Fatty acid and triacylglycerol synthesis
Rational and precise engineering of industrial oil and wax production in plants will need to be predicated on a comprehensive understanding of the metabolic pathways for the biosynthesis of fatty acids and their accumulation into TAG and WE storage lipids, and of the genetic control of these pathways. Based on biochemical research over several decades, seed oil biosynthesis had, until relatively recently, come to be viewed as an essentially linear process comprised of sequential fatty acid synthesis, modification and accumulation processes. These pathways and their component enzymes have been described in significant detail in various previous reviews (Baud and Lepiniec, 2010; Li-Beisson et al., 2010; Napier and Graham, 2010; Ohlrogge and Chapman, 2011; Snyder et al., 2009; Wallis and Browse, 2010; Weselake et al., 2009).
In essence, the classical view has been that saturated or monounsaturated acyl chains of various lengths generated via the fatty acid synthesis pathway in the plastid (Figure 2) are exported to the cytosolic compartment from where they are sequentially assembled onto the sn-1, sn-2 and sn-3 positions of the glycerol backbone by the reactions of the Kennedy pathway (Figure 3) to form TAGs that are then sequestered into oil bodies (oleosomes) to serve as energy reserves to support eventual seed germination. Prior to their incorporation into TAG, the acyl chains exported from the plastid are able to undergo a range of enzyme-mediated modifications (Figure 2). The most common of these are the further sequential desaturation at Δ12 and Δ15 positions to produce the polyunsaturated fatty acids (PUFAs) linoleic and α-linolenic. However, in many plants (and other organisms), unusual enzymatic modifications can occur that result in diverse fatty acid structures of potential industrial interest (such as shown in Figure 1). A notable example is the divergent family of Fad2-encoded enzymes that have evolved differing catalytic functions from the ancestral Δ12-desaturase (Figure 4). These catalyse the synthesis of a range of hydroxy, epoxy, acetylenic and conjugated fatty acid structures (Broun et al., 1998a; Cahoon et al., 1999; Dyer et al., 2002; Lee et al., 1998; van der Loo et al., 1995; Nam and Kappock, 2007; Qiu et al., 2001; Sperling et al., 2000). Enzymes for the conversion of oleic acid to cyclopropanoic acid fatty acids have also been identified, for example in Sterculia (Bao et al., 2002) and cottonseed (Yu et al., 2011). Also, there are fatty acid elongase (FAE) systems that operate in the cytosol for the elongation of oleic acid to eicosenoic acid (C20:1Δ11) and erucic acid.
Figure 3. Contemporary diagrammatic representation of the known metabolic routes by which acyl groups from the acyl-PC and acyl-CoA pools can be directly or indirectly channelled to triacylglycerol (TAG). The traditional linear Kennedy pathway is shown in green and more recently defined routes in blue. Acyl pools are shown as solid rectangular boxes, and principal enzymes involved are shown as open oval boxes. ACS, acyl-CoA synthase; DGAT, diacylglycerol acyltransferase; GPAT, glycerol-3-phosphate acyltransferase; LPAAT, lysophosphatidic acid acyltransferase; LPCAT, lysophosphatidylcholine acyltransferase; PAP, 3-sn-phosphatidate phosphohydrolase; PDAT, phosphatidylcholine diacylglycerol acyltransferase; PDCT, phosphatidylcholine diacylglycerol cholinephosphotransferase; PLA, phospholipase A; PLC, phospholipase C; PLD, phospholipase D.
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Figure 4. Examples of unusual fatty acid structures originating from modifications to the structure of oleic acid (C18:1) in the acyl-PC pool from the action of divergent Fad2 enzymes and CPFA synthase. RA, ricinoleic acid; ESA, eleostearic acid; VA, vernolic acid; CA, crepenynic acid; DHSA, dihydrosterculic acid; C18:2, linoleic acid.
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In recent years, however, the discovery and characterization of several additional enzymes involved in acyl exchange within the cytosolic compartment has challenged this essentially linear view of TAG assembly. Newly identified acyl exchange enzymes appear to be playing significant roles in the transfer of fatty acids from the site of modification on phosphatidylcholine (PC) to deposition on TAG molecules. They may also be providing specialized pathways for the channelling of unusual, and potentially damaging, acyl groups away from membranes and into storage lipids and thus have potentially significant consequences for attempts to metabolically engineer unusual fatty acid production in plant oils. In particular, a number of enzymes have now been uncovered that mediate the transfer of acyl groups directly from PC to TAG without passage through the acyl-coenzyme A (CoA) pool. One such route involves the enzyme phosphatidylcholine diacylglycerol acyltransferase (PDAT) that transfers an acyl group from the sn-2 position of PC directly to the sn-3 position of TAG in seed of Arabidopsis (Ståhl et al., 2004). It has recently been shown that PDAT and diacylglycerol acyltransferase (DGAT) are the two main enzymes responsible for TAG synthesis in Arabidopsis seeds (Zhang et al., 2009). Mutation in one gene and down-regulation by RNA interference (RNAi) of the other led to seeds virtually lacking TAG, whereas double mutants were both pollen and embryo lethal (Zhang et al., 2009). Similar to the situation with DGAT, PDAT has been shown to be expressed both in tissues that accumulate TAG (oil palm mesocarp) and in similar tissues that are essentially devoid of TAG (date palm mesocarp) (Bourgis et al., 2011). The relative contribution of DGAT and PDAT in seed TAG synthesis can vary drastically between even evolutionary closely related species, like sunflower and safflower (W. Banas, A. Sanchez, A. Banas and S. Stymne, pers. commun.).
An alternative route, by which acyl groups present on PC can be moved directly to the TAG backbone, is through exchange of the polar head group between PC and diacylglycerol (DAG), the immediate precursor of TAG. The enzyme responsible, phosphatidylcholine diacylglycerol cholinephosphotransferase (PDCT), was recently described in Arabidopsis and shown to act by transferring the complete phosphocholine head group to directly form DAG, thereby effectively providing the entire glycerol backbone from PC as a precursor of TAG (Lu et al., 2009). Mutational inactivation of this gene results in the lowering of oil content and polyunsaturation because of reduced flux of acyl groups from PC to DAG. Furthermore, when the PDCT mutation was combined with mutations in the lysophosphatidylcholine acyltransferase genes (LPCAT1 and LPCAT2) that are responsible for acyl loading and editing of PC, there was a dramatic reduction in PUFA content in the seed TAG, down to one-third the wild-type level. These results indicate that PC acyl editing and phosphocholine head-group exchange between PC and DAG control the majority of acyl fluxes through PC to provide PUFA for TAG synthesis (Bates et al., 2012). Recent work also demonstrates that the acyl-CoA-independent transfer of acyl groups from PC to TAG catalysed by PDAT is dependent on an efficient acylation of lysophosphatidylcholine, the coproduct of the PDAT reaction. In the absence of a functional DGAT1, the Arabidopsis seed TAG is mainly synthesized by PDAT and disruption of LPCAT2 activity caused drastic reduction in seed TAG content and severely disturbed seed development (Xu et al., 2012).
Another related acyl-CoA-independent route to DAG involves phospholipase D (PLD) that catalyses the removal of just the choline group from PC to form phosphatidic acid (PA), which can then be dephosphorylated by 3-sn-phosphatidate phosphohydrolase (PAP) to form DAG. Suppression of PLD in developing soybean seeds has recently been shown to alter both the total content and the level of unsaturation of soybean seed oil, suggesting that it may be playing a role in the conversion of PC to TAG during oil synthesis, either by a direct catalytic action to produce phosphatidic acid (PA), or by exerting a regulatory influence over other acyl exchange enzymes (Lee et al., 2011). In both of these head-group removal reactions (PDCT and PLD), the acyl groups at position sn-1 and sn-2 of PC retain their positioning on the TAG backbone.
Taken overall, the relatively recent uncovering of PDAT and PDCT has now provided clear evidence for the existence of metabolic routes for direct (acyl-CoA independent) transfer of acyl groups from PC to each of the three positions on TAG during seed oil biosynthesis. These pathways presumably operate in addition to the classical Kennedy pathway. However, it is tempting to speculate that the full picture is still not elucidated, especially given the significant numbers of uncharacterized acyltransferase genes evident in recent and current genomic and transcriptomic studies of several oil-bearing plants that accumulate unusual fatty acids, such as flax (Wang et al., 2012), Hiptage benghaliensis and Bernardia pulchella (A.G. Green, S. Singh and X.-R. Zhou, unpublished data). Many of these gene families could harbour additional enzymes for direct acyl exchange either between PC and the various TAG intermediates, or amongst the intermediates themselves. For example, there have been biochemical indications of possible DAG/DAG acyl exchange yielding a monoacylglycerol (MAG) and a TAG (Stobart et al., 1997). However, no gene dedicated to such a function has yet been identified, although it has been reported that a soluble form of yeast PDAT enzyme has some low level of DAG/DAG transacylation activity (Ghosal et al., 2007). Recent work could not detect any DAG/DAG transacylase activity in sunflower or safflower membranes, despite good PDAT activity (W. Banas, A. Sanchez, A. Banas and S. Stymne, pers. commun.) casting some doubts over any significant contribution of such enzyme activity in TAG synthesis in oil seeds. Interestingly, PDAT can also transfer acyl groups from sn-1 of PC to TAG, albeit at a lower rate (about 30%) than with acyl groups from the sn-2 position (W. Banas, A. Sanchez, A. Banas and S. Stymne, pers. commun.; Ståhl et al., 2004). As acyl groups at sn-1 position of PC can undergo desaturation and structural modification at that position (Bao et al., 2003; Stymne et al., 1992), it is possible that PDAT and other enzymes might also be involved in channelling these acyl groups from PC to TAG.
Thus, the picture of TAG synthesis that is now emerging is one of a complex network of acyltransferase reactions mediating the movement of acyl groups between pools of acyl-PC, acyl-CoA and TAG precursors within the cytosolic compartment (Figure 3). Acyl moieties exported from the plastid could take various potential metabolic routes through these networks towards assembly on TAG, the routes taken probably differing between plant species, based on presence, activity and coordinated regulatory control of alternative enzymatic steps. For example, recent studies with soybean (Bates and Browse, 2011) and Arabidopsis (Bates and Browse, 2012) have identified the presence of kinetically distinct DAG pools during oil accumulation. Diacylglycerol (DAG) used in TAG assembly was predominantly derived from PC, whereas DAG produced through de novo synthesis (Kennedy pathway) was mainly used to produce PC. As fatty acids esterified to PC are substrates for modifications, such as desaturation, hydroxylation or epoxygenation, the fatty acid composition of the PC-derived DAG pool may be markedly different to that of the de novo DAG pool. The extent of this shunting of DAG through PC, compared with direct conversion to TAG, may vary significantly between different species and could be a significant underlying factor in the varying ability of some wild plants to incorporate unusual fatty acids into TAG, as well as the difficulty in engineering some oilseeds to accumulate unusual fatty acids.
Importantly, particular routes through this metabolic network may have become specialized for particular unusual acyl groups in certain plants. This feature could be based on customized substrate specificity of the acyltransferases available, and there is already evidence of divergent substrate specificities within a number of acyltransferase families. For example, it is known that DGAT enzymes in castor bean, Vernonia galamensis and tung have strong preferences for DAG and CoA substrates containing ricinoleoyl, vernoleoyl and eleostearoyl moieties, respectively (Kroon et al., 2006; Shockey et al., 2006; Yu et al., 2006).
Wax ester synthesis
In contrast to the complex, and perhaps incompletely defined, metabolic pathways leading to TAG synthesis, the pathways resulting in WE are relatively simple and well understood (Lardizabal et al., 2000). In essence, synthesis of a WE is a two-step process involving, firstly, the conversion of a fatty acid to a fatty alcohol by the action of a fatty acid reductase (FAR), followed by the esterification of the fatty alcohol to a fatty acid by the action of a wax synthase (WS). This final step in WE synthesis can be viewed as being analogous to the final DGAT-mediated step in the formation of TAG, in that the fatty acid donor becomes esterified either through the primary OH group of the fatty alcohol or the through sn-3 OH group of DAG. This commonality has been reinforced by the discovery of DGAT enzymes with dual-function WS/DGAT activity (Kalscheuer and Steinbüchel, 2003; Li et al., 2008) and able to utilize a variety of acyl acceptors as substrates (Yen et al., 2005). Arguably, the best-studied metabolic pathway for WE synthesis is that occurring naturally in jojoba seed wax and its transgenic expression in oil-forming seeds (Lardizabal et al., 2000). Jojoba WE synthesis is located in the extra-plastidial compartments, and both FAR and WS are membrane associated and utilize acyl-CoA (Lardizabal et al., 2000; Metz et al., 2000). The structure of jojoba WE is dominated by C20:1 and C22:1 acyl and alcohol chains, and this is reflected in the substrate specificity of both the FAR and the WS enzymes (Lardizabal et al., 2000; Metz et al., 2000).