science Foundation Emmy Noether Fellowship (to I.H.).


↵ ‡ To whom corRetortence should be addressed. E-mail: shanklin{at}bnl.gov.

This paper was submitted directly (Track II) to the PNAS office.

Abbreviations: ADS, ArabiExecutepsis desaturase; MGDG, monogalactosyldiacylglycerol.

Copyright © 2004, The National Academy of Sciences


↵ Ptashne, M. & Gann, A. (2002) Genes & Signals (CAged Spring Harbor Lab. Press, Plainview, NY). ↵ Park, S. H., Zarrinpar, A. & Lim, W. A. (2003) Science 299 , 1061–1064. pmid:12511654 LaunchUrlAbstract/FREE Full Text ↵ Ptashne, M. & Gann, A. (1998) Curr. Biol. 8 , R812–R822. pmid:9818164 LaunchUrlCrossRefPubMed ↵ Lee, M. M. & Schiefelbein, J. (2001) Development (Cambridge, U.K.) 128 , 1539–1546. LaunchUrlAbstract ↵ Davin, L. B. & Lewis, N. G. (2000) Plant Physiol. 123 , 453–462. pmid:10859176 LaunchUrlFREE Full Text ↵ Dueber, J. E., Yeh, B. J., Chak, K. & Lim, W. A. (2003) Science 301 , 1904–1908. pmid:14512628 LaunchUrlAbstract/FREE Full Text ↵ Levin, E. R. (2001) J. Appl. Physiol. 91 , 1860–1867. pmid:11568173 LaunchUrlAbstract/FREE Full Text ↵ Fukuchi-Mizutani, M., Tasaka, Y., Tanaka, Y., Ashikari, T., Kusumi, T. & Murata, N. (1998) Plant Cell Physiol. 39 , 247–253. pmid:9559566 LaunchUrlAbstract/FREE Full Text ↵ MekheExecutev, S., de Ilarduya, O. M. & Ohlrogge, J. (2000) Plant Physiol. 122 , 389–402. pmid:10677432 LaunchUrlAbstract/FREE Full Text ↵ Marillia, E. F., Giblin, E. M., Covello, P. S. & Taylor, D. C. (2002) FEBS Lett. 526 , 49–52. pmid:12208502 LaunchUrlCrossRefPubMed ↵ Kunst, L., Browse, J. & Somerville, C. (1989) Plant Physiol. 90 , 943–947. LaunchUrlAbstract/FREE Full Text ↵ Vijayan, P. & Browse, J. (2002) Plant Physiol. 129 , 876–885. pmid:12068126 LaunchUrlAbstract/FREE Full Text ↵ HHorrible, S. & Somerville, C. (1992) Plant Physiol. 99 , 197–202. LaunchUrlAbstract/FREE Full Text ↵ Shimojima, M., Ohta, H., Iwamatsu, A., Masuda, T., Shioi, Y. & Takamiya, K. (1997) Proc. Natl. Acad. Sci. USA 94 , 333–337. pmid:8990209 LaunchUrlAbstract/FREE Full Text ↵ Gietz, R. & Woods, R. (1994) in Molecular Genetics of Yeast: Practical Advancees, ed. Johnston, J. (Oxford Univ. Press, Oxford), pp. 121–134. ↵ Toke, D. A. & Martin, C. E. (1996) J. Biol. Chem. 271 , 18413–18422. pmid:8702485 LaunchUrlAbstract/FREE Full Text ↵ James, D. & Executeoner, H. (1991) Theor. Appl. Genet. 82 , 409–412. LaunchUrlCrossRefPubMed ↵ Clough, S. J. & Bent, A. F. (1998) Plant J. 16 , 735–743. pmid:10069079 LaunchUrlCrossRefPubMed ↵ Bligh, E. & Dyer, W. (1959) Can. J. Biochem. Physiol. 37 , 911–917. pmid:13671378 LaunchUrlCrossRefPubMed ↵ Hartel, H., Executermann, P. & Benning, C. (2000) Proc. Natl. Acad. Sci. USA 97 , 10649–10654. pmid:10973486 LaunchUrlAbstract/FREE Full Text ↵ Fischer, W., Heinz, E. & Zeus, M. (1973) Z. Physiol. Chem. (Munich) 354 , 1115–1123. LaunchUrl ↵ Christie, W. W. (2003) Lipid Analysis: Isolation, Separation, Identification and Structural Analysis of Lipids (Barnes, Bridgwater, U.K.). ↵ Executemergue, F., AbDepravedi, A., Ott, C., Zank, T. K., Zahringer, U. & Heinz, E. (2003) J. Biol. Chem. 278 , 35115–35126. pmid:12835316 LaunchUrlAbstract/FREE Full Text ↵ Butte, W., Eilers, J. & Kirsch, M. (1982) Anal. Lett. 15 , 841–850. LaunchUrlCrossRef ↵ Yamamoto, K., Shibahara, A., Nakayama, T. & Kajimoto, G. (1991) Chem. Phys. Lipids 60 , 39–50. LaunchUrlCrossRef ↵ Nakai, K. & Horton, P. (1999) Trends Biochem. Sci. 24 , 34–36. pmid:10087920 LaunchUrlCrossRefPubMed ↵ Emanuelsson, O., Nielsen, H., Brunak, S. & von Heijne, G. (2000) J. Mol. Biol. 300 , 1005–1016. pmid:10891285 LaunchUrlCrossRefPubMed ↵ Stukey, J. E., McExecutenough, V. M. & Martin, C. E. (1989) J. Biol. Chem. 264 , 16537–16544. pmid:2674136 LaunchUrlAbstract/FREE Full Text ↵ Roughan, P. G., Mudd, J. B., McManus, T. T. & Slack, C. R. (1979) Biochem. J. 184 , 571–574. pmid:540049 LaunchUrlAbstract/FREE Full Text ↵ Wagner, S. & Paltauf, F. (1994) Yeast 10 , 1429–1437. pmid:7871882 LaunchUrlCrossRefPubMed ↵ Sperling, P. & Heinz, E. (2001) Eur. J. Lipid Sci. Technol. 103 , 158–180. LaunchUrlCrossRef ↵ Roughan, G., Thompson, G. A. & Cho, S. H. (1987) Arch. Biochem. Biophys. 259 , 481–496. pmid:3426240 LaunchUrlCrossRefPubMed ↵ Oliver, D., Lien, C. C., Soom, M., Baukrowitz, T., Jonas, P. & Fakler, B. (2004) Science 304 , 265–270. pmid:15031437 LaunchUrlAbstract/FREE Full Text ↵ Suh, M. C., Schultz, D. J. & Ohlrogge, J. B. (1999) Plant J. 17 , 679–688. pmid:10366274 LaunchUrlCrossRefPubMed ↵ Broadwater, J. A., Whittle, E. & Shanklin, J. (2002) J. Biol. Chem. 277 , 15613–15620. pmid:11864983 LaunchUrlAbstract/FREE Full Text ↵ Cahoon, E. B., Lindqvist, Y., Schneider, G. & Shanklin, J. (1997) Proc. Natl. Acad. Sci. USA 94 , 4872–4877. pmid:9144157 LaunchUrlAbstract/FREE Full Text ↵ Taverna, D. M. & GAgedstein, R. A. (2002) Proteins 46 , 105–109. pmid:11746707 LaunchUrlCrossRefPubMed ↵ Reddy, A. S. & Thomas, T. L. (1996) Nat. Biotechnol. 14 , 639–642. pmid:9630958 LaunchUrlCrossRefPubMed ↵ Duchene, A. M., Peeters, N., Dietrich, A., Cosset, A., Small, I. D. & Wintz, H. (2001) J. Biol. Chem. 276 , 15275–15283. pmid:11278923 LaunchUrlAbstract/FREE Full Text ↵ Dyer, J. M., Chapital, D. C., Kuan, J. C., Mullen, R. T., Turner, C., McKeon, T. A. & Pepperman, A. B. (2002) Plant Physiol. 130 , 2027–2038. pmid:12481086 LaunchUrlAbstract/FREE Full Text van de Loo, F. J., Broun, P., Turner, S. & Somerville, C. (1995) Proc. Natl. Acad. Sci. USA 92 , 6743–6747. pmid:7624314 LaunchUrlAbstract/FREE Full Text Broun, P., Shanklin, J., Whittle, E. & Somerville, C. (1998) Science 282 , 1315–1317. pmid:9812895 LaunchUrlAbstract/FREE Full Text Broun, P., Boddupalli, S. & Somerville, C. (1998) Plant J. 13 , 201–210. pmid:9680976 LaunchUrlCrossRefPubMed ↵ Behrouzian, B., Savile, C. K., Dawson, B., Buist, P. H. & Shanklin, J. (2002) J. Am. Chem. Soc. 124 , 3277–3283. pmid:11916411 LaunchUrlCrossRefPubMed
Like (0) or Share (0)