biosynthesis of phytosterol esters: identiï¬cation of a

Biosynthesis of Phytosterol Esters: Identification of aSterol O-Acyltransferase in Arabidopsis1[OA]

Qilin Chen2, Lee Steinhauer, Joe Hammerlindl, Wilf Keller, and Jitao Zou*

Plant Biotechnology Institute, National Research Council Canada, Saskatoon, Saskatchewan, Canada S7N 0W9

Fatty acyl esters of phytosterols are a major form of sterol conjugates distributed in many parts of plants. In this study wereport an Arabidopsis (Arabidopsis thaliana) gene, AtSAT1 (At3g51970), which encodes for a novel sterol O-acyltransferase.When expressed in yeast (Saccharomyces cerevisiae), AtSAT1 mediated production of sterol esters enriched with lanosterol.Enzyme property assessment using cell-free lysate of yeast expressing AtSAT1 suggested the enzyme preferred cycloartenol asacyl acceptor and saturated fatty acyl-Coenyzme A as acyl donor. Taking a transgenic approach, we showed that Arabidopsisseeds overexpressing AtSAT1 accumulated fatty acyl esters of cycloartenol, accompanied by substantial decreases in estercontent of campesterol and b-sitosterol. Furthermore, fatty acid components of sterol esters from the transgenic lines wereenriched with saturated and long-chain fatty acids. The enhanced AtSAT1 expression resulted in decreased level of free sterols,but the total sterol content in the transgenic seeds increased by up to 60% compared to that in wild type. We conclude thatAtSAT1 mediates phytosterol ester biosynthesis, alternative to the route previously described for phospholipid:sterolacyltransferase, and provides the molecular basis for modification of phytosterol ester level in seeds.

Plant sterols, known generally as phytosterols, areintegral components of the membrane lipid bilayer(Demel and De Kruyff, 1976; Schuler et al., 1991). Un-like animal systems in which cholesterol is most oftenthe lone final product of sterol synthesis, each plantspecies has its own characteristic distribution of phy-tosterols, with the three most common phytosterols innature being b-sitosterol, campesterol, and stigmas-terol (Benveniste, 1986, 2004). In addition to free sterolform, phytosterols are also found in the form of con-jugates, particularly fatty acyl sterol esters (SEs). SEscan be found in lipid bodies in the cytoplasm andare present at a substantial level in seeds. In canola(Brassica napus), for example, phytosterols constituteabout 0.5% of seed oil (Sabir et al., 2003), 35% of whichis in the form of SEs (Harker et al., 2003). The bio-chemical process of sterol acylation is believed to playa role in maintaining the free sterol content of cellmembranes at their physiological levels (Schaller,2004).

The effectiveness of phytosterols as a dietary com-ponent to lower serum cholesterol level in humans hasbeen well documented in medical research for morethan half a century (Moghadasian et al., 1997, 1999;Hendriks et al., 1999; Sierksma et al., 1999; Piironenet al., 2000; van Rensburg et al., 2000; Awad et al., 2001,2003; Bouic, 2001). It also has been suggested that theaverage diet of the western world has a phytosterolintake far below prehistoric levels (Yankah and Jone,2001). These studies led to recommendations of con-sumption of phytosterol-enriched food products bythe general population and particularly mildly hyper-cholesterolemic subjects. However, limited quantitiesof phytosterols are currently a major barrier in satis-fying the demands for such functional foods. Based ona firmly established concept that 3-hydroxy-3-methyl-glutaryl-CoA reductase (HMGR) plays a rate-limitingrole in the flux control of sterol biosynthesis (Bach,1986; Gondet et al., 1992, 1994; Chappell et al., 1995),efforts aimed at increasing phytosterol content havebeen concentrated on enhancing the activity of HMGRthrough introducing modified HMGR that is resistantto regulation (Chappell et al., 1995; Re et al., 1995;Schaller et al., 1995; Harker et al., 2003; Hey et al.,2006). Nonetheless, evidence suggesting the existenceof other regulatory step(s) in the flux control of sterolbiosynthesis can be found in some studies, particularlythat of Gondet et al. (1994) who studied a sterol-overproducing tobacco (Nicotiana tabacum) mutantLAB1-4, with HMGR levels approximately 3-fold higherthan normal (Gondet et al., 1994). LAB1-4 had a 10-foldstimulation of sterol content in calli as compared towild type, mostly in the form of SEs, but a mere 3-foldstimulation in the regenerated leaves. Intriguingly,both type of tissues had the same stimulation factorof HMGR (3-fold). Therefore, there appears to be other

1 This work was supported by the National Research CouncilCanada-Crops for Enhanced Human Health Program and a NaturalScience and Engineering Research Council of Canada grant (grantno. NSERC RGPIN 327217–06 to J.Z.). Q.C. is a recipient of the Post-Doctoral Fellowship for Visiting Government Laboratories from theNatural Science and Engineering Research Council of Canada.

2 Present address: Department of Biology, University of Saskatch-ewan, Saskatoon, Saskatchewan, Canada S7N 5E2.

* Corresponding author; e-mail [email protected] author responsible for distribution of materials integral to

the findings presented in this article in accordance with the policydescribed in the Instructions for Authors (www.plantphysiol.org) is:Jitao Zou ([email protected]).

[OA] Open Access articles can be viewed online without a sub-scription.

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contributing factor(s), independent of HMGR, partici-pating in regulating total sterol content.

Despite the ubiquitous presence of phytosterol es-ters in plant cells, the metabolic process of sterol acyla-tion remains poorly understood. Sterol esterificationis suggested to take place using several acyl donorsincluding phospholipid, diacylglycerol, or triacylglyc-erol (Zimowski and Wojciechowski, 1981a, 1981b;Dyas and Goad, 1993). A recent report on the phos-pholipid:sterol O-acyltransferase gene from Arabidop-sis (Arabidopsis thaliana; AtPSAT) established a routeof transacylation between phospholipids and sterolsin plant SE synthesis (Banas et al., 2005). AtPSAT is alecithin:cholesterol acyltransferase protein family mem-ber homologous to human soluble lecithin:cholesterolacyltransferase. In this study, we took a functional com-plementation approach using a yeast (Saccharomycescerevisiae) strain in which two sterol O-acyltransferase-encoding genes, ARE1 and ARE2, are disrupted. Weidentified a novel Arabidopsis sterol O-acyltransferaseAtSAT1, which is structurally related to the acyl-CoAcholesterol acyltransferase (ACAT) in animal systems.We present evidence that overexpression of AtSAT1alters SE synthesis in Arabidopsis, providing elevatedlevels of commercially desirable phytosterols.

RESULTS

Cloning of AtSAT1 from Arabidopsis

Taking a yeast functional complementation strategy(Bach and Benveniste, 1997), we focused our searchof a plant sterol O-acyltransferase to the superfamilyof membrane-bound O-acyltransferases (MBOATs;Hofmann, 2000), because the common biochemicalfunctionality of the MBOAT family proteins is to me-diate the transfer of organic acids in ester or thioesterforms to hydroxyl groups of membrane-embeddedacyl acceptors.

The yeast mutant SCY059 produces only a re-sidual amount of SE because the two principle sterolO-acyltransferase genes, ARE1 and ARE2, were inter-rupted (Yang et al., 1996). Employing this strain, weintroduced members of the Arabidopsis MBOAT fam-ily to test functional complementation. Fifteen cDNAscorresponding to MBOAT family candidate genes(listed in ‘‘Materials and Methods’’) were cloned intothe yeast expression vector pYES2.1 under the controlof GAL1 promoter. The yeast transformants weresubjected to GAL1 induction condition and harvestedfor neutral lipid analysis. When separated by thin-layerchromatography (TLC), SCY059 harboring At3g51970produced an additional band in the neutral lipid ex-traction (Fig. 1A). HPLC analysis also revealed similarresults from these cells (Fig. 1B). Both the TLC Rf valueand the HPLC retention time of the novel productswere consistent with that of SE.

To verify the identity of the product, the HPLC frac-tion was collected and subjected to alkaline hydrolysis(saponification). The saponification extract was deriv-

atized with N,O-bis(trimethysily)-trifluoroacetamide 11% trimethylchlorosilane (BSTFA 1 1% TMCS) anddetected as trimethylsilyl (TMS) derivatives by gaschromatography (GC)-mass spectrometry (MS). GCprofile revealed four major peaks from the sterolfraction (Fig. 2A). All the GC fractions were identifiedby searching the National Institute of Standards andTechnology (NIST) 2.0 mass spectra library. The firstpeak, also present in the control strain, correspondedto squalene. The other three peaks displayed massspectra identical to that of ergosterol, lanosterol, and4,4-dimethyl-8,24-cholestadienol (also known as 4,4-dimethylzymosterol), respectively (Fig. 2A). The fivepeaks from the fatty acid fraction were found to be14:0, 16:1, 16:0, 18:1, and 18:0 fatty acids (Fig. 2B).

Figure 1. TLC and HPLC separation of neutral lipids from SCY059harboring control vector (vector-only), SCY059 expressing AtSAT1(AtSAT1), and a parental strain SCY062 (WT). A, Neutral lipids sepa-rated through TLC developed in hexane/diethyl ether/acetic acid (40/10/1.5). B, Normal-phase HPLC separation of neutral lipids extractedfrom SCY059 expressing AtSAT1 (AtSAT1) and SCY059 harboring thevector (vector-only). Arrows denote the novel product found in SCY059expressing AtSAT1.

Arabidopsis Phytosterol O-Acyltransferase

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These results confirmed that expression of At3g51970conferred SCY059 an ability to produce SE andAt3g51970 was then tentatively designated as AtSAT1.

AtSAT1 Is Structurally Related to Acyl-CoA SterolAcyltransferases of Yeast and Animal Origins

AtSAT1 is predicted to be an integral membraneprotein with eight transmembrane domains, and has aputative signal peptide with a cleavage site at the 19thamino acid from the N terminus, which was proposedto direct nascent proteins into the secretory pathway,including the endoplasm reticulum (Nielsen et al.,1997). Deduced amino acid sequence alignment showsthat AtSAT1 is structurally related to acyl-CoA sterolacyltransferase from yeast and animal systems (Fig. 3),among which ACATs have been extensively studied.The RxWNxxVxxxLxxxVY motif present in all ACATs,commonly believed to be crucial for fatty acyl-CoAbinding (Guo et al., 2001), is also found in AtSAT1. A

His residue (His245 in AtSAT1) located within a longstretch of hydrophobic amino acids is present at the Cterminus of AtSAT1. This conserved His residue wassuggested to be a part of the active site (Hofmann,2000). However, when compared with other ACATs,AtSAT1 protein is a shorter polypeptide, lacking theextended N-terminal region found in other sterolacyltransferases. Moreover, two conservative motifs,FYxDWWN and H/YSF previously suggested as be-ing crucial for ACAT activity (Cao et al., 1996; Oelkerset al., 1998), were also absent in AtSAT1.

Heterologous Expression of AtSAT1 in Yeast ProducesMainly Lanosterol Esters

We estimated SE content of the yeast strains by quan-tifying sterols released through alkaline hydrolysis. Asreported previously (Yang et al., 1996), the vector-onlytransformant of SCY059 still produced a trace amountof SE (0.027 mmol/g dry weight [DW] of yeast cells).The same strain heterologously expressing AtSAT1 in-creased SE production by 10-fold to 2.7 mmol/g DW,though still lower than 9.4 mmol/g DW of parentalwild-type strain SCY062. The levels of free sterols inSCY059, SCY059/AtSAT1, and SCY062 were found at18.1 mmol/g DW, 14.9 mmol/g DW, and 13.2 mmol/gDW, respectively.

Considering SCY059 still retained a low level of SEsynthesis capacity, there was the possibility of AtSAT1being a facilitator, enhancing the residual SE synthesisactivity in the mutant strain. If this were true, SCY059harboring AtSAT1 would likely produce SE with aprofile of sterol or fatty acid moiety similar to that ofthe vector-alone transformant. This was experimen-tally proven untrue. As shown in Figure 4A, the molarcomposition of the fatty acid species saponified fromthe SE collected from the vector-only strain and theSEs produced by SCY059/AtSAT1 were different. Like-wise, the sterol composition of the SE also displayedmajor differences due to the expression of AtSAT1 (Fig.4B). The residual SE in SCY059/empty vector con-sisted of mainly ergosterol. On the other hand, expres-sion of AtSAT1 resulted in production of SE containingchiefly lanosterol, which contributed to 88.1% of totalsterol moiety. Thus, AtSAT1 appeared to preferentiallyacylate lanosterol when expressed in yeast.

Substrate Preference of AtSAT1

In light of substrate preferences of ACAT-relatedenzymes reported to date, we examined fatty acyldonor preferences of AtSAT1 using cell-free lysate ofyeast with various fatty acyl-CoAs. Lipids extractedfrom in vitro reactions were separated by TLC and theincorporation of [3H]lanosterol into SE was measured.Because the cell-free lysate was expected to containcertain amount of acyl-CoA, reactions without addedacyl-CoA were set as blank controls for enzyme activ-ity calculation. As shown in Figure 5A, AtSAT1 hada substrate preference in the order of 16:0 . 18:0 .

Figure 2. Analysis of TMS derivatives of sterols and fatty acids of SE.A, GC profiles of sterols saponified from SE of yeast strain SCY059harboring control vector (I) and SCY059 expressing AtSAT1 (II). B, GCprofile of TMS derivatives of fatty acids released from SE of AtSAT1yeast transformant. The TMS derivatives of peaks in the profiles weresubsequently subjected to mass spectrum analysis and the mass spec-tra were found as being identical to: 1, Squalene; 2, ergosterol; 3,lanosterol; 4, 4,4-dimethyl-8,24-cholestadienol; 5, myristic acid (14:0);6, palmitoleic acid (16:1); 7, palmitic acid (16:0); 8, oleic acid (18:1);and 9, stearic acid (18:0).

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16:1 . 18:1. 18:2 was found to negatively affect sterolO-acyltransfease activity.

We used stigmasterol, b-sitosterol, and cycloartenolas well as two sterols of yeast origin, lanosterol andergosterol, to assess [14C]16:0-CoA acylation. Assess-ment of sterol substrate preferences entailed a methyl-

b-cyclodextrin wash to extract free sterols from thepreparations (Rodal et al., 1999), due to the concernthat endogenous sterols may interfere with substratespecificity assessment. However, there apparently wasstill a substantial amount of endogenous sterols re-maining, because the background value in the absence

Figure 3. Sequence alignment of AtSAT1 with other reported sterol acyltransferases. Alignment was performed with CLUSTALVfrom the DNASTAR package run with default multiple alignment parameters (gap opening penalty: 10, gap extension penalty:10). Accession numbers of proteins are: AtSAT1 (Arabidopsis, AAQ65159); HsACAT1 (Homo sapiens, NP_003092); HsACAT2(H. sapiens, NP_003569); MmACAT1 (Mus musculus, Q61263); MmACAT2 (M. musculus, NP_666176); RnACAT1 (Rattusnorvegicus, O70536); RnACAT2 (R. norvegicus, NP_714950); ScARE1 (yeast, P25628); and ScARE2 (yeast, P53629). Residuesboxed in dashed or solid lines were previously reported as conserved functional motifs in ACATs. The black bar on the bottommarks a shared domain, and the arrow indicates the His likely to be an active-site residue (Hofmann, 2000).





of added sterol substrate was nonetheless high. Wealso attempted acetone extraction of the membranefractions to reduce the endogenous sterol levels. Un-fortunately the acetone treatment essentially elimi-nated the enzyme activity (data not shown). This ledus to use reactions without exogenous sterol additionsas control experiments. Under our assay conditions,the yeast strain harboring empty vector displayed areduced 16:0 acylation activity in the presence ofexogenous sterol. Addition of ergosterol to the yeastlysate expressing AtSAT1 also gave rise to a negativevalue on SE formation. Such a negative effect ofexogenous sterol addition on sterol O-acyltransferaseactivity was previously reported for animal ACAT(Tavani et al., 1982; Yang et al., 1997) and the Arabi-dopsis AtPSAT1 assays (Banas et al., 2005). Despitethese results, we could consistently detect SE synthesisactivity with most of the sterol substrates whenAtSAT1 was expressed. The highest activity was foundwith cycloartenol, followed by b-sitosterol, lanosterol,and stigmasterol (Fig. 5B).

Because phosphatidylcholine (PC) has been previ-ously implicated as an acyl donor for SE synthesis(Banas et al., 2005), the ability of AtSAT1 to use thislipid as substrate was assessed using dipalmitoyl-PCwith [14C]label in both the sn-1 and sn-2 fatty acylmoieties. We found that, while the PC could indeedsustain SE synthesis above the background of the

control yeast strain, it was nonetheless at a level one-fiftieth of that when 16:0-CoA was provided. We alsoexpressed AtSAT1 in the are1are2lro1dga1 quadrupleyeast mutant devoid of triacylglycerol (Sandager et al.,2002). While the formation of triacylglycerols was notdetectable, the quadruple yeast mutant expressingAtSAT1 was still capable of producing SE. This resultwas significant in two respects: first, it suggested thatAtSAT1 did not require triacyglycerols (TAGs) as afatty acyl donor for SE biosynthesis, and second, itcould not acylate diacylglycerol to produce TAG.

Expression Profile of AtSAT1

Based on transcript profile obtained from Geneves-tigator at https://www.genevestigator.ethz.ch/, AtSAT1is expressed in all tissues examined, but particularlyabundant in the elongation zone of roots followedby developing microspore and germinating seedlings.

Figure 4. Fatty acid profile and sterol composition of SE in yeast strains.A, Mol % of fatty acid species released from SE of SCY059 expressingAtSAT1 in comparison to that of the vector-only control and theparental strain (WT). B, Profiles of the sterol moieties of SE. The SE wassaponified in methanolic-KOH for 2 h at 80�C. The fatty acids weretransmethylated with methanolic HCl and free sterols were derivatizedwith BSTFA (1% TMCS).

Figure 5. Substrate preference assessment of AtSAT1. A, Fatty acyl-CoAselectivity of AtSAT1. Cell-free lysates were assayed for acylation of[3H]lanosterol. Fatty acyl-CoAs used for the assay were palmitoyl-CoA(16:0), palmitoleoyl-CoA (16:1), stearoyl-CoA (18:0), oleoyl-CoA (18:1),and linoleoyl-CoA (18:2). B, Sterol preference of AtSAT1. Cell-freelysates were assessed for acylation of ergosterol, lanosterol, stigmasterol,b-sitosterol, and cycloartenol in the presence of [14C]palmitoyal-CoA.Reactions without exogenous sterol (or fatty acyl-CoA) served ascontrol. Enzyme activity was calculated based on the difference ofsterol acylation between reactions with added sterol (or fatty acyl-CoAs) substrates and reactions without sterols (or fatty acyl-CoAs). Thenegative values in some reactions were caused by high backgroundsterol acylation in the control in the absence of exogenous sterolsubstrates.

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Previous published works (Ting et al., 1998; Hernandez-Pinzon et al., 1999) have shown the accumulation of SEin developing pollen and seed. AtSAT1 expression levelincreased as pollen development proceeded from uni-nucleate microspore to bicellular pollen, tricellularpollen, and mature pollen (Honys and Twell, 2004).Increased expression of AtSAT1 was also an evidenttrend during seed development.

Overexpression of AtSAT1 in Arabidopsis Leads to

Cycloartenol Ester Overproduction

To further confirm that AtSAT1 is involved in SEbiosynthesis in plants, we cloned AtSAT1 cDNA into a

plant expression vector under the control of a seed-specific napin promoter (Josefsson et al., 1987). Thetransgenic plants showed no apparent developmentaldifference from wild-type plants. Real-time quantita-tive (qRT)-PCR revealed that AtSAT1 was approxi-mately 20-fold overexpressed in developing siliques ofmost AtSAT1 transgenic lines (Fig. 6).

Seeds harvested from four wild-type plants and 12transgenic plant (T1) lines, raised simultaneously inthe same growth chamber, were selected for sterolanalysis. AtSAT1 overexpression led to increases in SEby more than 2-fold and total sterol content by 64% insome transgenic lines (Table I). Overexpression ofAtSAT1 in seed, as exemplified by one transgenic lineshown in Fig. 7, also drastically altered the SE-sterolprofile. In wild-type seeds, the two major SE species,b-sitosterol ester and campesterol ester, accounted for80.4% and 17.5% of total SE, respectively. Cycloartenolester was detected as a minor component at a distantthird with 2.1%. Strikingly, the AtSAT1 transgenicseeds produced cycloartenol ester as the most prom-inent component, representing up to an average of64.1% of total SE. On the other hand, the esters ofb-sitosterol and campesterol were reduced to averagesof 25.2% and 5.2%, respectively. We also found that24-methylene cycloartenol was increased from non-detectable level in wild-type seeds to 5.6% in thetransgenic line seeds. The enhanced SE overproduc-tion also caused reduction of free sterol from 58.8% inwild type to 28.3% of total sterols in transgenic plants.

We further analyzed the profile of the fatty acidcomponent of the SEs in seeds overexpressing AtSAT1(Table II). In comparison to wild-type seeds, there wasa drastic increase in the molar percentage of 16:0, from

Figure 6. qRT-PCR analysis of transcript levels of AtSAT1. Each samplecontained RNA extracted from pooled developing siliques. qRT-PCRwas performed using Applied Biosystem StepOne Real-Time PCRsystem. CT was automatically determined by software StepOne (version1.0). AtSAT1 expression levels were calculated through normalizationwith b-Actin-8 by the formula 22(CTAtSAT1 2 CTb-actin). Data represent themean and SD of three replicates.

Table I. Content of phytosterol esters and free sterols in wild-type and AtSAT1 transgenic seeds

Arabidopsis (Columbia) plants were transformed with vector pSE129TAtSAT1, in which AtSAT1 was under the control of a napin promoter. The fourwild-type (WT) controls and 12 AtSAT1 overexpression (OE) lines were raised under identical conditions. Lipids were extracted in chloroform/methanol (2/1) from WT and transgenic seeds, followed by separation via normal-phase HPLC. SE fractions were collected and saponified in 7.5%methanolic KOH. Released free sterols were derivatized with BSTFA containing 1% TMCS and quantified by GC. The data were presented as averagesof two analyses. Camp, Campesterol; Sito, sitosterol; Cyclo, cycloartenol; MethyCyclo, 24-methylene cycloartenol; nd, nondetectable.

LinesSE Composition

Total SE SterolFree Sterol Composition

Total Free Sterol Total SterolCamp Sito Cyclo Methy-Cyclol Camp Sito Cyclo Methy Cyclo

% % % % mg/mg % % % % mg/mg mg/mg

WT1 16.3 79.8 4.0 nd 1.7 12.1 74.9 1.3 11.8 2.1 3.8WT2 18.2 78.9 2.9 nd 1.6 10.3 69.7 0.0 20.1 2.6 4.2WT3 17.3 82.7 0.0 nd 1.8 11.8 66.4 0.6 21.2 2.9 4.7WT4 18.1 80.2 1.7 nd 1.6 14.8 69.8 0.9 14.6 2.1 3.7OE1 2.7 18.0 73.0 6.3 4.5 16.3 74.2 1.7 7.7 1.2 5.7OE2 5.3 27.8 61.5 5.5 3.5 12.5 69.3 2.5 15.7 2.1 5.6OE3 7.3 34.5 52.3 5.8 3.4 11.6 70.3 1.1 17.1 1.8 5.2OE4 5.3 26.4 63.7 4.7 4.0 11.0 66.4 0.9 21.8 1.8 5.9OE5 4.6 22.0 67.7 5.7 4.6 11.1 63.9 2.1 22.9 1.7 6.3OE6 4.1 19.7 70.1 6.1 5.0 12.1 64.0 1.9 22.1 1.8 6.7OE7 3.7 19.4 71.8 5.2 5.2 14.2 68.1 2.2 15.6 1.4 6.6OE8 7.0 30.5 57.6 5.0 3.1 16.7 67.2 2.6 13.5 1.5 4.6OE9 5.6 27.8 60.9 5.8 3.8 18.7 73.7 3.4 4.3 1.2 5.0OE10 4.5 20.6 69.5 5.4 4.3 15.1 67.9 2.5 14.5 1.4 5.7OE11 6.9 31.2 56.1 5.9 3.2 12.3 67.1 1.9 18.8 2.0 5.2OE12 5.0 24.3 64.5 6.3 3.8 12.7 68.7 1.5 17.1 1.3 5.1





less than 15% to approaching 30%. Significant increaseswere also apparent in the composition of very long-chain fatty acids 20:0 and 20:1. The overall changes infatty acid component of the SEs in the transgenic linescan be summarized as saturated and long-chain fattyacids replacing unsaturated fatty acids in wild-typeseeds, particularly 18:2 and 18:3. The fatty acid com-position of SE in AtSAT1-overexpressed seeds was con-sistent with the in vitro fatty-CoA specificity results.

DISCUSSION

In this study we identified AtSAT1 as a sterolO-acyltransferase from Arabidopsis that belongs tothe MBOAT family. Our conclusion on the functionalidentity of AtSAT1 was based on the following linesof evidence: (1) heterologous expression of AtSAT1enabled a yeast SE-deficient mutant SCY059 to syn-thesize a considerable amount of SE; (2) AtSAT1 me-diated the synthesis of SE with sterol and fatty acidprofiles distinctively different from that of the traceamount of SE synthesized in the yeast mutant strain;and (3) transgenic overexpression of AtSAT1 in Arab-idopsis substantially increased SE content in seeds.Furthermore, since we did not find detectable levels ofTAG when expressing AtSAT1 in the are1are2lro1dga1quadruple yeast mutant, it appears that AtSAT1 couldnot acylate diacylglycerol for TAG biosynthesis.

Expression of AtSAT1 in yeast strain SCY059 re-sulted in the accumulation of SE consisting mainly oflanosterol, rather than the most abundant sterol beingergosterol, the end product of the sterol biosynthesispathway in yeast (Zweytick et al., 2000), suggestingthat AtSAT1 possesses a certain degree of sterol sub-strate specificity. The SE composition resulting fromexpression of AtSAT1 was clearly different from re-ports of overexpressing the ARE1 and ARE2 genes inSCY059, which produced SE enriched in zymosteroland erogosterol (Jensen-Pergakes et al., 2001). Thus, itcan also be inferred that sterol preference of AtSAT1 isdistinctively different from that of the yeast sterolO-acyltransferases. AtSAT1 also appeared to possesssubstrate preference toward fatty acyl donors, display-ing the highest activity when saturated fatty acyl-CoAs(16:0 and 18:0) were provided. This is in contrast tothe Arabidopsis phospholipid sterol O-acyltransferaseAtPSAT, which discriminates against saturated fattyacids and was suggested to play a minor role in thesynthesis of SE with saturated fatty acids (Banas et al.,2005). On this basis, it is important to note that pre-viously published work has established that SE insome tissues of Brassica and Arabidopsis are highlyenriched in saturated fatty acids (Hernandez-Pinzonet al., 1999).

Data obtained from transgenic plants were generallyin line with results of enzyme property assessment per-formed through heterologous expression of AtSAT1in yeast. Consistent with the high enzyme activitydetected with cycloartenol in our in vitro assays, en-

hanced AtSAT1 expression drastically advanced accu-mulation of cycloartenol esters in Arabidopsis seeds.Likewise, for the fatty acid components of the SEs,we observed increases in the molar ratio of saturatedfatty acids, including 16:0, 18:0, and 20:0. Elevatedsterol acylation with the long-chain monounsaturatedeicosenoic acid was also evident. On the other hand,the polyunsaturated fatty acids were proportionallyreduced.

Previous biochemical studies suggested that thereare several potential acyl donors for SE biosynthesisin plants (Garcia and Mudd, 1978a, 1978b, 1978c;Zimowski and Wojciechowski, 1981a, 1981b), includingacyl-CoA, phospholipids, diacylglycerol, and triglyc-eride. In the case of AtSAT1, triglyceride can be ruledout because TAG-deficient are1are2lro1dga1 quadrupleyeast mutant expressing AtSAT1 produced SE. WhenPC was supplied as an acyl donor the detected sterolacylation was above residual level of the control, butfar below that in the presence of acyl-CoA. It is unclearwhether PC served as an immediate acyl donor orindirectly through mobilization of acyl group from PCto the fatty acyl-CoA pool. We suggest that AtSAT1 isan acyl-CoA:sterol acyltransferase. However, since thespecific activity in our assay was lower than that ofother reported ACATs (Macauley et al., 1986; Yanget al., 1997), and our in vitro assays with yeast celllysate was complicated by a high background of sterol

Figure 7. GC profiles of sterols saponified from SE extracted from seedsof wild type (WT) and one representative AtSAT1 overexpression line.Lipids were extracted from seeds with chloroform:methanol (2:1, v/v)and separated though normal-phase HPLC. SE fractions were collectedand saponified with 7.5% KOH in 95% methanol. The resulting freesterols were derivatized with BSTFA:pyridine (1:1, v/v). Identification ofmass spectra and assignation of GC/MS peaks was carried out by alibrary search (NIST, version 2.0). 1, Cholesterol (internal standard);2, campesterol; 3, b-sitosterol; 4, cycloartenol; and 5, 24-methylenecycloartenol.

Chen et al.




acylation, this supposition should be approached withcaution. In light of the fact that AtSAT1 belongs to alarge family of proteins in which some have promis-cuous substrate preference, the substrate specificity ofAtSAT1 requires further study.

SE has been long proposed to be the storage andtransport form of sterols for their intracellular move-ment (Kemp et al., 1967; Janiszowska and Kasprzyk,1977) and movement between tissues (Holmer et al.,1973). Radiolabeling experiments in Apium graveolenssuspension cell culture indicated SE pool is underrapid turnover with esterification and hydrolysis pro-cess occurring concomitantly (Dyas and Goad, 1993). Itwas also proposed that esterification and hydrolysisshould occur alternatively to newly synthesized sterolintermediates for orderly structural modification untilend product 4-desmethylsterols were formed. In ourstudy, seed-specific overexpression of AtSAT1 in Arab-idopsis resulted in the accumulation of cycloartenolesters, which otherwise occur only as a minor sterolcomponent in wild-type seeds. Taken together withthe in vitro sterol substrate preference results, thesimplest explanation for such a metabolic outcome isthat AtSAT1 selectively channeled cycloartenol intothe ester pool (Fig. 8). Cycloartenol is the first tetracy-clic sterol precursors in plant sterol biosynthesis path-way, and is considered an intermediate in thephytosterol biosynthesis pathway equivalent to lanos-terol in yeast and animal systems (Benveniste 1986;Lovato et al., 2000; Kolesnikova et al., 2006; Suzukiet al., 2006). After cycloartenol, several major biochem-ical steps including methyltransferase, demethylation,and desaturation reactions need to take place beforethe final products of b-sitosterol and campesterol canbe formed (Benveniste, 2004). The elevated expressionof AtSAT1 could enhance the esterification of cyclo-artenol, thereby removing cycloartenol from the main-

stream of sterol biosynthetic pathway into the formationof esters. This is consistent with our results in thetransgenic seeds where the contents of b-sitosterol andcampesterol, regardless if in the form of free or ester-ified forms, were reduced when compared to wildtype. A question remains as to the physiological sig-nificance of such a role of AtSAT1 in plant metabolismand development. While highly speculative, we sug-gest that because cycloartenol has much better micel-lar solubility than the sterol end products, a role forAtSAT1 is to synthesize cycloartenol esters, which mayserve as a significant form of transported sterol con-jugates in certain developmental stages.

Incorporating phytosterols into the diet may be aneffective approach to lower total and LDL cholesterollevels in humans, but free phytosterols are difficult toincorporate into commercial foods because of theirlow solubility. Fortunately, phytosterol esters can bedissolved in vegetable oil at a concentration 10 timeshigher than that of the free phytosterols. In this study,we found that overexpression of AtSAT1 could lead toenhanced total sterol content, primarily due to theincrease in cycloartenol ester level. Cycloartenol isparticularly enriched in oryzanol derived from rice(Oryza sativa) bran oil (Rukmini and Raghuram, 1991).It has been suggested that cycloartenol is the effectiveingredient in rice bran oil for lowering plasma non-HDL cholesterol level (Wilson et al., 2007). We believethat the engineering outcome of the current research islikely to be repeated in crops species, particularlycanola, offering a novel genetic engineering target forenhanced phytosterol content in vegetable oil. Wemust note, although the increase in seed oil SE contentwas significant, increase in total sterol content was stillmodest. Further studies on the regulation of phytoste-rol biosynthesis are clearly required to improve phy-tosterol content in seeds.

Table II. Fatty acid composition of SEs in wild-type (WT) and AtSAT1 overexpression (OE) plants

WT and transgenic seeds were from the same lines as that in Table I. SE purified through HPLC wastransmethylated in methanolic HCl. The resultant fatty acid methyl ester products were quantified by GC forfatty acid mole ratio calculation. The data were presented as averages of two analyses.

Lines 16:0 16:1 18:0 18:1 18:2 18:3 20:0 20:1

WT1 9.0 3.1 3.0 6.6 50.7 23.3 0.1 4.3WT2 16.0 3.6 7.9 11.2 38.6 17.3 2.4 3.0WT3 13.5 3.1 4.0 11.6 33.9 22.7 2.1 8.3WT4 14.0 7.3 4.7 9.1 41.0 20.8 0.1 3.1OE1 28.3 2.6 14.5 11.5 9.1 4.1 8.1 21.9OE2 29.3 2.3 9.2 9.0 13.1 6.0 8.3 22.6OE3 24.7 2.4 8.0 9.4 17.9 8.7 7.5 21.4OE4 22.6 6.8 8.6 14.4 15.8 8.3 6.0 17.6OE5 32.1 2.2 8.7 8.8 11.6 4.3 9.1 23.2OE6 29.1 1.6 8.5 9.8 10.8 4.6 9.0 26.5OE7 28.5 2.1 8.3 8.7 12.3 5.0 9.3 25.7OE8 29.1 2.9 8.3 9.0 11.9 4.5 9.2 24.9OE9 25.3 2.3 10.2 10.9 15.8 6.8 7.2 21.5OE10 27.6 2.4 9.7 8.4 11.9 5.6 9.4 25.0OE11 24.4 0.9 7.6 9.8 18.3 8.1 8.4 22.6OE12 23.7 2.2 8.8 11.6 15.2 6.3 8.2 24.0





MATERIALS AND METHODS

TOPA TA Cloning and Yeast Complementation

RT-PCR of the open reading frames of Arabidopsis (Arabidopsis thaliana)

MBOAT family genes At1g12460, At1g34490, At1g34500, At1g34520, At1g57600,

At3g08930, At3g51970, At5g01460, At5g55320, At5g55330, At5g55340, At5g55350,

At5g55360, At5g55370, and At5g55380 was performed with primer pairs

designed based on sequences of gene annotation available on The Arabidopsis

Information Resource Web site. The primer pair for At3g51970 cDNA was

5#-CCATGGCGAGTTTCATCAAGGCAT-3# (forward primer) and 5#-GGC-

AGGGTTAAAAAAGATATGCGGTCAGT-3# (reverse primer). The cDNA was

cloned into vector pYES2.1 using pYES2.1 TOPO TA cloning kit according to the

manufacturer’s protocol (Invitrogen), and subsequently introduced into yeast

(Saccharomyces cerevisiae) strain SCY059 (MATa, his 3-11,15 leu 2-3, 112 trp 1-1, ura

3-1, kan 1-100, ade2 met14Dare1THIS3 are2TLEU2), kindly providedby Dr. Stephen

Sturley (Columbia University), and a quadruple mutant strain (are1 are2 lro1

dgat1) from Dr. Sten Stymne (Swedish University of Agricultural Sciences).

Yeast Growth, Lipid Extraction, and HPLC Analysis

Single colonies of vector-only or AtSAT1 transformed yeast were inocu-

lated in 10 mL of synthetic complete medium with Leu, His, and Ura omitted,

and grown overnight at 28�C. Twenty-milliliters of fresh medium was then

added and the cultures were further maintained overnight. Neutral lipid

fraction was extracted following an established protocol (Folch et al., 1957)

and separated through normal-phase HPLC on Agilent HPLC 1100 system by

the method detailed by Moreu et al. (1996).

Qualitative Analysis of Sterol and Fatty Acid Releasedfrom SE

SEs were saponified for 2 h in 1 mL of 7.5% methanolic KOH at 80�C. The

released sterol was extracted twice with hexane and dried under N2 stream.

The aqueous layer was neutralized with HCl and extracted twice with hexane

to obtain free fatty acids. The fatty acid fraction was derivatized with BSTFA

containing 1% TMCS and sterol derivatives were obtained in a mixture of

BSTFA and pyridine (1:1, v/v) at room temperature for 1 h, and separated on a

Hewlett-Packard 6890 N GC with a capillary column DB-5 (Hewlett-Packard

HP5, 35 m length 3 0.25 mm diameter 3 0.25 mm thickness). GC-MS analysis

was accomplished using an Agilent 5973 mass selective detector coupled to an

Agilent 6890 N gas chromatograph. The mass selective detector was run under

standard electron impact conditions (70 eV), scanning an effective mass-to-

charge ratio range of 40 to 700 at 2.26 scans/s. The mass spectra were

compared with entries in the NIST mass-spectral database, version 2.0.

Quantitative Analysis of SE

After HPLC separation of neutral lipid, SE fraction was collected, dried

under N2 stream, and saponified in 7.5% KOH in 95% methanol. Separation

and quantification of sterol species was performed on a Hewlett-Packard 6890

series GC with a capillary column DB-5 and flame ionization detector. The

peaks of each sterol species were compared to the cholesterol (internal

standard) peak to determine the amount of each sterol present. Fatty acid

methyl esters were separated on GC with DB23 column. The quantification

was performed in triplicate with three different batches of yeast cultures. The

content of SE was expressed as micromoles of sterol moiety saponified from

SE on a dry yeast cell weight basis.

Sterol O-Acyltransferase Activity Assay Using

Cell-Free Lysates

AtSAT1 expression induction in yeast cells was performed according to

manufacturer’s protocol for pYES2.1 TOPO TA expression kit (Invitrogen). The

cells, resuspended in 1 mL cell wall-breaking buffer, were shaken vigorously in

the presence of acid-washed glass beads. The resultant homogenate was

centrifuged at 1,500g for 5 min at 4�C. The supernatant was removed, aliquoted,

and stored at 276�C. For sterol substrate specificity determination, the super-

natant was washed twice with 80 mM methyl-b-cyclodextrin (Sigma) for

removal of free sterols embedded in endoplasmic reticulum membrane (Rodal

et al., 1999). Protein concentration was measured using Bio-Rad Protein Assay

kit for final AtSAT1 activity calculation. Substrate specificity was determined

by a previously described method (Yang et al., 1997) with slight modification.

For fatty acid-CoA substrate specificity, a reaction mixture containing 200 mg

microsomal protein, 10 mg unlabeled lanosterol, 200 mg bovine serum albumin,

and 30 mg reduced glutathione in a final volume of 400 mL of 0.1 M potas-

sium phosphate buffer, pH 7.4, was sonicated for 5 min. One microliter of

[3H]lanosterol (1 mCi/mL, 10 mCi/mmol, American Radiolabeled Chemicals)

was added and incubated at 30�C with 100 rpm shaking for 30 min to allow

incorporation of sterols into the membrane. Sterols were suspended in reaction

buffer with the help of Triton-100 at a ratio of 30:1 (Tritus/sterol, w/w). The

reaction was initiated by adding 50 mL 0.4 mM acyl-CoA (C16:0, C16:1, C18:0,

C18:1, and C18:2, respectively). Reaction without exogenous acyl-CoA was

employed as control. The reaction mixture for sterol substrate preference

determination was essentially the same as that for acyl-CoA specificity assays

except that [14C]palmitoyl-CoA (50 mL, 10nCi/nmol, 180 mM) was utilized to

measure SE production. Reactions were terminated by the addition of 3 mL of

chloroform/methanol (2:1, v/v). Cholesterol oleate was added as a carrier. The

chloroform layer containing lipid was removed and dried under a N2 stream,

redissolved in 100 mL chloroform, and then applied to a J.T. baker Si250 PATLC

silica plate gel. After visualization with iodine, the bands corresponding to SE

were scraped off and subjected to scintillation counting. The specific activity of

AtSAT1 activity was determined as picomoles of acyl-CoA or lanosterol

converted to SE per milligram protein per minute (pmol mg21 min21).

L-a-dipalmitoyl PC (dipalmitoyl-1-a-[14C]PC, 55 mCi/mmol, 0.1 mCi/mL)

was used to examine if AtSAT1 was capable of using PC as fatty acyl donor.

The reaction mixture was essentially the same as that for sterol substrate

specificity analysis, except that 2 mL of the 14C-labeled PC in ethanol:toluene

(1:1) solution was added to the reaction mixture. A similar assay with

[14C]16:0-CoA (55mCi/mmol, 0.1 mCi/mL) as acyl donor was also performed

in parallel for comparison.

Generation of Transgenic Plants

A cDNA of AtSAT1 was PCR amplified with primers FP-XbaI (5#-CAA-

GAATCTAGAATGGCGAGTTTCATCAAGGCA-3#) and RP-KpnI (5#-GAT-

CACTCAAGTTACCACACACGGCAGGGTTA-3#), and ligated into the XbaI

Figure 8. A proposed role for AtSAT1 in the sterol biosynthesis path-way. The diagram accentuates the section where the conversion ofsqualene to sterol end products occurs. Solid lines denote single-stepenzymatic reactions and dashed lines represent multiple steps. Theopen arrows suggest the roles of AtSAT1 in sterol biosynthesis.

Chen et al.




and KpnI sites of pSE129 vector. The AtSAT1 sequence and the construct

integrity were verified by sequencing. The resulting gene expression cassette

in the transformation vector contained a canola (Brassica napus) seed-specific

napin promoter and Agrobacterium NOS terminator flanking the 5# and 3# ends

of the AtSAT1 cDNA, respectively. A single colony of Agrobacterium (GV3101)

carrying napin:AtSAT1 transformation vector was cultured in Luria-Bertani

medium containing 50 mg mL21 kanamycin and 25 mg mL21 gentamycin.

Plant transformation was performed through vacuum infiltration. Selection of

T0 transgenic seeds was carried on media containing one-half Murashige and

Skoog Gamborg medium, 0.8% (w/v) phytagar, 3% (w/v) Suc, 50 mg mL21

kanamycin, and 50 mg mL21 timentin.

qRT-PCR and Data Analysis

To quantify transcript levels, total RNA was extracted from siliques of

wild-type and transgenic T2 plants using Qiagen NEasy Plant Mini kit. One

microgram of DNase I-treated RNA was used as template for cDNA synthesis

(Invitrogen superscript First-Strand Synthesis kit). qRT-PCR was performed

on Applied Biosystem StepOne Real-Time PCR system in triplicates. The

b-actin-8 gene (AT1G49240) was used as endogenous standard for expression

level calculation. The cDNA samples were 50 times diluted for amplication of

SYBR green-labeled PCR fragments (Power SYBR Green PCR Master mix,

Applied Biosystems) by using gene-specific primers (forward 5#-CTTGA-

GATCATCCTTGCCGCCACAGC-3# and reverse 5#-AGTGATGTCGCTAGG-

TACGGCTTGTTG-3#) for AtSAT1. The expression of b-actin-8 was assayed

using forward primer 5#-AACAGCAGAACGGGAAATTGTGAGA-3# and

reverse primer 5#-TGGAGGAGCTGGTTTTCGAGGT-3#. The corresponding

amplicon size of b-actin-8 and AtSAT1 are 101 and 99 bp, respectively. The

cycling parameters included an initial DNA denaturation step at 95�C for

10 min, followed by 40 cycles of PCR with DNA denaturation at 95�C for 15 s,

and primer annealing and extension at 60�C for 1 min. qRT-PCR reactions with

RNA templates (minus RT) were used as controls to check for DNA contam-

ination or amplification of nonspecific products. Controls with no added

template were included for each primer pairs to ensure primer dimmer was

not interfering with amplification detection. qRT-PCR results were captured

and analyzed using software StepOne (ver. 1.0). The cycle number at which

the fluorescence passed the cycle threshold (CT), automatically determined by

the software StepOne (version 1.0), was used for AtSAT1 expression level quan-

titation. AtSAT1 relative expression levels in each cDNA sample were obtained

through normalization to b-actin-8 by the formula 22(CTAtSAT1 2 CTb-actin-8).

Seed Sterol Extraction and Quantification

Six to 8 mg of mature seeds from wild-type and AtSAT1 transgenic lines

were suspended in 2 mL of chloroform:methanol (2:1, v/v) containing 20 mg

cholesterol and 20 mg cholesteryl myristate as internal standards. The sus-

pension was probe sonicated, washed with 0.9% NaCl, and then vigorously

vortexed. The bottom chloroform layer was transferred into a new solvent-

washed tube. The aqueous layer was extracted twice with 2 mL of chloroform:

methanol (2:1, v/v), and the organic layers were combined and dried down

under N2 stream. The residue was dissolved in 200 mL hexane. SE and

free sterol were separated and collected through HPLC. The SE fraction was

saponified in 2 mL 7.5% KCl in 95% methanol and the resultant free sterols

were repeatedly extracted with hexane. Sterol-TMS derivatives were qualita-

tively and quantitatively analyzed following the same method for yeast sterol

analysis. Campesterol, b-sitosterol, cycloartenol, and methylenecycloartenol

were identified by searching NIST 2.0 mass spectrum library or standard

chemicals.

Sequence data from this article can be found in the GenBank/EMBL data

libraries under accession number NM_115056.

ACKNOWLEDGMENTS

We thank Dr. Stephen L. Sturley for providing the yeast strain SCY059 and

Dr. Sten Stymne for providing the quadruple disrupted yeast strain devoid of

TAG synthesis. We thank Mike Giblin and Darwin Reed for technical

guidance in HPLC separation of SEs, and Ms. Kim Bryce, project manager

for the National Research Council Canada-Crops for Enhanced Human

Health Program. Helpful discussion with Drs. Sten Stymne, Pat Covello, and

Mark Smith are acknowledged. This is National Research Council Canada

publication number 48407.

Received July 25, 2007; accepted September 6, 2007; published September 20,

2007.

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biosynthesis of phytosterol esters: identiï¬cation of a

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