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Keywords:

  • cuticular waxes;
  • fatty acid elongation;
  • chain lengths;
  • esters;
  • hydrocarbons;
  • industrial products

Summary

  1. Top of page
  2. Summary
  3. Introduction
  4. Plant cuticular wax composition and biosynthesis
  5. Current applications and commercial sources of waxes
  6. Metabolic engineering for wax ester production
  7. Potential for biotechnological alkane production
  8. Conclusions and perspectives
  9. Acknowledgements
  10. References

Due to their unique physical properties, waxes are high-value materials that are used in a variety of industrial applications. They are generated by chemical synthesis, extracted from fossil sources, or harvested from a small number of plant and animal species. As a result, the diversity of chemical structures in commercial waxes is low and so are their yields. These limitations can be overcome by engineering of wax biosynthetic pathways in the seeds of high-yielding oil crops to produce designer waxes for specific industrial end uses. In this review, we first summarize the current knowledge regarding the genes and enzymes generating the chemical diversity of cuticular waxes that accumulate at the surfaces of primary plant organs. We then consider the potential of cuticle biosynthetic genes for biotechnological wax production, focusing on selected examples of wax ester chain lengths and isomers. Finally, we discuss the genes/enzymes of cuticular alkane biosynthesis and their potential in future metabolic engineering of plants for the production of renewable hydrocarbon fuels.


Introduction

  1. Top of page
  2. Summary
  3. Introduction
  4. Plant cuticular wax composition and biosynthesis
  5. Current applications and commercial sources of waxes
  6. Metabolic engineering for wax ester production
  7. Potential for biotechnological alkane production
  8. Conclusions and perspectives
  9. Acknowledgements
  10. References

Primary plant surfaces are impregnated with waxes produced by epidermal cells (Riederer and Müller, 2006). These cuticular waxes are complex mixtures of C20–C34 straight-chain aliphatics derived from very-long-chain fatty acids (VLCFAs), and in certain plant species also include alicyclic and aromatic compounds such as triterpenoids, alkaloids, phenylpropanoids and flavonoids. Plant cuticular waxes serve as a protective barrier against water loss, UV light, pathogens and insects. In addition, they are valuable raw materials for a variety of industrial applications. Wax mixtures derived from different plant sources have unique chemical compositions that determine their physical properties, and therefore their potential applications and industrial value.

At present, cuticular waxes are commercially harvested from only a small number of plant species, so the structural diversity of their wax constituents is limited. In addition, these plant species are mostly grown in tropical areas and are agronomically not well suited to commercial production. These apparent shortcomings of plant surface wax production can be circumvented through genetic engineering approaches using established high-yielding oil crops as a platform. By introducing wax biosynthetic pathways into oilseeds, waxes with optimal chemical compositions for various specialty markets could be produced, including high-value lubricants, cosmetics and pharmaceuticals, as well as high-energy fuels.

In this review, we present the chemical diversity of plant cuticular wax mixtures and summarize our understanding of the biosynthetic pathways involved in generating this diversity; provide an overview of commercial sources and uses of waxes, and of current limitations of wax production; discuss how engineering of wax biosynthetic pathways in target crops might be exploited for the production of novel waxes with specific chain-length distributions in oilseeds; and describe how wax biosynthetic pathways can be used in metabolic engineering of plants for the production of hydrocarbon biofuels. This information complements recent reviews that have focused on the chemical composition (Jetter et al., 2006), biosynthesis (Kunst et al., 2006) and biological functions of plant cuticular waxes (Bargel et al., 2006; Riederer and Müller, 2006).

Plant cuticular wax composition and biosynthesis

  1. Top of page
  2. Summary
  3. Introduction
  4. Plant cuticular wax composition and biosynthesis
  5. Current applications and commercial sources of waxes
  6. Metabolic engineering for wax ester production
  7. Potential for biotechnological alkane production
  8. Conclusions and perspectives
  9. Acknowledgements
  10. References

Cuticular wax composition varies between different species and organs essentially in two respects: chain-length distribution and compound class composition (Jetter et al., 2006). This diversity is established during wax biosynthesis in epidermal cells (Kunst et al., 2006), and involves two types of pathways: those for the elongation of fatty acid wax precursors to assorted chain lengths and those for modifying them into wax components with various functional groups. Aliphatic compound classes ubiquitously present in cuticular wax mixtures are alkanes, primary alcohols, aldehydes and fatty acids ranging in chain length between 20 and 34 carbons, as well as alkyl esters up to C60 in length (Figure 1).

image

Figure 1.  Structures of major components occurring in plant cuticular wax mixtures. (a) Ubiquitous compound classes lacking functional groups (alkanes) or with primary functional groups. Typically, series of compounds with wide ranges of chain lengths are present in these classes. n and m indicate the number of methylene (CH2) groups, and can range from 18 to 32. (b) Wax constituents with secondary functional groups accumulate to high concentrations in the wax of certain plant species, usually with very narrow chain-length and isomer distributions. Typical chain lengths and isomers are shown for selected combinations of hydroxyl and carbonyl functionalities.

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The cuticular waxes from many plant species comprise roughly equal amounts of the various compound classes, with no particular class predominating. For example, alkanes, aldehydes, primary alcohols, fatty acids and alkyl esters each contribute 9–42% of the leaf wax of Zea mays (Bianchi et al., 1984). In contrast, the wax mixtures from many other plant species contain high percentages of a single compound class. Hordeum vulgare leaf wax, for example, contains 89% of primary alcohols, together with only 0.2–9% of alkanes, aldehydes, fatty acids and alkyl esters (Giese, 1975).

Compound chain length

Variation in the chain length of wax compounds is generated during synthesis of VLCFA wax precursors. This process involves several enzyme complexes in various cellular compartments. The first phase, the de novo fatty acid synthesis of C16 and C18 acyl chains, is catalysed by the soluble fatty acid synthase (FAS) complex localized in the plastid stroma (Ohlrogge and Browse, 1995; Ohlrogge et al., 1993), and proceeds through a cycle of four reactions utilizing intermediates attached to acyl carrier protein (ACP). In each cycle, comprising the condensation of a C2 moiety originating from malonyl ACP to acyl ACP, the reduction of β-ketoacyl ACP, the dehydration of β-hydroxyacyl ACP and the reduction of trans2–enoyl ACP, the acyl chain is extended by two carbons. Three different FAS complexes participate in the production of C18 fatty acids in the plastid. They differ in their β-ketoacyl-acyl carrier protein synthase (KAS) condensing enzymes, which have strict acyl chain-length specificities: KASIII (C2–C4; Clough et al., 1992), KASI (C4–C16) and KASII (C16–C18; Shimakata and Stumpf, 1982). The two reductases and the dehydratase have no particular acyl chain-length specificity and are shared by all three plastidial elongation complexes (Stumpf, 1984).

The second phase (Figure 2), the extension of the C16 and C18 fatty acids to VLCFA chains, is carried out by fatty acid elongases (FAE; von Wettstein-Knowles, 1982), multienzyme complexes bound to the endoplasmic reticulum membrane (Kunst and Samuels, 2003; Xu et al., 2002; Zheng et al., 2005). To reach the ER-associated fatty acid elongation sites, saturated C16 and C18 acyl groups must be hydrolysed from the ACP by an acyl ACP thioesterase, exported from the plastid, and esterified to CoA. Two classes of acyl ACP thioesterases, designated FATA and FATB, have been described in plants. The FATA class exhibits a strong preference for 18:1 ACP in vitro, while the FATB thioesterases predominantly use saturated fatty acids (Voelker, 1996). The involvement of the FATB thioesterase in cuticular wax biosynthesis has been confirmed by analyses of the Arabidopsis fatb mutant, which exhibits a major reduction in its wax load (Bonaventure et al., 2003). The specifics of fatty acid export from the plastid, CoA esterification and transport to the ER are not well understood. Fatty acids released from ACP by a thioesterase in the plastid undergo conversion to acyl CoAs by a long-chain acyl CoA synthetase (LACS) in the outer envelope membrane. Of the nine LACS genes annotated in the Arabidopsis genome (Shockey et al., 2002), only one, LACS9, has been demonstrated to encode a plastid envelope enzyme (Schnurr et al., 2002). However, loss of function of LACS9 does not result in reduced export of acyl groups from the chloroplast, or a wax-deficient phenotype (Schnurr et al., 2002), suggesting that the LACS isozyme primarily responsible for CoA esterification of fatty acids en route to wax biosynthesis has yet to be identified. Movement of the fatty acyl group from the thioesterase to LACS has been proposed to occur by some type of facilitated diffusion (Koo et al., 2004), but the exact mechanism of transfer is not known. An alternative model for fatty acid export from the plastid was recently suggested by Bates et al. (2007). Their radiolabelling studies revealed that 16:0 and 18:1 fatty acids synthesized de novo in the plastid can be incorporated into phosphatidylcholine (PC), perhaps by direct acylation of lyso-PC. The acyl groups removed from PC by acyl editing may then be fed into the acyl CoA pool. However, mechanistic details and the relevance of this process for epidermal wax formation have not been established.

image

Figure 2.  Wax biosynthetic pathways. Repeated cycles of four enzymatic steps first elongate acyl CoA precursors. They are then modified by one of (up to) five different reactions into various compound classes. Preferred chain lengths are indicated by numbers. Characterized enzymes catalysing key biosynthetic steps are shown in blue (CER6, condensing enzyme=β-ketoacyl CoA synthase; KCR, β-ketoacyl CoA reductase; dehydratase, β-hydroxyacyl CoA dehydratase; CER10, enoyl CoA reductase; CER4, fatty acyl CoA reductase; WSD1, wax ester synthase; MAH1, mid-chain alkane hydroxylase).

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Translocation of fatty acids to the ER, where additional acyl chain elongation and modification of VLCFAs to diverse aliphatic wax components take place, appears to involve plastid-associated membranes (PLAMs; Andersson et al., 2007). Physical manipulation of GFP-labelled ER strands, using laser scalpels and optical tweezers, experimentally verified the intimate connection between the plastid and the ER of Arabidopsis leaf protoplasts. Therefore, PLAMs have been proposed to be major routes for lipid transfer between the two organelles.

Elongation of C16 and C18 fatty acids to VLCFAs involves cycles of four consecutive enzymatic reactions analogous to those of the FAS (Figure 2), and results in a two-carbon extension of the acyl chain per cycle. The chain lengths of aliphatic wax components are typically in the range of 20–34 carbons, thus multiple elongation cycles are needed to extend the acyl chain to its final length. The differential effects of inhibitors on incorporation of radiolabelled precursors into wax components of various chain lengths, and analyses of mutants with defects in fatty acid elongation, demonstrated that sequential acyl chain extensions are carried out by several distinct FAEs with unique substrate chain-length specificities (von Wettstein-Knowles, 1993). Specificity of each elongation reaction resides in the condensing enzyme of the FAE complex (Lassner et al., 1996; Millar and Kunst, 1997). Consistent with the requirement for fatty acyl precursors of diverse chain lengths for the synthesis of cuticular waxes, a family of 21 FAE condensing enzyme-like sequences has been identified in the A. thaliana genome (Dunn et al., 2004). An unrelated ELO-like gene family of putative condensing enzymes, related to the Saccharomyces cerevisiae condensing enzymes ELO1, ELO2 and ELO3, has also been annotated (Dunn et al., 2004). It is not known how many of these putative condensing enzymes participate in wax production and how many different condensing enzymes are needed for the elongation of a C18 to a C34 fatty acyl CoA, as single condensing enzymes may catalyse multiple elongation steps. The only wax-specific condensing enzyme characterized to date is CER6 (Fiebig et al., 2000; Hooker et al., 2002; Millar et al., 1999), which is involved in the elongation of fatty acyl CoAs longer than C22.

Unlike the condensing enzymes, the other three enzyme activities of the FAE complex, the β-ketoacyl reductase, β-hydroxyacyl dehydratase and enoyl reductase, are shared by all VLCFA elongase complexes. Thus, these three enzymes have broad substrate specificities and generate a variety of acyl products used to make different classes of lipids (Millar and Kunst, 1997). Because genetic screens in Arabidopsis did not result in isolation of mutants defective in the reductases or the dehydratase, suggesting that these enzymes are essential and/or functionally redundant (Millar and Kunst, 1997), genes encoding the β-ketoacyl reductase and enoyl reductase were cloned by homology to the corresponding sequences from Saccharomyces cerevisiae (Beaudoin et al., 2002; Kohlwein et al., 2001). Two β-ketoacyl reductase (KCR) genes are present in both the A. thaliana and maize (Zea mays) genomes. The maize genes, named GL8A and GL8B (Dietrich et al., 2005; Perera et al., 2003; Xu et al., 2002), are not only expressed in the epidermis, but also in internal tissues. Attempts to generate double mutants by crossing gl8a × gl8b failed because embryos carrying both mutations were not viable. Thus, the KCR has an essential function in plants, most likely in the production of sphingolipids (Dietrich et al., 2005).

An A. thaliana single-copy gene was identified as an enoyl reductase (ECR) candidate. Heterologous expression of the putative plant ECR gene rescued the temperature-sensitive lethality of yeast tsc13-1elo2Δ cells (Gable et al., 2004), demonstrating that it encodes a functional ECR. The A. thaliana ECR gene is ubiquitously expressed, and the protein physically interacts with the Elo2p and Elo3p condensing enzymes when expressed in yeast (Gable et al., 2004). The A. thaliana ECR was shown to be identical to CER10 (Zheng et al., 2005), the protein defective in one of the original A. thaliana eceriferum mutants isolated by Koornneef et al. (1989). These eceriferum (literally ‘not bearing wax’) mutants lack epicuticular wax crystals and therefore have glossy green inflorescence stems that can easily be recognized in visual screens. Biochemical analysis of the cer10 mutant demonstrated that the ECR gene product is involved in the VLCFA elongation that is required for synthesis of all the VLCFA-containing lipids, including cuticular waxes, seed triacylglycerides and sphingolipids (Zheng et al., 2005). Although the plant dehydratase remains unknown, recent identification of the yeast β-hydroxyacyl dehydratase PHS1 (Denic and Weissman, 2007) should permit cloning and characterization of this enzyme from plants.

Compound classes

In addition to variations in the chain-length distributions, cuticular wax mixtures from diverse plants and plant organs also contain various constituent compound classes. These compounds vary in the nature and position of the (typically oxygen-containing) functional groups, with the extreme case of hydrocarbons that are devoid of functional groups (Jetter et al., 2006). Five or more parallel reactions (or pathways), all competing for the VLCFA CoA precursors, can be envisioned leading to these ubiquitous wax components: (i) acyl reduction, (ii) esterification with an alkyl alcohol, (iii) hydrolysis, (iv) aldehyde formation and (v) alkane formation (Figure 2). Knowledge of all the wax biosynthetic reactions will assist in their exploitation for biotechnological production of individual compounds and/or mixtures of compounds with specific combinations of functional groups.

In virtually all vascular plants, wax compound classes with predominantly even numbers of carbons are produced by the so-called acyl reduction pathway (Figure 2; Kunst et al., 2006). The most important of these compounds are primary alcohols and alkyl esters. The latter are essentially dimeric compounds, in which the primary alcohols are bonded to acyl groups, most commonly C16, C18 or VLCFAs (>C20; Figure 3). Primary alcohols are thus central metabolites of wax biosynthesis, and their formation from VLCFA CoA esters has been studied extensively.

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Figure 3.  Array of biosynthetic reactions leading to wax esters. First, the variety of chain lengths is generated by elongation, leading to C22 fatty acid (‘ic’) precursors in seeds (black arrows) and including all chain lengths up to C32 in epidermal cells (orange arrows). Then, individual acyl precursors are reduced to the corresponding alcohols (‘ol’) (green arrows), and alcohols and acyl CoAs of various chain lengths are combined into esters (blue arrows). Depending on the specificity of the elongase (KCS), acyl reductase (FAR) and ester synthase (WS) enzymes, various mixtures of ester isomers and chain lengths can be generated. Arabidopsis stem surface wax contains esters with predominantly C16 acyl and C22–C30 alkyl groups.

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Two reduction steps are required to transform acyl precursors into primary alcohols, and aldehydes must occur as intermediates of the reaction sequence. It has been much debated whether both reduction steps are catalysed by one fatty acyl reductase (FAR), or whether two separate enzymes are necessary for alcohol formation. There is currently substantial evidence for the existence of a one-enzyme system in a number of plant species, including the green alga Euglena gracilis (Kolattukudy, 1970) as well as the angiosperms jojoba (Simmondsia chinensis; Pollard et al., 1979), pea (Pisum sativum; Vioque and Kolattukudy, 1997) and A. thaliana (Rowland et al., 2006). For example, functional expression of genes specifying alcohol-forming FARs from jojoba (Metz et al., 2000) and A. thaliana (Rowland et al., 2006) in heterologous systems demonstrated that alcohol biosynthesis from VLCFAs in these species is carried out by a single alcohol-forming FAR. In contrast, biochemical feeding experiments that allowed isolation of an aldehyde intermediate suggest that the two-step process of alcohol formation operates in Brassica oleracea (Kolattukudy, 1971). However, similar biochemical evidence from other species and molecular information supporting the two-step process in any system is currently lacking.

It is generally assumed that primary alcohols serve as precursors for ester biosynthesis. However, detailed analyses of esterified and free alcohols of various mutants of A. thaliana only recently demonstrated a clear correlation of alcohol chain lengths in both types of compounds, indicating that the free alcohols are indeed incorporated into the wax esters (Lai et al., 2007). In addition, this study revealed that the levels of free alcohols are limiting for ester formation. Thus, a pool of primary alcohols, generated in the A. thaliana epidermal cells, is available either for export towards the cuticle or for esterification with an acyl CoA. Other plant species exhibit large variations in compositions of cuticular wax esters, characterized in some cases by broad distributions of acyl and/or alkyl moieties and in other cases by relatively high preferences for certain isomers (Figure 4).

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Figure 4.  Diversity of acyl and alkyl compositions of wax esters from three plant species. Waxes were extracted from leaf surfaces and analysed by GC-MS (= 3). The relative acyl composition for each ester chain length was determined from the abundances of MS fragments [RCO2H2]+, and used to calculate overall acyl and alkyl distributions across ester chain lengths.

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In higher plants, mammals and bacteria, ester biosynthesis is catalysed by one of three classes of wax synthase (WS) enzymes: jojoba-type WS, mammalian WS, and WS/DGAT bifunctional enzymes. Jojoba-type WS uses a wide range of saturated and unsaturated acyl CoAs ranging from C14 to C24, with 20:1 as the preferred acyl and 18:1 as the preferred alcohol substrate (Lardizabal et al., 2000). In A. thaliana, there are 12 wax synthases with high homology to the jojoba WS, but none have yet been characterized. Mammalian WS enzymes do not have homologues in plants, and have highest activities with C12–C16 acyl CoAs and alcohols shorter than C20 (Cheng and Russell, 2004a,b). A bifunctional WS/DGAT enzyme from Acinetobacter calcoaceticus has a preference for C14 and C16 acyl CoA together with C14–C18 alcohols (Stöveken et al., 2005). Nearly a hundred WS/DGAT homologues have been identified from over 20 other micro-organisms so far (Wältermann et al., 2007), and ten sequences in the A. thaliana genome have also been annotated as WS/DGATs. One of these enzymes, WSD1, has been characterized and shown to be responsible for the formation of cuticular wax esters in A. thaliana stems (R.J., L.K., F. Li, X. Wu and A.L. Samuels, University of British Columbia, Canada, unpublished results). The enzyme utilizes mostly saturated C16 acyl CoA precursors, showing that this upstream precursor of wax production must be co-localized in the cell with the primary alcohols, which are synthesized far downstream in the wax biosynthetic pathway.

Two additional compound classes with predominantly even-numbered chain lengths, aldehydes and free fatty acids, are also found in the wax mixture of most plant species, albeit usually at relatively low concentrations. Currently, our knowledge on their biosynthesis is very limited. Formation of free fatty acids must involve hydrolysis of the elongated acyl CoA precursors (Figure 2). However, it is not clear whether this reaction occurs spontaneously or whether it is enzyme-catalysed. Aldehyde formation requires reduction of acyl CoA precursors, and may occur as an intermediate step during alcohol formation (see above), during alkane formation (see below), or independently of either of these pathways (Figure 2). Only a single wax aldehyde-forming reductase enzyme has been partially purified to date, and the gene encoding this enzyme has not been identified (Vioque and Kolattukudy, 1997).

A separate set of wax biosynthetic reactions is responsible for the formation of compounds with predominantly odd numbers of carbons (Figure 2). Examples of such compound classes include the alkanes, secondary alcohols and ketones that occur together in many wax mixtures and typically share similar chain-length distributions (Jetter et al., 2006). Early biochemical experiments led to a model describing the biosynthesis of these compounds as a two-stage process, with a first set of reactions transforming VLCFA precursors into alkanes and a second series of reactions modifying them into secondary alcohols and ketones (Kolattukudy, 1965). Subsequent experiments confirmed the central role of alkanes in this pathway (Kolattukudy, 1968; Kolattukudy and Brown, 1974; Kolattukudy et al., 1974), either as intermediates en route to mid-chain functionalized compounds or as end products if the downstream reactions are missing. Overall, the second stage of the pathway is relatively well characterized, whereas the first part remains poorly understood.

Although conversion of VLCFA precursors into alkanes could proceed directly in one reaction, the net acyl decarboxylation is apparently brought about by a sequence of transformations. This multistep pathway is supported by the fact that a number of different A. thaliana mutants with alkane-deficient cuticular wax mixtures have been described (Hannoufa et al., 1993; Jenks et al., 1995; Rashotte et al., 2001, 2004). Cloning of several of these mutated genes (CER1, CER2 and CER3/WAX2) revealed that the proteins they encode contain motifs similar to known biosynthetic enzymes (Aarts et al., 1995; Ariizumi et al., 2003; Chen et al., 2003; Kurata et al., 2003; Negruk et al., 1996; Rowland et al., 2007; Xia et al., 1996). While this suggests a potential enzymatic role for these proteins, their exact function remains unknown. Due to this lack of molecular information, it is currently not possible to predict the exact number of reaction steps involved in the conversion of acyl precursors into alkanes, the nature of these steps or the resulting intermediates.

Two alternative pathways have been proposed for the conversion of acyl compounds into alkanes, which vary in the central reaction in which a C1 unit is cleaved off (Bianchi, 1995; Bognar et al., 1984; Chibnall and Piper, 1934). The difference lies in the nature of the immediate precursor from which cleavage occurs and whether the C1 unit is CO or CO2 (decarbonylation versus decarboxylation). Only one model, which describes alkane formation as the decarbonylation of an aldehyde intermediate, has been tested experimentally to some extent (Cheesbrough and Kolattukudy, 1984). However, conclusive molecular genetic and biochemical evidence for either model is lacking, leaving alkane formation as the least understood part of wax biosynthesis.

In A. thaliana leaves, alkanes are the major odd-numbered product, while a high level of secondary alcohols and ketones accompanies alkanes in the stem wax, as well as in wax from B. oleracea leaves (Baker, 1974; Jenks et al., 1995). In these instances, a second stage of the pathway is additionally involved, transforming alkanes first into secondary alcohols and then into ketones (Figure 2). This reaction sequence is well-supported by chemical evidence correlating chain-length and isomer compositions of all three compound classes (Jenks et al., 1995), and by biochemical evidence provided by feeding experiments and detailed studies of label positions in resulting products (Kolattukudy and Liu, 1970; Kolattukudy et al., 1971, 1973). Recently, a reverse genetic approach led to the discovery of a cytochrome P450 enzyme that is involved in secondary alcohol and ketone formation in A. thaliana (Greer et al., 2007). The protein is a mid-chain alkane hydroxylase (MAH1) catalysing two consecutive reactions by first hydroxylating the central CH2 group of alkanes, and then probably re-binding the resulting secondary alcohol for a second hydroxylation of the same carbon. Overall, this confirms the original hypothesis that the pathway involves alkanes as central intermediates that may be further oxidized depending on plant species and organ.

Waxes from certain taxa and/or organs can also contain other compound classes (Figure 1), most prominently aromatic esters and compounds with two hydroxyl or carbonyl functions (diols, ketols, ketoaldehydes and diketones; Jetter et al., 2006). These wax constituents can be regarded as downstream or side products of the ubiquitous biosynthetic reactions forming the common product classes as described above. This implies that additional enzymes, expressed at high levels in certain plant species, can intercept intermediates and/or final products of the ubiquitous pathways before they are exported to the cuticle. As these enzymes can apparently handle the pre-formed wax compounds, they could be added in a modular fashion to the standard pathways in heterologous expression systems. This would allow stepwise addition and modification of secondary functional group(s), and substantially increase the chemical diversity of biotechnologically produced wax mixtures. The necessary biochemical and molecular genetic information on the biosynthesis of these compound classes is currently not available. However, cloning and characterization of the genes involved may become possible in the near future, once the standard wax biosynthetic pathways are better understood in A. thaliana, so that the rapidly growing genomic information from other species (Pennisi, 2007) can be further exploited.

With the isolation and characterization of a number of key genes involved in modification of VLCFA precursors into the diverse wax compound classes, important information on the intracellular localization of wax biosynthetic pathways has emerged. The site of primary alcohol formation appears to be the ER, as shown by localization of the alcohol-forming Arabidopsis enzyme CER4 after expression in yeast (Rowland et al., 2006). This is in contrast with mammalian FARs, which are associated with peroxisomes (Burdett et al., 1991; Cheng and Russell, 2004a,b), and therefore the localization of the CER4 FAR will have to be verified in planta. Meanwhile, the subsequent enzyme in the wax biosynthetic pathway, the wax ester synthase WSD1, has been localized to the ER, (R.J., L.K., F. Li, X. Wu and A.L. Samuels, University of British Columbia, Canada, unpublished results). Similarly, the mid-chain alkane hydroxylase MAH1 (CYP96A15), which catalyses the last two steps of the alkane-pathway in A. thaliana stems, is also confined to the ER (Greer et al., 2007). These two downstream pathway enzymes are thus co-localized with the VLCFA-generating FAEs (Kunst and Samuels, 2003; Xu et al., 2002; Zheng et al., 2005), and it is very likely that the entire wax biosynthesis process occurs in a single subcellular compartment. All the wax biosynthetic enzymes and precursors are therefore expected to be present in the ER of epidermal cells, leading to accumulation of all intermediates and products in the extensive membrane system of this organelle.

Current applications and commercial sources of waxes

  1. Top of page
  2. Summary
  3. Introduction
  4. Plant cuticular wax composition and biosynthesis
  5. Current applications and commercial sources of waxes
  6. Metabolic engineering for wax ester production
  7. Potential for biotechnological alkane production
  8. Conclusions and perspectives
  9. Acknowledgements
  10. References

Wax applications

From the applied perspective, waxes are defined as mixtures of lipophilic compounds that are solid at room temperature, range from transparent to opaque, and are ductile and easy to polish (Illmann et al., 1983; Warth, 1956). Physical parameters used to characterize waxes include hardness, cure speed, melting point or range, pour point, viscosity, (low) surface tension, adhesive strength, optical transparency and durability, and thermal expansion coefficient (Anwar et al., 1999; Imai et al., 2001; Kim and Mahlberg, 1995; Kobayashi et al., 2005; McMillan and Darvell, 2000).

Due to their special properties, waxes are used as lubricants, adhesives, coatings, sealants, impregnation materials and adjuvants in formulations of (bio)active compounds. A wide range of commercially important final products rely on waxes, including automobiles, textiles, papers and specialty inks, pesticides, candles, plastics and wood–plastic composites, furniture and shoe polish, household cleaners, cosmetics, dental treatment products, drugs (lozenge coating) and food (chewing gum, cheese packaging, confectionery coating).

Wax sources

To meet the demand for material applications, waxes are currently generated by chemical syntheses, obtained from geological deposits originating from past organisms (fossil waxes) or obtained from living organisms (recent waxes) (Illmann et al., 1983). The vast majority of these waxes are based either on alkane or ester structures: synthetic waxes are mainly generated by the Fischer–Tropsch process (CO + H2) and olefin (ethylene, propylene) polymerization, giving rise to mixtures of normal and branched alkanes (Illmann et al., 1983; Schulz, 1999; Warth, 1956). Fossil waxes, on the other hand, are extracted from crude oil and coal deposits, yielding alkanes and alkyl ester mixtures (together with the corresponding free acids and alcohols), respectively (Illmann et al., 1983; Warth, 1956).

Beeswax and wool wax are the prime commodities of recent waxes from animal sources (Tulloch, 1971; Warth, 1956), whereas the most important plant sources for commercial production of waxes are carnauba (Copernicia cerifera), candelilla (Euphorbia cerifera and E. antisyphilitica), ouricouri (Syagros coronata), sugar cane (Saccharum sp.) and jojoba (Simmondsia chinensis). All these recent waxes are relatively rich in aliphatic esters, with varying overall chain lengths of both acyl and alkyl groups, and contain characteristic admixtures of cinnamates, hydroxyesters and lactones, steryl esters, estolides and alkanes (Basson and Reynhardt, 1988; Holloway, 1984; Illmann et al., 1983; Lamberton and Redcliffe, 1960; Regert et al., 2005; Vandenburg and Wilder, 1967; Warth, 1956).

Chemical diversity of commercial waxes

The chemical diversity that is currently available is relatively broad for the wax alkanes, which include a wide variety of isomeric branching patterns and chain lengths. For example, alkane isomers with various patterns of methyl branches can be synthesized through polymerization of propylene and/or co-polymerization of propylene and ethylene (Illmann et al., 1983; Warth, 1956). Furthermore, broad mixtures of (synthetic and fossil) alkanes with diverse chain lengths can be distilled into mixtures with desired chain-length ranges and average carbon numbers, or even purified into single chain lengths (Illmann et al., 1983; Warth, 1956).

The chemical nature of wax esters allows a similar diversity of isomers and chain lengths through variation of acyl and/or alcohol carbon numbers (Figure 3). However, this diversity is presently not commercially exploited, because the natural wax sources are characterized by ester mixtures with relatively narrow ranges of chain lengths in their acyl and alcohol moieties, and therefore also of overall ester chain lengths (Illmann et al., 1983). To increase the wax ester diversity and expand the array of ester applications, chemical variations beyond those available in the current sources will have to be explored. Examples of highly desirable modifications in wax ester structures include in-chain and ω-terminal functional groups on the alkyl and/or on the acyl chains, as well as further variations in average chain lengths and chain-length distributions. The target wax esters will also have to accumulate to high levels in waxes of the source plants to make them viable industrial raw materials. These goals can only be accomplished by genetic engineering of wax biosynthetic pathways in oilseed crops as described below.

Because of their special chain-length composition, some of the wax ester mixtures from natural sources have proven to be important commercial commodities. For example, wax esters consisting of 20:1 fatty acid bonded to 20:1 and 22:1 alcohols are known to have outstanding lubrication properties combined with high resistance to hydrolysis and oxidation (Carlsson, 2006). It has been estimated that there is a market for millions of tonnes of these esters in lubrication alone (Carlsson, 2006). However, these special wax commodities can currently be commercially extracted only from jojoba seeds, i.e. their production relies on the agriculture of a single plant species (Carlsson, 2006; Purcell et al., 2000; Yermanos, 1975). Jojoba cultivation is limited by special growth conditions and low yields with respect to time and agricultural area, resulting in the high cost of jojoba oil and its almost exclusive use for high-value products such as cosmetics and specialty lubricants. Genetic engineering of the jojoba-type wax biosynthetic pathway in a conventional oilseed crop would result in a new cost-effective supply of these wax esters and enable their extensive use.

Exploitation of plant cuticular waxes

With the exception of the esters produced in jojoba seeds, all other commercial plant waxes are harvested more or less directly from plant surfaces, where they are deposited by the epidermal cells. Cuticular wax biosynthesis is largely controlled by developmental genetic programs, resulting in fairly constant, specific compositions for each plant species and organ. Plant cuticular waxes are therefore chemically much more diverse than all the other wax sources, and this greater chemical diversity goes hand in hand with the variations in wax physical properties that are desirable for industrial applications. At present, however, the chemical diversity of plant cuticular waxes is not being exploited because waxes are commercially harvested from only a small number of plant species. For example, the carnauba palm (Copernicia cerifera) is grown exclusively for cuticular wax production. Its large leaves are covered by an exceptionally thick layer of wax reaching a coverage of 300–1000 μg cm−2 of plant surface (Tulloch, 1976). This greatly facilitates mechanical wax harvest, but yields only 10–100 kg per hectare (Da Silva et al., 1999; Johnson and Nair, 1985). The vast majority of other plant species have leaf wax coverages in the range of 1–100 μg cm−2 (Jetter et al., 2006), but these low wax amounts can be offset by large surface areas reached in crop fields (Gower et al., 1999). For example, wheat fields are estimated to contain approximately 10–200 kg of wax per hectare (Austin et al., 1986; Bianchi and Corbellini, 1977). However, substantial investments would be necessary to make harvesting this wax source commercially viable. Sugar cane (Saccharum sp.) is the only crop species from which cuticular wax is currently exploited as a side-commodity, as wax is easily accessible by extraction of the filter cakes from sugar production. Approximately 40–240 kg of wax can be produced per hectare of sugar cane, assuming average crop yields of 50 000 kg ha−1, with filter cakes amounting to 4% of the mass and waxes to 2–12% of the filter cake (US patent 3931258; Paturau, 1982; Azzam, 2006; FAOSTAT, 2008).

In addition to the modest wax coverages and the lack of structural diversity in the wax mixtures, wax utilization from plant surfaces is also limited by the poor agronomic properties and special growth conditions of plant species currently used for wax production. In order to make the surface wax a lucrative side-commodity in the future, the wax coverage and composition of temperate plant species would have to be genetically manipulated in a controlled way. This is currently not feasible, however, because neither the regulation of cuticle-forming genes nor the effect of changed wax composition on the critical cuticle functions have been investigated in any detail.

Potential for plant wax production in seeds

The apparent shortcomings of wax production on plant surfaces can be circumvented by genetic engineering approaches using established high-yielding oil crops as a platform. By introducing wax biosynthetic pathways into oilseeds, waxes with optimal chemical compositions for various specialty markets could be produced, including high-value lubricants, cosmetics and pharmaceuticals as well as high-energy fuels. Potential wax yields from oilseed engineering can be estimated based on the current yields for major plant oil commodities. For example, Brassica seed oil yields are in the range of 500–4000 kg ha−1, with typical values for Canadian canola between 1500 and 1800 kg ha−1 (FAOSTAT, 2008). Even though wax yields from engineered oilseed crops will probably be lower than the current oil amounts, this would represent a several-fold increase over current surface wax yields. That this estimated potential is realistic can be seen from comparison with jojoba, the only species known to accumulate wax in its seeds, which yields 75–750 kg of wax per hectare (Botti et al., 1998).

Metabolic engineering for wax ester production

  1. Top of page
  2. Summary
  3. Introduction
  4. Plant cuticular wax composition and biosynthesis
  5. Current applications and commercial sources of waxes
  6. Metabolic engineering for wax ester production
  7. Potential for biotechnological alkane production
  8. Conclusions and perspectives
  9. Acknowledgements
  10. References

Metabolic engineering of high-yielding oilseed species is a rapid, cost-effective and rational approach for mitigating current limitations in the chemical diversity and yield of surface wax crops. Success will require the following individual steps to be accomplished: (i) elucidation of wax biosynthetic pathways, (ii) reconstitution of selected pathways, one at a time, in transgenic systems, (iii) modification of other lipid biosynthetic pathways by up- or down-regulation of certain enzymes, to control flux and to generate appropriate product mixtures, and (iv) integration of additional unique modification steps into pathways. The first step in this process is nearing completion, and the second step is currently being attempted. The remaining steps can be tackled in the near future. The following example illustrates the whole process. (i) Wax ester biosynthesis is relatively well understood, with genes cloned and characterized from at least one plant species for each enzymatic step involved (Lardizabal et al., 2000; Metz et al., 2000; Rowland et al., 2006). As the early FAE steps of this pathway are shared with formation of sphingolipids (Dietrich et al., 2005; Zheng et al., 2005), which are essential membrane components of all cells, it is likely that sufficient quantities of VLCFA precursors for wax ester production will be available in all target species and tissues. (ii) Heterologous overexpression of a fatty acyl reductase (FAR) together with a wax ester synthase (WS) should therefore lead to wax ester formation. (iii) Wax ester formation can be manipulated to enhance flux or generate novel products. For example, downregulation of triacylglycerol biosynthesis competing for fatty acid percursors should increase wax ester production. On the other hand, up-regulation of steroid biosynthesis should increase the levels of steryl alcohols and result in greater production of steryl esters and/or mixtures of wax and steryl esters. (iv) To further increase the wax ester diversity, additional enzymes may be co-expressed that would lead to the hydroxylation, desaturation or other modifications of the hydrocarbon chains of either the acyl or alkyl moieties.

Proof of concept exists that jojoba-type wax esters (C38–C44) can be produced at high levels by engineering of oilseeds (Lardizabal et al., 2000). A recent study concluded that production of wax esters by introduction of a three-enzyme biosynthetic pathway in the crucifer Crambe abyssinica is a viable enterprise for the EU (Carlsson, 2006). This may lead to high volume production of wax esters at substantially reduced cost, and to their use for general automotive lubrication applications, for example, as transmission and hydraulic fluids.

Wax esters with a vast array of compositions of constituent fatty acids and alcohols are present in various plant species (Figures 3 and 4). Unfortunately, it is currently not clear whether the chain-length compositions of esters from various plant species are governed by the chain-length specificities of the enzymes involved and/or by substrate availability. Chemical evidence for A. thaliana stem wax showed that epidermal ester biosynthesis was limited by wax alcohol pools, but the study did not address enzyme specificity (Lai et al., 2007). Biochemical characterization of various wax ester synthases is currently under way. Once the substrate specificities of these enzymes are known, it will be possible to increase the chemical diversity of wax esters through introduction of the desired enzymes in transgenic crops. The various types of wax esters will have unique properties and will serve as substrates for the production of high-value specialty lubricants, cosmetics and pharmaceuticals. Additional wax ester diversity can be generated by engineering of artificial enzymatic steps into pathways that do not normally occur in nature in a single species, to introduce novel functional groups in either the acid or alcohol moieties of the esters.

Examples of novel wax products with broad industrial applications include esters containing acyl and/or alkyl moieties with C=C double bonds, cyclopropane rings and methyl branches. Interestingly, a recent study on wax hydrocarbons in barley spikes showed that all three structural features are biosynthetically related (von Wettstein-Knowles, 2007), implying that a small set of enzymes can convert elongated wax precursors into these compounds. It was hypothesized that a desaturase introduces a double bond with high positional specificity, and then a cyclopropane synthase and/or a methyl transferase generate(s) branched structure(s). Although the exact nature of the involved enzymes remains to be determined, their potential biochemical function makes them very important targets for cloning and future wax engineering studies.

Esters consisting of C20 and C22 alcohols and a hydroxy fatty acid, for example ricinoleic acid (18:1 − OH), are another type of novel wax product. The presence of hydroxy fatty acids disrupts the packing of hydrocarbon chains, thereby reducing the melting temperature of the wax esters and improving their lubrication properties at low temperatures (Carlsson, 2006).

Wax ester fatty acid and alcohol components are also valuable industrial raw materials, and wax esters containing ricinoleic acid would be of particular interest due to high demand for this chemical as an additive to base oils in lubricant formulations, and as a feedstock for the manufacture of nylon, surfactants, paints, cosmetics and biodegradable polymers for medical applications (Ogunniy, 2006). Castor oil, in which nearly 90% of acyl residues are ricinoleic acid, is currently the only commercial source of hydroxy fatty acids (Atsmon, 1989; Hogge et al., 1991). However, castor beans are far from an ideal source, as they require manual harvesting and contain complex allergens together with the potent toxin ricin. Castor is also not a temperate climate crop, making it necessary to import castor oil into many countries from India and China, with irregular supplies and fluctuating prices. Engineering of an existing oilseed crop to replace castor bean as a major source of hydroxy fatty acids is therefore highly desirable. To date, attempts to produce oils rich in ricinoleic acid have not been successful due to a general lack of understanding of the mechanisms involved in channelling this unusual fatty acid from PC, where it is synthesized, to storage triacylglycerols (Jaworski and Cahoon, 2003). In contrast to triacylglycerol synthesis, the enzymology of wax ester production is not as complex, so engineering of crop plants that efficiently incorporate ricinoleic acid into wax esters may be a viable alternative.

To further increase the structural diversity of genetically engineered wax esters, biosynthetic genes from organisms other than plants should be considered. Wax esters occur in a wide variety of organisms (Kolattukudy, 1976), including mammals (Jakobsson et al., 2006; Yen et al., 2005), birds (Dekker et al., 2000; Haribal et al., 2005; Sweeney et al., 2004), fungi (Cooper et al., 2000) and bacteria (Ishige et al., 2002, 2003). Mammalian and bacterial ester formation is fairly well understood, providing gene candidates for ester synthases with various substrate/product chain-length preferences (see description of ester biosynthesis above). Corresponding genes from birds, fungi and other organisms, once they are identified and characterized, will help to further broaden the repertoire of designer wax esters, for example with methyl branched moieties. Similarly, genes involved in the formation of cuticular alkanes of insects (Howard and Blomquist, 2005) should be explored as candidates for engineering hydrocarbons (see below).

The biosynthetic pathways for the production of novel commercial wax esters can be engineered in oilseed crops that are well suited for their synthesis and utilization. At present, two crop species are being considered as platforms for ester production, Crambe abyssinica and Brassica carinata (Carlsson, 2006). These crops are not intended for food production and will grow wherever other Brassica oilseed crops are cultivated. C. abyssinica has an advantage over B. carinata in that it is a high-yielding crop, and does not out-cross with any other agricultural species (Carlsson, 2006). However, despite its inferior seed yield and some out-crossing with other Brassica crops, B. carinata may have preference because it can be easily and efficiently transformed.

Possible bottlenecks for wax production in oilseeds will also have to be addressed for each species, including the low germination rates of transgenic lines and the intracellular autotoxicity of waxes accumulating in seed embryo cells, whereas they would be exported to the plant surface when produced in epidermal cells (Bird et al., 2007; Panikashvili et al., 2007; Pighin et al., 2004). In addition, the physical behaviour of VLCFA derivatives in seeds will have to be tested in these transgenic crops. It has to be noted that jojoba, the only currently available model for seed wax accumulation, has wax esters with relatively short chain lengths and a substantial amount of unsaturated acyl and alcohol moieties. The resulting low melting points make jojoba wax esters liquid at ambient temperatures. In contrast, longer-chain fully saturated esters are solid at room temperature, and it is not clear whether this might affect their accumulation in transgenic seeds.

Potential for biotechnological alkane production

  1. Top of page
  2. Summary
  3. Introduction
  4. Plant cuticular wax composition and biosynthesis
  5. Current applications and commercial sources of waxes
  6. Metabolic engineering for wax ester production
  7. Potential for biotechnological alkane production
  8. Conclusions and perspectives
  9. Acknowledgements
  10. References

Alkanes, the other large group of currently used very-long-chain wax compounds, can be generated by chemical synthesis and extracted from fossil sources with sufficient chemical diversity and at very low cost (Schulz, 1999; Warth, 1956). It is therefore not commercially attractive to produce alkane-rich waxes using biotechnological approaches. Nevertheless, biosynthesis of cuticular alkanes has great potential for application in commodities other than waxes. One important future market for these hydrocarbons is in the fuel sector, where gasoline and diesel are currently provided by crude fossil oil consisting of various hydrocarbons. Much of our transportation system relies on these hydrocarbons, because the highly reduced carbon contained in them has maximum chemical energy. Hydrocarbons can be replaced to some extent by other compounds including ethanol and biodiesel (Doran-Peterson et al., 2008; Dyer et al., 2008; Pauly and Keegstra, 2008), but for some applications (e.g. aircraft), high-energy hydrocarbon fuels will remain essential. Therefore, it is highly desirable to develop renewable hydrocarbon sources.

Production of hydrocarbon-rich biofuels can be accomplished by biotechnological approaches harnessing photosynthetic organisms. To that end, a number of enzymes have to be combined so that plant compounds, most importantly fatty acids, can be utilized and transformed into the desired products. While the enzymes necessary for the hydrocarbon assembly have not been characterized from any organism to date, they are known to be present in the epidermis of higher plants, where they are capable of converting fatty acids into the hydrocarbons that accumulate in the cuticular wax deposited on the plant surface (see above). The cuticular hydrocarbons are synthesized at too low a level for direct industrial use. However, this system serves as an ideal model for studying hydrocarbon formation, and can be exploited for identification and isolation of genes and enzymes for bio-gasoline production by genetic engineering.

The central reaction of the alkane-forming pathway, i.e. the transition step leading from even- to odd-numbered carbon chains, is thought to involve the loss (rather than addition) of one carbon atom from the acyl precursors (see above). However, the biochemical details of this reaction have not been determined. Several genes involved in alkane formation have been cloned (CER1, CER2 and CER3/WAX2), but all attempts to characterize their protein products have failed so far. As multiple steps are probably required for the conversion of fatty acids to hydrocarbons, additional enzymes and corresponding genes might have to be identified to complete the pathway. Once all the genes/enzymes have been established, the knowledge of this pathway can be exploited for elucidation of analogous pathways leading to the formation of short- and medium-chain, as well as branched-chain, hydrocarbons.

Conclusions and perspectives

  1. Top of page
  2. Summary
  3. Introduction
  4. Plant cuticular wax composition and biosynthesis
  5. Current applications and commercial sources of waxes
  6. Metabolic engineering for wax ester production
  7. Potential for biotechnological alkane production
  8. Conclusions and perspectives
  9. Acknowledgements
  10. References

To overcome current limitations in the supply of waxes, to generate wax commodities with new physical and chemical properties that do not normally occur in a single species, and to create entirely new arrays of wax-derived products with desired chain lengths and functional groups for the chemical industry, it will be necessary to harness the wax biosynthetic diversity present in nature and genetically engineer wax biosynthetic pathways capable of making such specialty chemicals. These rationally designed wax biosynthetic pathways will be introduced into the seeds of oil crops dedicated to industrial use to avoid threat to the existing food and feed systems, and will result in production of renewable, high-value waxes that will be able to compete with petroleum-based products, thus reducing our dependency on fossil oils. In addition, a better understanding of cuticular alkane biosynthesis might provide renewable sources for high-energy hydrocarbons and lead to applications that do not rely on the physical properties of waxes.

Acknowledgements

  1. Top of page
  2. Summary
  3. Introduction
  4. Plant cuticular wax composition and biosynthesis
  5. Current applications and commercial sources of waxes
  6. Metabolic engineering for wax ester production
  7. Potential for biotechnological alkane production
  8. Conclusions and perspectives
  9. Acknowledgements
  10. References

This work has been supported by the Natural Sciences and Engineering Research Council (Canada), the Canada Research Chairs Program, and the Canadian Foundation for Innovation.

References

  1. Top of page
  2. Summary
  3. Introduction
  4. Plant cuticular wax composition and biosynthesis
  5. Current applications and commercial sources of waxes
  6. Metabolic engineering for wax ester production
  7. Potential for biotechnological alkane production
  8. Conclusions and perspectives
  9. Acknowledgements
  10. References
  • Aarts, M.G.M., Keijzer, C.J., Stiekema, W.J. and Pereira, A. (1995) Molecular characterization of the CER1 gene of Arabidopsis involved in epicuticular wax biosynthesis and pollen fertility. Plant Cell, 7, 21152127.
  • Andersson, M.X., Goksor, M. and Sandelius, A.S. (2007) Optical manipulation reveals strong attracting forces at membrane contact sites between endoplasmic reticulum and chloroplasts. J. Biol. Chem. 282, 11701174.
  • Anwar, M., Khan, H.U., Nautiyal, S.P., Agrawal, K.M. and Rawat, B.S. (1999) Solubilised waxes and their influence on the flow properties of lube oil base stocks. Pet. Sci. Technol. 17, 491501.
  • Ariizumi, T., Hatakeyama, K., Hinata, K., Sato, S., Kato, T., Tabata, S. and Troiyama, K. (2003) A novel male-sterile mutant of Arabidopsis thaliana, faceless pollen–1, produces pollen with a smooth surface and an acetolysis-sensitive exine. Plant Mol. Biol. 53, 107116.
  • Atsmon, D. (1989) Castor. In Oil Crops of the World (Robbelen, G., Downey, K.R. and Ashri, A., eds). New York: McGraw-Hill, pp. 438447.
  • Austin, R.B., Morgan, C.L. and Ford, M.A. (1986) Dry matter yields and photosynthetic rates of diploid and hexaploid Triticum species. Ann. Bot. 57, 847857.
  • Azzam, A.M. (2006) Separation and analysis of wax from Egyptian sugar cane filter press cake. Fette Seifen Anstrichmittel, 86, 247250.
  • Baker, E.A. (1974) The influence of environment on leaf wax development in Brassica oleracea var. gemmifera. New Phytol. 73, 955966.
  • Bargel, H., Koch, K., Cerman, Z. and Neinhuis, C. (2006) Structure–function relationships of the plant cuticle and cuticular waxes – a smart material? Funct. Plant Biol. 33, 893910.
  • Basson, I. and Reynhardt, E.C. (1988) An investigation of the structures and molecular dynamics of natural waxes: II. carnauba wax. J. Phys. D, 21, 14291433.
  • Bates, P.D., Ohlrogge, J.B. and Pollard, M. (2007) Incorporation of newly synthesized fatty acids into cytosolic glycerolipids in pea leaves occurs via acyl editing. J. Biol. Chem. 282, 3120631216.
  • Beaudoin, F., Gable, K., Sayanova, O., Dunn, O. and Napier, J.A. (2002) A Saccharomyces cerevisiae gene required for heterologous fatty acid elongase activity encodes a microsomal β-keto-reductase. J. Biol. Chem. 277, 1148111488.
  • Bianchi, G. (1995) Plant waxes. In Waxes: Chemistry, Molecular Biology and Functions (Hamilton, R.J., ed.). Dundee, UK: The Oily Press, pp. 175222.
  • Bianchi, G. and Corbellini, M. (1977) Epicuticular wax of Triticum aestivum Demar 4. Phytochemistry, 16, 943945.
  • Bianchi, G., Avato, P. and Salamini, F. (1984) Surface waxes from grain leaves and husks of maize (Zea mays L.). Cereal Chem. 61, 4547.
  • Bird, D., Beisson, F., Brigham, A., Shin, J., Greer, S., Jetter, R., Kunst, L., Wu, X., Yephremov, A. and Samuels, L. (2007) Characterization of Arabidopsis WBC11/ABCG11, an ATP binding cassette (ABC) transporter that is required for cuticular lipid secretion. Plant J. 52, 485498.
  • Bognar, A.L., Paliyath, G., Rogers, L. and Kolattukudy, P.E. (1984) Biosynthesis of alkanes by particulate and solubilized enzyme preparations from pea leaves (Pisum sativum). Arch. Biochem. Biophys. 235, 817.
  • Bonaventure, G., Salas, J.J., Pollard, M. and Ohlrogge, J.B. (2003) Disruption of the FATB gene in Arabidopsis demonstrates an essential role of saturated fatty acids in plant growth. Plant Cell, 15, 10201033.
  • Botti, C., Prat, L., Palzkill, D. and Canaves, L. (1998) Evaluation of jojoba clones grown under water and salinity stresses in Chile. Ind. Crops Prod. 9, 3945.
  • Burdett, K., Larkins, L.K., Das, A.K. and Hajra, A.K. (1991) Peroxisomal localization of acyl-coenzyme A reductase (long chain alcohol forming) in guinea pig intestine mucosal cells. J. Biol. Chem. 266, 1220112206.
  • Carlsson, A.S.(2006) Production of Wax Esters in Crambe. EpoBIO Report. Newbury, UK: CLC Press, http://www.epobio.net/pdfs/0611CrambeWaxEstersReport_c.pdf .
  • Cheesbrough, T.M. and Kolattukudy, P.E. (1984) Alkane biosynthesis by decarbonylation of aldehydes catalyzed by a particulate preparation from Pisum sativum. Proc. Natl Acad. Sci. USA, 81, 66136617.
  • Chen, X., Goodwin, S.M., Boroff, V.L., Liu, X. and Jenks, M.A. (2003) Cloning and characterization of the WAX2 gene of Arabidopsis involved in cuticle membrane and wax production. Plant Cell, 15, 11701185.
  • Cheng, J.B. and Russell, D.W. (2004a) Mammalian wax biosynthesis. I. Identification of two fatty acyl-coenzyme A reductases with different substrate specificities and tissue distributions. J. Biol. Chem. 279, 3778937897.
  • Cheng, J.B. and Russell, D.W. (2004b) Mammalian wax biosynthesis. II. Expression cloning of wax synthase cDNAs encoding a member of the acyltransferase enzyme family. J. Biol. Chem. 279, 3779837807.
  • Chibnall, A.C. and Piper, S.H. (1934) The metabolism of plant and insect waxes. Biochem. J. 28, 22092219.
  • Clough, R., Matthis, A.L., Barnum, S.R. and Jaworski, J.G. (1992) Purification and characterization of 3–ketoacyl-acyl carrier protein synthase from spinach: a condensing enzyme utilizing acetyl-CoA to initiate fatty acid synthesis. J. Biol. Chem. 267, 2099220998.
  • Cooper, L.L.D., Oliver, J.E., De Vilbiss, E.D. and Doss, R.P. (2000) Lipid composition of the extracellular matrix of Botruytis cinerea. Phytochemistry, 53, 293298.
  • Da Silva, J.A., Da Cunha, P.B. and Meunier, I.M.J. (1999) Modelagem da producao cerifera de carnauba Copernicia prunifera (Miller) H.E. Moore, no municipio de campo maior – Piaui. CERNE, 5, 6168, in Portuguese.
  • Dekker, M.H.A., Piersma, T. and Sinninghe Damste, J.S. (2000) Molecular analysis of intact preen waxes of Calidris canutus (Aves: Scolopacidae) by gas chromatography/mass spectrometry. Lipids, 35, 533541.
  • Denic, V. and Weissman, J.S. (2007) A molecular caliper mechanism for determining very long-chain fatty acid length. Cell, 130, 663677.
  • Dietrich, C.R., Perera, M.A.D.N., Yandeau-Nelson, M., Meeley, R.B., Nikolau, B.J. and Schnable, P.S. (2005) Characterization of two GL8 paralogs reveals that the 3–ketoacyl reductase component of fatty acid elongase is essential for maize (Zea mays L.) development. Plant J. 42, 844861.
  • Doran-Peterson, J., Cook, D.M. and Brandon, S.K.(2008) Microbial conversion of sugars from plant biomass to lactic acid or ethanol. Plant J. 54, 582592.
  • Dunn, T.M., Lynch, D.V., Michaelson, L.V. and Napier, J.A.(2004) A post-genomic approach to understanding sphingolipid metabolism in Arabidopsis thaliana. Ann. Bot. 93, 483497.
  • Dyer, J., Stymne, S., Green, A. and Carlsson, A.(2008) High value oils from plants. Plant J. 54, 640654.
  • FAOSTAT (2008) ProdSTAT: Crops. Rome: Food and Agriculture Organization of the United Nations.
  • Fiebig, A., Mayfield, J.A., Miley, N.L., Chau, S., Fischer, R.L. and Preuss, D. (2000) Alterations in CER6, a gene identical to CUT1, differentially affect long-chain lipid content on the surface of pollen and stems. Plant Cell, 12, 20012008.
  • Gable, K., Garton, S., Napier, J.A. and Dunn, T.M. (2004) Functional characterization of the Arabidopsis thaliana orthologue of Tsc13p, the enoyl reductase of the yeast microsomal fatty acid elongating system. J. Exp. Bot. 55, 543545.
  • Giese, B.N. (1975) Effects of light and temperature on the composition of epicuticular wax of barley leaves. Phytochemistry, 14, 921929.
  • Gower, S.T., Kucharik, C.J. and Norman, J.M. (1999) Direct and indirect estimation of leaf area index, fAPAR, and net primary production of terrestrial ecosystems. Remote Sens. Environ. 70, 2951.
  • Greer, S., Wen, M., Bird, D., Wu, X., Samuels, L., Kunst, L. and Jetter, R.(2007) The cytochrome P450 enzyme CYP96A15 is the mid-chain alkane hydroxylase responsible for formation of secondary alcohols and ketones in stem cuticular wax of Arabidopsis thaliana. Plant Physiol. 145, 653667.
  • Hannoufa, A., McNevin, J. and Lemieux, B. (1993) Epicuticular waxes of eceriferum mutants of Arabidopsis thaliana. Phytochemistry, 33, 851855.
  • Haribal, M., Dhondt, A.A., Rosane, D. and Rodriguez, E. (2005) Chemistry of preen gland secretions of passerines: different pathways to same goal? Why? Chemoecology, 15, 251260.
  • Hogge, L.R., Taylor, D.C., Reed, D.W. and Underhill, E.W. (1991) Characterization of castor bean neutral lipids by mass spectrometry/mass spectrometry. J. Am. Oil Chem. Soc. 68, 863868.
  • Holloway, P.J.(1984) Surface lipids of plants and animals. In Handbook of Chromatography, Lipids. Vol. 1 (Mangold, H.K., Zweig, G. and Sherma, J. eds). Boca Raton, FL: CRC Press, pp. 347380.
  • Hooker, T.S., Millar, A.A. and Kunst, L. (2002) Significance of the expression of the CER6 condensing enzyme for cuticular wax production in Arabidopsis. Plant Physiol. 129, 15681580.
  • Howard, R.W. and Blomquist, G.J. (2005) Ecological, behavioral, and biochemical aspects of insect hydrocarbons. Annu. Rev. Entomol. 50, 371393.
  • Illmann, G., Schmidt, H., Brotz, W., Michalczyk, G., Payer, W., Frohning, C.D., Dietsche, W., Hohner, G. and Wildgruber, J. (1983) Wachse. In Ullmanns Lexikon der Technischen Chemie, Vol. 24 . (Bartholome, E., Biekert, E. and Hellmann, H. eds). Weinheim, Germany: Wiley VCH, pp. 149.
  • Imai, T., Nakamura, K. and Shibata, M. (2001) Relationships between the hardness of an oil–wax gel and the surface structure of the wax crystals. Colloids Surf. A, 194, 233237.
  • Ishige, T., Tani, A., Takabe, K., Kawasaki, K., Sakai, Y. and Kato, N. (2002) Wax ester production from n–alkanes by Acinetobacter sp. strain M–1: ultrastructure of cellular inclusions and role of acyl coenzyme A reductase. Appl. Environ. Microbiol. 68, 11921195.
  • Ishige, T., Tani, A., Sakai, Y. and Kato, N. (2003) Wax ester production by bacteria. Curr. Opin. Microbiol. 6, 244250.
  • Jakobsson, A., Westerberg, R. and Jacobsson, A. (2006) Fatty acid elongases in mammals: their regulation and roles in metabolism. Prog. Lipid Res. 45, 237249.
  • Jaworski, J. and Cahoon, E.B. (2003) Industrial oils from transgenic plants. Curr.Opin. Plant Biol. 6, 178184.
  • Jenks, M.A., Tuttle, H.A., Eigenbrode, S.D. and Feldmann, K.A. (1995) Leaf epicuticular waxes of the eceriferum mutants in Arabidopsis. Plant Physiol. 108, 369377.
  • Jetter, R., Kunst, L. and Samuels, A.L.(2006) Composition of plant cuticular waxes. In Biology of the Plant Cuticle (Riederer, M. and Müller, C.eds). Oxford: Blackwell, pp. 145181.
  • Johnson, D.V. and Nair, P.K.R. (1985) Perennial crop-based agroforestry systems in Northeast Brazil. Agrofor. Syst. 2, 281292.
  • Kim, E.-S. and Mahlberg, P.U. (1995) Glandular cuticle formation in Cannabis (Cannabaceae). Am. J. Bot. 82, 12071214.
  • Kobayashi, M., Saitoh, M., Ishida, K. and Yachi, H. (2005) Viscosity properties and molecular structure of lube base oil prepared from Fischer–Tropsch waxes. J. Jpn Pet. Inst. 48, 365372.
  • Kohlwein, S.D., Eder, S., Oh, C.-S., Martin, C.E., Gable, K., Bacikova, D. and Dunn, T. (2001) Tsc13p is required for fatty acid elongation and localizes to a novel structure at the nuclear–vacuolar interface in Saccharomyces cerevisiae. Mol. Cell. Biol. 21, 109125.
  • Kolattukudy, P.E. (1965) Biosynthesis of wax in Brassica oleracea. Biochemistry, 4, 18441855.
  • Kolattukudy, P.E. (1968) Tests whether a head to head condensation mechanism occurs in the biosynthesis of n–hentriacontane, the paraffin of spinach and pea leaves. Plant Physiol. 43, 14661470.
  • Kolattukudy, P.E. (1970) Reduction of fatty acids to alcohols by cell-free preparations of Euglena gracilis. Biochemistry, 9, 10951102.
  • Kolattukudy, P.E. (1971) Enzymatic synthesis of fatty alcohols in Brassica oleracea. Arch. Biochem. Biophys. 142, 701709.
  • Kolattukudy, P.E. (1976) Chemistry and Biochemistry of Natural Waxes. Amsterdam: Elsevier.
  • Kolattukudy, P.E. and Brown, L. (1974) Inhibition of cuticular lipid biosynthesis in Pisum sativum by thiocarbamates. Plant Physiol. 53, 903906.
  • Kolattukudy, P.E. and Liu, T.-Y.J. (1970) Direct evidence for biosynthetic relationships among hydrocarbons, secondary alcohols and ketones in Brassica oleracea. Biochem. Biophys. Res. Commun. 41, 13691374.
  • Kolattukudy, P.E., Jaeger, R.H. and Robinson, R. (1971) Biogenesis of nonacosan-15–one in Brassica oleracea. Phytochemistry, 10, 30473051.
  • Kolattukudy, P.E., Buckner, J.S. and Liu, T.-Y.J. (1973) Biosynthesis of secondary alcohols and ketones from alkanes. Arch. Biochem. Biophys. 156, 613620.
  • Kolattukudy, P.E., Croteau, R. and Brown, L. (1974) Structure and biosynthesis of cuticular lipids. Hydroxylation of palmitic acid and decarboxylation of C28, C30 and C32 acids in Vicia faba flowers. Plant Physiol. 54, 670677.
  • Koo, A.J.K., Ohlrogge, J.B. and Pollard, M.(2004) On the export of fatty acids from the chloroplast. J. Biol. Chem. 279, 1610116110.
  • Koornneef, M., Hanhart, C.J. and Thiel, F. (1989) A genetic and phenotypic description of eceriferum (cer) mutants in Arabidopsis thaliana. J. Hered. 80, 118122.
  • Kunst, L. and Samuels, A.L. (2003) Biosynthesis and secretion of plant cuticular wax. Prog. Lipid Res. 42, 5180.
  • Kunst, L., Jetter, R. and Samuels, A.L.(2006) Biosynthesis and transport of plant cuticular waxes. In Biology of the Plant Cuticle (Riederer, M. and Müller, C.eds). Oxford: Blackwell, pp. 182215.
  • Kurata, T., Kawabata-Awai, C., Sakuradani, E., Shimizu, S., Okada, K. and Wada, T. (2003) The YORE–YORE gene regulates multiple aspects of epidermal cell differentiation in Arabidopsis. Plant J. 36, 5566.
  • Lai, C., Kunst, L. and Jetter, R. (2007) Composition of alkyl esters in the cuticular wax on inflorescence stems of Arabidopsis thaliana cer mutants. Plant J. 50, 189196.
  • Lamberton, J.A. and Redcliffe, A.H. (1960) The chemistry of sugar-cane wax. I. The nature of sugar-cane wax. Aust. J. Chem. 13, 261268.
  • Lardizabal, K.D., Metz, J.G., Sakamoto, T., Hutton, W.C., Pollard, M.R. and Lassner, M.W. (2000) Purification of a jojoba embryo wax synthase, cloning of its cDNA, and production of high levels of wax in seeds of transgenic Arabidopsis. Plant Physiol. 122, 645655.
  • Lassner, M.W., Lardizabal, K. and Metz, J.G. (1996) A jojoba β-ketoacyl-CoA synthase cDNA complements the canola fatty acid elongation mutation in transgenic plants. Plant Cell, 8, 281292.
  • McMillan, L.C. and Darvell, B.W. (2000) Rheology of dental waxes. Dent. Mater. 16, 337350.
  • Metz, J.G., Pollard, M.R., Anderson, L., Hayes, T.R. and Lassner, M.W. (2000) Purification of a jojoba embryo fatty acyl-coenzyme A reductase and expression of its cDNA in high erucic acid rapeseed. Plant Physiol. 122, 635644.
  • Millar, A.A. and Kunst, L. (1997) Very-long-chain fatty acid biosynthesis is controlled through the expression and specificity of the condensing enzyme. Plant J. 12, 121131.
  • Millar, A.A., Clemens, S., Zachgo, S., Giblin, E.M., Taylor, D.C. and Kunst, L. (1999) CUT1, an Arabidopsis gene required for cuticular wax biosynthesis and pollen fertility, encodes a very-long-chain fatty acid condensing enzyme. Plant Cell, 11, 825838.
  • Negruk, V., Yang, P., Subramanian, M., McNevin, J.P. and Lemieux, B. (1996) Molecular cloning and characterization of the CER2 gene of Arabidopsis thaliana. Plant J. 9, 137145.
  • Ogunniy, D.S. (2006) Castor oil: a vital industrial raw material. Bioresour. Technol. 97, 10861091.
  • Ohlrogge, J. and Browse, J. (1995) Lipid biosynthesis. Plant Cell, 7, 957970.
  • Ohlrogge, J.B., Jaworski, J.G. and Post-Beittenmiller, D. (1993) De novo fatty acid biosynthesis. In Lipid Metabolism in Plants (Moore, T.S., ed.). Boca Raton, FL: CRC Press, pp. 332.
  • Panikashvili, D., Savaldi-Goldstein, S., Mandel, T., Yifhar, T., Franke, R.B., Höfer, R., Schreiber, L., Chory, J. and Aharoni, A. (2007) The Arabidopsis DESPERADO/AtWBC11 transporter is required for cutin and wax secretion. Plant Physiol. 145, 13451360.
  • Paturau, J.M. (1982) By-Products of the Cane Sugar Industry. Amsterdam: Elsevier.
  • Pauly, M. and Keegstra, K.(2008) Cell wall carbohydrates and their modifications as a resource for biofuels. Plant J. 54, 559568.
  • Pennisi, E. (2007) The greening of plant genomics. Science, 317, 317.
  • Perera, M.A.D.N., Dietrich, C.R., Meeley, R., Schnable, P.S. and Nikolau, B.J. (2003) Dissecting the maize epicuticular wax biosynthetic pathway via the characterization of an extensive collection of glossy mutants. In Advanced Research on Lipids (Murata, N., Yamada, M., Nishida, I., Okuyama, H., Sekiya, J. and Hajime, W., eds). Dordrecht, The Netherlands: Kluwer Academic, pp. 225228.
  • Pighin, J.A., Zheng, H., Balakshin, L.J., Goodman, I.P., Western, T.L., Jetter, R., Kunst, L. and Samuels, A.L. (2004) Plant cuticular lipid export requires an ABC transporter. Science, 306, 702704.
  • Pollard, M.R., McKeon, T., Gupta, L.M. and Stumpf, P.K. (1979) Studies on biosynthesis of waxes by developing Jojoba seed. 2. The demonstration of wax biosynthesis by cell-free homogenates. Lipids, 14, 651662.
  • Purcell, H.C., Abbott, T.P., Holser, R.A. and Phillips, B.S. (2000) Simmondsin and wax ester levels in 100 high-yielding jojoba clones. Ind. Crops Prod. 12, 151157.
  • Rashotte, A.M., Jenks, M.A. and Feldmann, K.A. (2001) Cuticular waxes on eceriferum mutants of Arabidopsis thaliana. Phytochemistry, 57, 115123.
  • Rashotte, A.M., Jenks, M.A., Ross, A.S. and Feldmann, K.A. (2004) Novel eceriferum mutants in Arabidopsis thaliana. Planta, 219, 513.
  • Regert, M., Langlois, J. and Colinart, S. (2005) Characterisation of wax works of art by gas chromatographic procedures. J. Chromatogr. A, 1091, 124136.
  • Riederer, M. and Müller, C. (2006) Biology of the Plant Cuticle. Oxford: Blackwell.
  • Rowland, O., Zheng, H., Hepworth, S.R., Lam, P., Jetter, R. and Kunst, L. (2006) CER4 encodes an alcohol-forming fatty acyl-CoA reductase involved in cuticular wax production in Arabidopsis. Plant Physiol. 142, 866877.
  • Rowland, O., Lee, R., Franke, R., Schreiber, L. and Kunst, L. (2007) The CER3 wax biosynthetic gene from Arabidopsis thaliana is allelic to WAX2/YRE/FLP1. FEBS Lett. 581, 35383544.
  • Schnurr, J.A., Shockey, J.M., De Boer, G.-J. and Browse, J.A. (2002) Fatty acid export from the chloroplast. Molecular characterization of a major plastidial acyl-coenzyme A synthetase from Arabidopsis. Plant Physiol. 129, 17001709.
  • Schulz, H. (1999) Short history and present trends of Fischer–Tropsch synthesis. Appl. Catal. A, 186, 312.
  • Shimakata, T. and Stumpf, P.K. (1982) Purification and characterizations of β-ketoacyl-[acyl-carrier-protein] reductase, β-hydroxyacyl-[acyl-carrier-protein] dehydrase, and enoyl-[acyl-carrier-protein] reductase from Spinacia oleracea leaves. Arch. Biochem. Biophys. 218, 7791.
  • Shockey, J.M., Fulda, M.S. and Browse, J.A. (2002) Arabidopsis contains nine long-chain acyl-coenzyme A synthetase genes that participate in fatty acid and glycerolipid metabolism. Plant Physiol. 129, 17101722.
  • Stöveken, T., Kalscheuer, R., Malkus, U., Reichelt, R. and Steinbüchel, A. (2005) The wax ester synthase/acyl coenzyme A:diacylglycerol acyltransferase from Acinetobacter sp. strain ADP1: characterization of a novel type of acyltransferase. J. Bacteriol. 187, 13691376.
  • Stumpf, P.K. (1984) Fatty acid biosynthesis in higher plants. In Fatty Acid Metabolism and its Regulation (Numa, S., ed.). Amsterdam: Elsevier, pp. 155179.
  • Sweeney, R.J., Lovette, I.J. and Harvey, E.L. (2004) Evolutionary variation in feather waxes of passerine birds. Auk, 121, 435445.
  • Tulloch, A.P. (1971) Beeswax: structure of the esters and their component hydroxy acids and diols. Chem. Phys. Lipids, 6, 235265.
  • Tulloch, A.P. (1976) Chemistry of waxes of higher plants. In Chemistry and Biochemistry of Natural Waxes (Kolattukudy, P.E., ed.). Amsterdam: Elsevier, pp. 235287.
  • Vandenburg, L.E. and Wilder, E.A. (1967) Aromatic acids of carnauba wax. J. Am. Oil Chem. Soc. 44, 659662.
  • Vioque, J. and Kolattukudy, P.E. (1997) Resolution and purification of an aldehyde-generating and an alcohol-generating fatty acyl-CoA reductase from pea leaves (Pisum sativum L.). Arch. Biochem. Biophys. 340, 6472.
  • Voelker, T.A.(1996) Plant acyl-ACP thioesterases: chain-length determining enzymes in plant fatty acid biosynthesis. In Genetic Engineering, Vol. 18 (Setlow, J.K.ed.). New York: Plenum Press, pp. 111131.
  • Wältermann, M., Stöveken, T. and Steinbüchel, A. (2007) Key enzymes for biosynthesis of neutral lipid storage compounds in prokaryotes: properties, function and occurrence of wax ester synthases/acyl-CoA:diacylglycerol acyltransferases. Biochemie, 89, 230242.
  • Warth, A.H.(1956) The Chemistry and Technology of Waxes. New York: Reinhold Publishers.
  • Von Wettstein-Knowles, P.(1982) Biosynthesis of epicuticular lipids as analysed with the aid of gene mutations in barley. In Biochemistry and Metabolism of Plant Lipids (Wintermans, J.F.G.M. and Kuiper, P.J.C.eds). Amsterdam: Elsevier, pp. 6978.
  • Von Wettstein-Knowles, P. (1993) Waxes, cutin, and suberin. In Lipid Metabolism in Plants (Moore, T.S., ed.). Boca Raton, FL: CRC Press, pp. 128166.
  • Von Wettstein-Knowles, P. (2007) Analyses of barley spike mutant waxes identify alkenes, cyclopropanes and internally branched alkanes with dominating isomers at carbon 9. Plant J. 49, 250264.
  • Xia, Y., Nikolau, B.J. and Schnable, P.S. (1996) Cloning and characterization of CER2, an Arabidopsis gene that affects cuticular wax accumulation. Plant Cell, 8, 12911304.
  • Xu, X., Dietrich, C.R., Lessire, R., Nikolau, B.J. and Schnable, P.S. (2002) The endoplasmic reticulum-associated maize GL8 protein is a component of the acyl-coenzyme A elongase involved in the production of cuticular waxes. Plant Physiol. 128, 924934.
  • Yen, C.-L.E., Brown, C.H., IV, Monetti, M. and Farese, R.V., Jr (2005) A human skin multifunctional O–acyltransferase that catalyzes the synthesis of acylglycerols, waxes, and retinyl esters. J. Lipid Res. 46, 23882397.
  • Yermanos, D.M. (1975) Composition of jojoba seed during development. J. Am. Oil Chem. Soc. 52, 115117.
  • Zheng, H., Rowland, O. and Kunst, L. (2005) Disruptions of the Arabidopsis enoyl-CoA reductase gene reveal an essential role for very-long-chain fatty acid synthesis in cell expansion during plant morphogenesis. Plant Cell, 17, 14671481.