| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
|
First published online May 11, 2007; 10.1104/pp.107.099432 Plant Physiology 144:1267-1277 (2007) © 2007 American Society of Plant Biologists OPEN ACCESS ARTICLE
Monoacylglycerols Are Components of Root Waxes and Can Be Produced in the Aerial Cuticle by Ectopic Expression of a Suberin-Associated Acyltransferase1,[W],[OA]Department of Plant Biology, Michigan State University, East Lansing, Michigan 48824
The interface between plants and the environment is provided for aerial organs by epicuticular waxes that have been extensively studied. By contrast, little is known about the nature, biosynthesis, and role of waxes at the root-rhizosphere interface. Waxes isolated by rapid immersion of Arabidopsis (Arabidopsis thaliana) roots in organic solvents were rich in saturated C18-C22 alkyl esters of p-hydroxycinnamic acids, but also contained significant amounts of both  - and  -isomers of monoacylglycerols with C22 and C24 saturated acyl groups and the corresponding free fatty acids. Production of these compounds in root waxes was positively correlated to the expression of sn-glycerol-3-P acyltransferase5 (GPAT5), a gene encoding an acyltransferase previously shown to be involved in aliphatic suberin synthesis. This suggests a direct metabolic relationship between suberin and some root waxes. Furthermore, when ectopically expressed in Arabidopsis, GPAT5 produced very-long-chain saturated monoacylglycerols and free fatty acids as novel components of cuticular waxes. The crystal morphology of stem waxes was altered and the load of total stem wax compounds was doubled, although the major components typical of the waxes found on wild-type plants decreased. These results strongly suggest that GPAT5 functions in vivo as an acyltransferase to a glycerol-containing acceptor and has access to the same pool of acyl intermediates and/or may be targeted to the same membrane domain as that of wax synthesis in aerial organs.
Suberin and cutin are ubiquitous extracellular lipid polymers found in plants. Each polymer is insoluble in organic solvents, but is found in association with solvent-extractable waxes. Cutin and its cuticular and epicuticular waxes form the cuticle layer covering all aerial organs of plants. The cuticle protects plants from biotic and abiotic stresses, limits gas and water exchange, and is likely involved in developmental processes during plant growth (Kolattukudy, 2001  ; Nawrath, 2006). Usually, cutin polyesters are composed largely of C16 and C18  -hydroxy fatty acid monomers, which often have midchain functionality, such as epoxy, secondary hydroxyl, or vicinal diol groups (Kolattukudy, 2001; Nawrath, 2002). Glycerol is also a monomer and has been shown to be esterified to -hydroxy fatty acids (Graça et al., 2002). In Arabidopsis (Arabidopsis thaliana) and oilseed rape (Brassica napus), the polyesters of leaf and stem epidermis and isolated cuticles are unusual in that they contain high proportions of ,-dicarboxylic acids and, particularly, that derived from linoleic acid (Bonaventure et al., 2004; Franke et al., 2006). Cuticular waxes are derived from very-long-chain (C22-C34) saturated fatty acids. Alkanes are common, but a wide variety of neutral lipids are found (Kolattukudy, 1980; Jetter et al., 2006). The chemical genetics of plant cuticular waxes has been extensively studied (Kunst and Samuels, 2003). Arabidopsis stem epicuticular waxes are dominated by nonacosane and its 14-hydroxy, 15-hydroxy, and 15-oxo derivatives, but contain smaller amounts of free fatty acids (FFAs), aldehydes, primary alcohols, and wax esters (Rashotte et al., 2001). Thus, despite their cooccurrence, there are few structural similarities between cuticular waxes and cutin monomers.
Suberin and its associated waxes form the suberin layer, which is often characterized by electron-translucent and electron-dense lamellae observed by transmission electron microscopy. Suberin is present in many external as well as internal tissues and has an important role in controlling water and solute fluxes. Its deposition is often induced by wounding or stress stimuli, thereby providing a barrier against pathogen invasion (Lulai and Corsini, 1998
A family of sn-glycerol-3-P acyltransferase (GPAT) genes has been identified in Arabidopsis, five members of which gave detectable enzyme activity when expressed heterologously in a gat1
We report here on the characterization of root waxes in the model plant Arabidopsis and show that they have a distinct composition and contain MAGs. We also demonstrate that the accumulation of these MAGs is positively correlated with the expression of GPAT5, suggesting a common pathway for the biosynthesis of aliphatic suberin and some of the suberin-associated waxes. Furthermore, we show that ectopic expression of GPAT5 under the control of the cauliflower mosaic virus (CaMV) 35S promoter leads to production of
MAGs Are Components of Arabidopsis Root Waxes
Stem and leaf surface waxes are operationally defined as the lipid material extracted by quickly dipping these organs into chloroform. Using the chloroform-dipping procedure, we examined the presence of surface lipids in roots of 7-week-old Arabidopsis plants. At this stage, the roots have undergone considerable secondary growth and the epidermis, cortex, and endodermis have been cast off, leaving the newly formed suberin-rich periderm as the root outer cell layers (Dolan and Roberts, 1995
The composition of waxes of Arabidopsis roots (Fig. 2C) was very distinct from that of Arabidopsis aerial parts (Rashotte et al., 2001 ) and significantly different from that reported for the subterranean storage organs of seven species, including Crucifers (Espelie et al., 1980). In Arabidopsis roots, esters of p-coumaric, caffeic, and ferulic acids with C18-C22 saturated fatty alcohols were the major component (47% [w/w]). Primary alcohols were present in significant amounts (10% [w/w]) with a chain length profile very similar to those esterified to hydroxycinnamic acids. This profile distinguishes them from the C26-C30 primary alcohols of Arabidopsis cuticular waxes in aerial organs (Rashotte et al., 2001). Substantial amounts of sterols and FFA were also observed (approximately 15% [w/w] each). Components that dominate aerial waxes, namely, nonacosane and its 15(14)-hydroxy and 15-oxo derivatives, were only minor contributors to root waxes (approximately 5% [w/w]). We will collectively call these major components found in the cuticular waxes of wild-type plants standard cuticular waxes.
The most unusual feature of Arabidopsis root waxes was the presence of both
Kinetics of chloroform-dipping extractions showed that, after a 10-s dip sufficient to extract 100% of stem waxes, the only root wax component completely recovered was alkanes (Fig. 2B). Other root wax components were extracted more slowly, most noticeably primary alcohols. Also, these kinetics of extraction showed that increasing dipping time did not allow the recovery of additional amounts of C16-C20 FFAs and sterols, although more of these lipids could be recovered by grinding the roots (data not shown). This indicated that there were distinct intracellular and wax fractions for C16-C20 FFAs and sterols.
Previously we noted that the T-DNA knockout mutant gpat5 had large reductions in suberin-like aliphatic monomers released by transesterification of the residual fraction of seeds and roots remaining after extensive delipidation (Beisson et al., 2007
However, analyses of the root waxes of wild type, two 35S::GPAT5-expressing lines (OE-1 and OE-2), and two knockout mutant lines (gpat5-1 and gpat5-2) revealed strong positive correlation between GPAT5 expression and the concentration of MAGs (Fig. 4
). An approximately 50% (w/w) reduction was observed in the mutants compared to a 70% (w/w) increase in the 35S::GPAT5-expressing lines. Both positional isomers of MAG were affected to a similar degree (data not shown). A positive correlation was also observed for C22-C30 FFAs (Fig. 4). Other root wax components were not significantly changed with altered GPAT5 expression (primary alcohols, sterols) or only weakly reduced by GPAT5 knockout and not increased by overexpression (alkyl ferulates; data not shown). The simple explanation for these results is that GPAT5 activity is limiting for the production of MAGs in waxes from wild-type roots. Null mutations of GPAT5 may not completely deplete MAGs from the suberin-associated waxes because of gene redundancy within the GPAT family. Indeed, all other GPATs, except one, are expressed in roots (Beisson et al., 2007
GPAT5 Overexpression Produces MAGs as Novel Components of Cuticular Waxes
To further demonstrate the link between GPAT5 and the production of MAGs, we characterized the stem cuticular waxes of the 35S::GPAT5-expressing plants. Scanning electron microscopy (SEM) of the stem surface showed a large reduction in wax crystal density compared to wild type (Fig. 5
). Stems of wild-type plants were covered primarily with columnar shaped crystals, although rods, tubes, vertical plates, and dendritic- and umbrella-like structures were also visible (Rashotte and Feldmann, 1998
Wax analysis identified both - and -isomers of MAGs, with saturated C22-C30 acyl groups, as novel components of the cuticular waxes from stems of 35S::GPAT5-expressing plants, constituting up to 20% (w/w) of the total wax load (Fig. 6
, inset). Whereas FFAs are only minor components of standard waxes (<5%), greatly elevated levels of C22-C30 FFAs (approximately 50% [w/w]) were also noted in the transgenes. The accumulated MAGs have a slightly altered acyl chain length distribution when compared to those present in the root waxes of Arabidopsis plants (Figs. 2 and 6). Tetracosanoic acid was the major FFA and acyl group in MAGs of stems. Another noticeable phenotype was a shift in distribution of the saturated wax esters, which constitute about 7% to 9% (w/w) of total wax load, from C40-C46 in wild type to C40-C54 in the transgenes. In wild-type wax esters, the predominant acyl group was C16, but in the transgene there was an additional contribution largely from the C22 acyl group (data not shown). The novel MAG and FFA products observed in stem waxes for 35S::GPAT5-expressing plants were also noted for surface waxes from leaves, siliques, and seeds (Table I
).
By analyzing the lipids in both the 30-s chloroform dip fraction and the remaining stem tissue, we demonstrated that two-thirds of very-long-chain fatty acids (VLCFAs; total acyl groups from MAGs and FFAs) were immediately extractable (Fig. 7A ). This compares with wild-type surface waxes, which are completely extracted, and with intracellular lipids (i.e. C16-C20 fatty acids), which are not extracted. Furthermore, the chain length distribution of FFA and MAG in these two fractions was similar, although the longer chain MAG species did appear to be preferentially found on the surface (Fig. 7B). Epidermal peel experiments confirmed that the MAGs plus FFAs remaining after the short chloroform dip are largely epidermal (>90% [w/w]), although whether their site of deposition is extracellular, intracellular, or both remains to be determined. Taken together, these findings show that ectopic expression of GPAT5 in the aerial parts of plants results in novel MAG, wax ester, and FFA products and that these products are largely secreted onto the plant surface, where they radically alter the crystal morphology of the epicuticular wax layer. Although lysophosphatidic acid might be expected to be the first product of the GPAT5 reaction (Zheng et al., 2003 ), analyses of the short chloroform dip and lipids extracted from residual stems failed to reveal any lysophosphatidic acid containing VLCFAs. Also, an analysis of stem cutin showed that no additional long-chain C22-C30 fatty acids were incorporated into this polyester (Supplemental Fig. S2).
GPAT5 Overexpression Reduces Accumulation of Wild-Type Cuticular Wax Components In addition to the appearance of novel components in 35S::GPAT5 overexpression lines, there was a reduction in the load of some standard wax components, most dramatically for the C29 alkane component of the decarboxylation/decarbonylation pathway, whereas C26-C30 primary alcohols remained approximately constant (Fig. 6). To confirm the reduction in standard waxes, we analyzed the accumulation of C22-C30 fatty acids (from MAGs plus FFAs) in stems from 6-week-old plants for 18 independent T2 35S::GPAT5 lines. An inverse relationship was observed between the amount of standard waxes present in each line and the amount of newly formed VLCFAs due to GPAT5 expression (Fig. 8 ).
Surface MAGs Are Produced in 35S::GPAT5-Expressing Tobacco Plants
Although the main focus of this study was Arabidopsis, some experiments conducted with tobacco (Nicotiana tabacum) suggest that the results seen with Arabidopsis may be more broadly applicable across the plant kingdom. First, when wild-type tobacco roots from 8-week-old plants were subjected to the rapid chloroform dip protocol and the lipids analyzed by GC-MS, trace amounts of docosanoyl and tetracosanoyl glycerols were detected. These were largely the
In this study, the composition of Arabidopsis root waxes is reported and MAGs with C22-C24 saturated acyl groups are identified as components of these waxes. This phytochemical analysis is complemented by characterization of Arabidopsis plants expressing the acyltransferase GPAT5 of the aliphatic suberin biosynthesis pathway under the control of the 35S promoter. The results show that (1) GPAT5 is not only involved in the synthesis of aliphatic suberin polymer (Beisson et al., 2007 ), but also in the synthesis of the MAGs present in the root waxes; (2) GPAT5 is an acyltransferase acting on a glycerol-based acceptor; and (3) the ectopic expression of this gene can lead to the production of MAGs in cuticular waxes of aerial organs. The significance of these results in terms of the relationships of aliphatic suberin to root waxes, the functional roles of GPAT5, and its utility in engineering cuticular waxes are discussed.
The presence in Arabidopsis root waxes of free and esterified saturated fatty acids and alcohols with chain lengths of C18-C22 and of hydroxycinnamate and glycerol building blocks (Fig. 2C) shows that, at least in Arabidopsis, root waxes have characteristics common to the monomers released upon depolymerization of root suberin (Franke et al., 2006
Concerning the location of suberin-associated waxes, it has been suggested that they are found within the electron-translucent region of the suberin lamellae (Soliday et al., 1979
The view that suberin-associated waxes are physically associated with and metabolically related to suberin is further substantiated genetically by the analysis of MAGs in both knockout and ectopic expression lines of GPAT5. The level of MAG and associated FFA in root waxes is strongly dependent on GPAT5 expression. Furthermore, GPAT5 controls the level of suberin monomers in young roots during development of the specialization zone (Beisson et al., 2007
Although GPAT5 is clearly involved in deposition of both waxes and polyesters, the metabolic relationships between the two product classes is not understood. In 7-week-old Arabidopsis roots, we observed about 350 µg/g fresh weight of suberin aliphatic monomers released by depolymerization (transesterification), and a similar amount of suberin-associated waxes (approximately 360 µg/g fresh weight). What is not known is the relative timing of wax and suberin deposition in peridermal tissues. The MAG, FFA, primary alcohols, and alkyl ferulates might be suberin precursors that have not been polymerized or they might simply be synthesized on the same pathway as suberin; or, with the exception of the alkyl ferulates, they might be derived by some postdeposition hydrolytic events. A comparison can be made with the inhibition of fatty acyl elongation in green-lint cotton fibers, which differentially reduces waxes over suberin (Schmutz et al., 1996
Because we observed MAG in the cuticular waxes in 35S::GPAT5 plants, whereas it is completely absent in the wild type, a principal conclusion of this study is that GPAT5 acts in vivo as an acyltransferase for a glycerol-containing acceptor. GPAT5 is annotated as a sn-glycerol-3-P acyl-CoA acyltransferase and has been shown to have such activity in yeast (Zheng et al., 2003
Concerning acyl donor specificity of GPAT5, a preference for very long acyl chains could be inferred based on strong reduction of such monomers in polyesters of the gpat5 mutants (Beisson et al., 2007
A major biosynthetic pathway of epidermal cells is the elongation of saturated fatty acyl-CoAs to produce precursors for surface waxes (Kunst and Samuels, 2003 Ectopic expression of GPAT5 driven by the CaMV 35S promoter causes a decrease in the levels of standard cuticular waxes in leaf, stem, and siliques (Table I; Fig. 8). However, this reduction is not observed in seeds, nor do we see a reduction in other long-chain aliphatics in roots. These differences may simply derive from relative differences in promoter strength in different tissues, differences that may vary between lines. However, a large number of other explanations may be invoked because we know almost nothing about relative pool sizes of intermediates, feedback inhibition, substrate channeling, temporal separation of accumulation of individual root wax components, and protein stability.
Previously, alterations in the content and composition of surface waxes and polyesters have been made largely through gene deletion (Kunst and Samuels, 2003
Plant Materials and Growth Conditions
All Arabidopsis (Arabidopsis thaliana) plants were in the Col-0 genetic background. The gpat5-1 and gpat5-2 mutants were previously isolated and characterized (Beisson et al., 2007
Genomic DNA was prepared from Arabidopsis leaf tissue using a plant mini DNA kit according to the manufacturer's instructions (Qiagen). Genomic DNA sequences encoding the GPAT5 gene (At3g11430) were amplified by PCR using forward primer (5'-CACACTCTAGAATGGTTATGGAGCAAGC-3') and reverse primer (5'-CACACGAGCTCTCAATGGAGACAAGG-3'). The PCR product was initially cloned into pGEM-T Easy vector, and then subcloned as an XbaI-SacI fragment into binary vector pBI121 to replace the GUS gene. The construct (35S::GPAT5) was introduced into Agrobacterium tumefaciens strain C58C1 for Arabidopsis vacuum infiltration (Bechtold et al., 1993
To view epicuticular waxes, sections of stems were treated in 1% (w/v) osmium tetroxide vapor for 24 h, air dried for 3 d, mounted onto standard aluminum stubs for JEOL SEM, and then sputter coated with around 30 nm of gold using an EMSCOPE SC-500 sputter coater. The images were taken with a JEOL 6400V scanning electron microscope.
Sudan Red 7B (Sigma) was prepared as a 0.05% (w/v) solution in PEG400:glycerol (1:1 [v/v]; Brundrett et al., 1991
Stems were dipped in chloroform for 30 s, the solvent evaporated under a stream of N2 gas, and tricosane, tricosanoic acid, monoheptadecanoin, and tridodecanoin added as internal standards. The waxes were derivatized by heating at 110°C for 10 min in pyridine:BSTFA (1:1 [v/v]). The silylated sample was analyzed by GC using a 30-m DB5-ht capillary column temperature programmed at 10°C min–1 to 370°C. Eluting components were quantified based on uncorrected peak areas from integrated flame ionization detector ion current. For molecular identification, a Hewlett-Packard 5890 GC-coupled MSD 5972 mass analyzer was used with the mass analyzer set in electron impact mode (70 eV) and scanning from 40 to 700 atomic mass units. Tobacco leaf epicuticular wax analysis was conducted as above, except that leaves were dipped in dichloromethane instead of chloroform (Severson et al., 1984
Arabidopsis roots were carefully and thoroughly washed in distilled water, blotted, then air dried at 50°C for 30 min, and dipped in chloroform for 1 min, unless otherwise stated. The extracts were passed through a glass wool-plugged column and evaporated to dryness under a stream of N2 gas. The waxes were derivatized and analyzed as described above for cuticular waxes. Due to the complex architecture of the roots, it is not practical to calculate the root wax load based on surface area; therefore, we report root wax load as micrograms per gram fresh weight. Inspection of root biomass of wild-type, mutant, and 35S::GPAT5 overexpression lines showed no obvious differences in morphology, nor did the average peridermal root diameter measured close to the crown vary significantly between lines, suggesting that per gram fresh-weight units are approximately proportional to surface area units.
Fatty acid content and composition of Arabidopsis tissues was analyzed directly by acidic transmethylation according to Li et al. (2006)
Statistical tests were performed using Microsoft Excel and Analyze-it (version 1.73) software.
The following materials are available in the online version of this article.
We thank Katrin Weber (Department of Plant Biology, Michigan State University) for assisting with plant transformations and Ewa Danielewics (Center for Advanced Microscopy, Michigan State University) for SEM analyses. Received March 12, 2007; accepted May 4, 2007; published May 11, 2007.
1 This work was supported by the National Research Initiative of the U.S. Department of Agriculture Cooperative State Research, Education, and Extension Service (grant no. 2005–35318–15419).  The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Mike Pollard (pollard9{at}msu.edu).
[W] The online version of this article contains Web-only data.
[OA] Open Access articles can be viewed online without a subscription. www.plantphysiol.org/cgi/doi/10.1104/pp.107.099432 * Corresponding author; e-mail pollard9{at}msu.edu; fax 517–353–1926.
Aharoni A, Dixit S, Jetter R, Thoenes E, van Arkel G, Pereira A (2004) The SHINE clade of AP2 domain transcription factors activates wax biosynthesis, alters cuticle properties, and confers drought tolerance when overexpressed in Arabidopsis. Plant Cell 16: 2463–2480 Bechtold N, Ellis J, Pelletier G (1993) In planta Agrobacterium-mediated gene transfer by infiltration of adult Arabidopsis thaliana plants. C R Acad Sci III 316: 1194–1199 Beisson F, Li Y, Bonaventure G, Pollard M, Ohlrogge J (2007) The acyltransferase GPAT5 is required for synthesis of suberin in seed coat and root of Arabidopsis. Plant Cell 19: 351–368 Bernards MA (2002) Demystifying suberin. Can J Bot 80: 227–240[CrossRef] Bernards MA, Lewis NG (1992) Alkyl ferulates in wound healing potato tubers. Phytochemistry 31: 3409–3412[CrossRef][ISI][Medline] Bernards MA, Lopez ML, Zajicek J (1995) Hydroxycinnamic acid-derived polymers constitute the polyaromatic domain of suberin. J Biol Chem 270: 7382–7386 Bonaventure G, Beisson F, Ohlrogge J, Pollard M (2004) Analysis of the aliphatic monomer composition of polyesters associated with Arabidopsis epidermis: occurrence of octadeca-cis-6, cis-9-diene-1,18-dioate as the major component. Plant J 40: 920–930[CrossRef][ISI][Medline] Broun P, Poindexter P, Osborne E, Jiang CZ, Reichmann J (2004) WIN1, a transcriptional activator of epidermal wax accumulation in Arabidopsis. Proc Natl Acad Sci USA 101: 4706–4711 Brundrett MC, Kendrick B, Peterson CA (1991) Efficient lipid staining in plant material with sudan red 7B or fluorol yellow 088 in polyethylene glycol-glycerol. Biotech Histochem 66: 111–116[ISI][Medline] Dolan L, Roberts K (1995) Secondary thickening in roots of Arabidopsis thaliana: anatomy and cell surface changes. New Phytol 131: 121–128[CrossRef][ISI] Espelie KE, Sadek NZ, Kolattukudy PE (1980) Composition of suberin-associated waxes from the subterranean storage organs of seven plants, parsnip, carrot, rutabaga, turnip, red beet, sweet potato and potato. Planta 148: 468–476[CrossRef][ISI] Franke R, Briesen I, Wojciechowski T, Faust A, Yephremov A, Nawrath C, Schreiber L (2006) Apoplastic polyesters in Arabidopsis surface tissues—a typical suberin and a particular cutin. Phytochemistry 66: 2643–2658[CrossRef][ISI] Graça J, Pereira H (1997) Cork suberin: a glyceryl based polyester. Holzforschung 51: 225–234[ISI] Graça J, Pereira H (1999) Glyceryl-acyl and aryl-acyl dimers in Pseudotsuga menziesii bark suberin. Holzforschung 53: 397–402[CrossRef][ISI] Graça J, Pereira H (2000) Suberin structure in potato periderm: glycerol, long-chain monomers, and glyceryl and feruroyl dimers. J Agric Food Chem 48: 5476–5483[CrossRef][ISI][Medline] Graça J, Santos S (2006) Linear aliphatic dimeric esters from cork suberin. Biomacromolecules 7: 2003–2010[CrossRef][ISI][Medline] Graça J, Schreiber L, Rodrigues J, Pereira H (2002) Glycerol and glyceryl esters of omega-hydroxyacids in cutins. Phytochemistry 61: 205–215[CrossRef][ISI][Medline] Gunstone FD (1967) An Introduction to Chemistry and Biochemistry of Fatty Acids and Their Glycerides. Chapman Hall, London, pp 141 Holloway PJ (1982) Structure and histochemistry of plant cuticular membranes: an overview. In DF Cutler, KF Alvin, CE Price, eds, The Plant Cuticle. Academic Press, London, pp 1–32 Jetter R, Kunst L, Samuels L (2006) Composition of plant cuticular waxes. In M Riederer, C Müller, eds, Biology of the Plant Cuticle. Annual Plant Reviews, Vol 23. Blackwell Scientific Publishers, Oxford, pp 145–175 Kolattukudy PE (1980) Cutin, suberin and waxes. In PK Stumpf, ed, The Biochemistry of Plants—A Comprehensive Treatise, Vol 4. Academic Press, New York, pp 571–645 Kolattukudy PE (2001) Polyesters in higher plants. Adv Biochem Eng Biotechnol 71: 1–49[Medline] Kunst L, Samuels AL (2003) Biosynthesis and secretion of plant cuticular wax. Prog Lipid Res 42: 51–80[CrossRef][ISI][Medline] Li Y, Beisson F, Pollard M, Ohlrogge J (2006) Oil content of Arabidopsis seeds: the influence of seed anatomy, light and plant-to-plant variation. Phytochemistry 67: 904–915[CrossRef][ISI][Medline] Lulai EC, Corsini DL (1998) Differential deposition of suberin phenolic and aliphatic domains and their roles in resistance to infection during potato tuber (Solanum tuberosum L.) wound-healing. Physiol Mol Plant Pathol 53: 209–222[CrossRef] Moire L, Schmutz A, Buchala A, Yan B, Stark RE, Ryser U (1999) Glycerol is a suberin monomer: new experimental evidence for an old hypothesis. Plant Physiol 119: 1137–1146 Molina I, Bonaventure G, Ohlrogge J, Pollard M (2006) The lipid polyester composition of Arabidopsis thaliana and Brassica napus seeds. Phytochemistry 67: 2597–2610[CrossRef][ISI][Medline] Murashige T, Skoog F (1962) A revised medium for rapid growth and bio-assays with tobacco tissue cultures. Physiol Plant 15: 473–497[CrossRef] Murphy RC (1993) Mass Spectroscopy of Lipids—Handbook of Lipid Research. Plenum Press, New York, pp 206 Nawrath C (2002) The biopolymers cutin and suberin. In CR Somerville, EM Meyerowitz, eds, The Arabidopsis Book. American Society of Plant Biologists, Rockville, MD, pp 1–14 Nawrath C (2006) Unraveling the complex network of cuticular structure and function. Curr Opin Plant Biol 9: 281–287[CrossRef][ISI][Medline] Rashotte AM, Feldmann KA (1998) Correlations between epicuticular wax structures and chemical composition in Arabidopsis thaliana. Int J Plant Sci 159: 773–779[CrossRef][ISI] Rashotte AM, Jenks MA, Feldmann KA (2001) Cuticular waxes on eceriferum mutants of Arabidopsis thaliana. Phytochemistry 57: 115–123[CrossRef][ISI][Medline] Rogers SG, Horsch RB, Fraley RT (1986) Gene transfer in plants: production of transformed plants using Ti plasmid vectors. Methods Enzymol 118: 627–640[CrossRef][ISI] Santos S, Graça J (2006) Glycerol- Schmutz A, Amrhein N, Ryser U (1993) Caffeic acid and glycerol are constituents of the suberin layers in green cotton fibres. Planta 189: 453–460[CrossRef][ISI] Schmutz A, Buchala A, Ryser U (1996) Changing the dimensions of suberin lamellae of green cotton fibers with a specific inhibitor of the endoplasmic reticulum-associated fatty acid elongases. Plant Physiol 110: 403–411[Abstract] Schmutz A, Jenny T, Ryser U (1994) A caffeoyl-fatty acid glycerol ester from wax associated with green cotton fiber suberin. Phytochemistry 36: 1343–1346[CrossRef][ISI] Schreiber L, Franke R, Hartmann K (2005) Wax and suberin development of native and wound periderm of potato (Solanum tuberosum L.) and its relation to peridermal transpiration. Planta 220: 520–530[CrossRef][ISI][Medline] Severson RF, Arrendale RF, Chortyk OT, Johnson AW, Jackson DM, Gwynn GR, Chaplin JF, Stephenson MG (1984) Quantitation of the major cuticular components from green leaf of different tobacco types. J Agric Food Chem 32: 566–570[CrossRef][ISI] Soliday CL, Kolattukudy PE, Davis RW (1979) Chemical and ultrastructural evidence that waxes associated with the suberin polymer constitute the major diffusion barrier to water vapor in potato tuber (Solanum tuberosum L.). Planta 146: 607–614[CrossRef][ISI] Suh MC, Samuels AL, Jetter R, Kunst L, Pollard M, Ohlrogge J, Beisson F (2005) Cuticular lipid composition, surface structure, and gene expression in Arabidopsis stem epidermis. Plant Physiol 139: 1649–1665 Zhang JY, Broeckling CD, Blancaflor EB, Sledge MK, Sumner LW, Wang ZY (2005) Overexpression of WXP1, a putative Medicago truncatula AP2 domain-containing transcription factor gene, increases cuticular wax accumulation and enhances drought tolerance in transgenic alfalfa (Medicago sativa). Plant J 42: 689–707[CrossRef][ISI][Medline] Zheng Z, Xia Q, Dauk M, Shen W, Selvaraj G, Zou J (2003) Arabidopsis AtGPAT1, a member of the membrane-bound glycerol-3-phosphate acyltransferase gene family, is essential for tapetum differentiation and male fertility. Plant Cell 15: 1872–1887 This article has been cited by other articles:
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
TOP
ABSTRACT
RESULTS
strain of yeast (Saccharomyces cerevisiae; Zheng et al., 2003
) bond of the glycerol backbone usually produces the base peak [M-CH2OTMSi]+. The m/z = 483 ion in 
-statistic = –0.63; two-tailed P = 0.0002 with normal approximation and correction for ties; n = 18).
90% [w/w] of total MAGs), whereas FFAs accounted for <5% of total MAG mass. Both straight and branched-chain acyl groups were present in the MAGs. This parallels the presence of iso- and anteiso-C27-C33 branched-chain hydrocarbons reported for tobacco epicuticular waxes (Severson et al., 1984
