- Email: [email protected]
Acetyl-CoA—Life at the metabolic nexus
Plant Science 176 (2009) 597–601
Contents lists available at ScienceDirect
Plant Science journal homepage: www.elsevier.com/locate/plantsci
Review
Acetyl-CoA—Life at the metabolic nexus David J. Oliver a,*, Basil J. Nikolau b, Eve Syrkin Wurtele a a b
Department of Genetics, Development and Cell Biology, Iowa State University, Ames, IA 50011, USA Department of Biochemistry, Biophysics and Molecular Biology, Iowa State University, Ames, IA 50011, USA
A R T I C L E I N F O
A B S T R A C T
Article history: Received 27 October 2008 Received in revised form 10 February 2009 Accepted 10 February 2009 Available online 20 February 2009
As a key intermediate in a number of different metabolic pathways, acetyl-CoA is independently produced and consumed in the plastids, mitochondria, peroxisomes and cytosol. Recent studies have led to new understanding of the enzymes that generate these different acetyl-CoA pools, including the role of ATP-citrate lyase in producing acetyl-CoA in the cytosol and of the pyruvate dehydrogenase complex for making it in the plastids. The involvement of a second plastid enzyme, acetyl-CoA synthetase, is more speculative but appears to be important in protecting plants from the two-carbon intermediates of fermentation by allowing the conversion of ethanol and acetaldehyde to fatty acids. New roles have also recently been proposed for triacylglycerols in energy storage and oil production in leaves. While additional biological functions for acetyl-CoA are being discovered there are also new proposals for using the chemistry of acetyl-CoA and fatty acid synthesis as the foundation for a new biologically based source of commodity chemicals. ß 2009 Elsevier Ireland Ltd. All rights reserved.
Keywords: Anaerobic metabolism Fatty acids Metabolic engineering Triacylglycerols
Contents 1. 2.
3.
4.
Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Recent breakthroughs in acetyl-CoA metabolism. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. Acetyl-CoA in the plastid . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. Acetyl-CoA in the cytosol. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3. Acetyl-CoA in the glyoxysomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4. Acetyl-CoA in the mitochondria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Some unanswered questions about acetyl-CoA metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1. The physiological function of acetate metabolism in plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2. Triacylglycerols in leaves . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Metabolic engineering of acetyl-CoA metabolism for production of biorenewable chemicals. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
1. Introduction Acetyl-CoA is a metabolite at a key point connecting catabolic and anabolic metabolism. It is produced by the catabolism of carbohydrates, lipids, and amino acids. At the same time it is the precursor for the synthesis of oils, membrane lipids, cuticle, suberin, isoprenoids, flavonoids, stilbenoids, some amino acids, and sometimes sugars. Because of the unique central metabolic
* Corresponding author. Tel.: +1 515 294 4118; fax: +1 515 294 1303. E-mail address: [email protected] (D.J. Oliver). 0168-9452/$ – see front matter ß 2009 Elsevier Ireland Ltd. All rights reserved. doi:10.1016/j.plantsci.2009.02.005
597 598 598 598 599 599 599 599 600 600 600 601
position occupied by acetyl-CoA, flux through this intermediate ultimately controls the rate that carbon reserves in plants can be converted into many important chemicals. Plans for engineering plants for optimal production of proteins, lipids, and specialty chemicals must consider the systems that produce acetyl-CoA as a substrate for these products. It is surprising therefore that so many questions about the basic biochemistry of acetyl-CoA production and use in plant cells remain unanswered. The purpose of this review is threefold. First, we provide a brief overview of recent breakthroughs in the understanding of acetylCoA metabolism in plants. Second, we discuss some unanswered questions concerning acetate metabolism. Finally, we describe a
598
D.J. Oliver et al. / Plant Science 176 (2009) 597–601
vision of how to change acetyl-CoA metabolic pathways to turn plants into engines for producing commodity chemicals. 2. Recent breakthroughs in acetyl-CoA metabolism Acetyl-CoA metabolism occurs in many subcellular compartments in a plant cell. The chloroplasts, peroxisomes, mitochondria, cytosol, and nucleus are shown in Fig. 1 [1,2]. The origins and metabolic fates of acetyl-CoA are distinct in each subcellular compartment, with the plastid and cytosol acetyl-CoA pools being used primarily in anabolic processes, whereas the mitochondrial and peroxisomal pools are used in both anabolic and catabolic processes. Acetyl-CoA is produced in plastids from pyruvate and acetate and supplies precursors for the biosynthesis of fatty acids, amino acids, and glucosinolates. Cytosolic acetyl-CoA is largely produced from citrate and provides carbon for elongation of fatty acids, which are converted to a variety of chemicals and is condensed yielding other polyketides. Cytosolic acetyl-CoA has
also been implicated in the synthesis of secondary metabolites including flavonoids, stilbenoids, and isoprenoids. Peroxisomal acetyl-CoA is formed from fatty acid breakdown and is the substrate for the glyoxylate shunt. In mitochondria acetyl-CoA is made from pyruvate oxidation and is the substrate for the TCA cycle and in some cases fatty acid biosynthesis. Because acetyl-CoA cannot readily cross membranes, the subcellular organization can be rationalized on the basis of providing plants unique regulatory mechanisms to control acetyl-CoA metabolism for different metabolic purposes. 2.1. Acetyl-CoA in the plastid Plastidic acetyl-CoA is synthesized by two enzyme systems, the pyruvate dehydrogenase complex (PDHC) and acetyl-CoA synthetase (ACS). ACS was discovered first [3] and for years it was considered a primary source of acetyl-CoA for lipid synthesis, although the source of its acetate substrate was never clear [4,5]. The discovery of PDHC provided a more direct connection between acetyl-CoA formation and central metabolism [6], still there was little indication as to which enzyme was the primary source of acetyl-CoA for fatty acid formation in plastids [7]. Several independent approaches now indicate that PDHC is the major source for lipid synthesis. The expression of the mRNA for the subunits of PDHC correlates temporally and spatially with lipid accumulation in Arabidopsis embryos, while ACS mRNA does not [8]. Furthermore, analysis of microarray data indicates that plastid PDHC and the fatty acid biosynthesis genes are co-expressed in a single regulon [9,10]. Also, knocking out the ACS gene in Arabidopsis does not alter lipid levels in seeds [11], while disrupting the E2 subunit of PDHC results in an early embryo lethal mutation [12]. These molecular genetic experiments have been reinforced by metabolic measurements that first suggested there was insufficient acetate to account for the rates of fatty acid accumulation [13] and showed that the flux rate through plastid pyruvate was sufficient to account for fatty acid synthesis in that organelle [14,15]. This body of research established the role of PDHC in producing acetyl-CoA for plastidic fatty acid synthesis, but left no obvious function for ACS. 2.2. Acetyl-CoA in the cytosol
Fig. 1. Acetyl-CoA metabolism occurs in at least five subcellular compartments. Acetyl-CoA is produced and consumed in the plastid, mitochondria, peroxisome, and cytosol. Because of its limited permeability through biological membranes and the apparent lack of acetyl-CoA transporters in plants, different sources of acetylCoA are needed in each of these compartments [2]. Acetyl-CoA is also required in the nucleus for acetylation of histones [46,47], however, nuclei contain large pores and it is not known whether it is synthesized in the nucleus or translocated from the cytosol. Isoprenoids are likely produced from IPP synthesized in three compartments; in the cytosol and peroxisomes this IPP would be acetyl-CoAderived (i.e., the mevalonic acid pathway). Identification of the localization(s) of all the enzymes from acetyl-CoA to IPP, combined with understanding the mechanisms of the variable permeability of IPP across organelle membranes, will clarify the subcellular source of acetyl-CoA [16,18,19]. Acetylation and malonation occur in the cytosol. In addition (omitted from this figure for clarity) a wide variety of acetylated and malonated metabolites, such as polypeptides, lignin, and specialized plant products, are present in other subcellular/extracellular compartments [48]. Though not well studied, it is likely that a number of these are synthesized in situ from a local pool of acetyl-CoA.
Cytosolic acetyl-CoA is essential for elongating the plastidproduced fatty acids for deposition in suberin, the cuticle, and seed oils. This acetyl-CoA pool is also necessary for producing flavonoids, stilbenoids, a range of malonylated and acetylated metabolites. In addition, it probably supplies IPP substrate to support the synthesis of several classes of isoprenoids, including membrane sterols, brassinosteroids, sesquiterpenes, and to a lesser extent plastid isoprenoids. The extent to which cytosolic acetylCoA contributes to isoprenoids is not well understood, because IPP is variably permeable to membranes, thus IPP produced in one compartment can potentially contribute to isoprenoids made in another compartment. In addition, while the (acetyl-CoA-derived) mevalonic acid pathway for isoprenoid synthesis is usually considered to be cytosolic [16] an exclusively cytosolic subcellular localization was questioned by the discovery of acetoacetyl-CoA thiolase in peroxisomes in a proteomics experiment [17], localization studies [18] indicating a part of the isoprenoid pathway is peroxisomal, and biochemical and informatics evidence suggest that IPP isomerases are targeted to plastids, peroxisomes and the cytosol [19]. Cytosolic acetyl-CoA is synthesized by a a4b4 heteromeric ATPcitrate lyase in Arabidopsis [1]. No other major source of cytosolic acetyl CoA has been identified to date (although the rapid incorporation of acetate into fatty acids extended in the cytosol
D.J. Oliver et al. / Plant Science 176 (2009) 597–601
suggests another pathway may exist to move carbon in plastid acetyl-CoA to the cytosol [20]). Indeed, even modestly decreasing the ATP-citrate lyase activity by antisense RNA technology results in small ‘‘bonsai’’ plants having complex metabolic changes and stress responses [2]. Many of these phenotypes can be overcome by providing exogenous malonate. The ultimate source of carbon for cytosolic acetyl-CoA is mitochondrial, with a citrate/oxaloacetate shuttle envisioned that, together with citrate synthase in the mitochondrial matrix and ATP-citrate lyase in the cytosol, moves acetate-equivalents from the matrix to the cytosol. Metabolic flux experiments seem to support this model [14,15]. 2.3. Acetyl-CoA in the glyoxysomes Acetyl-CoA within the glyoxysomes is produced from fatty acid breakdown and feeds into the glyoxylate shunt. This pathway is most important during early germination and is an essential process in the efficient conversion of fatty acids to sucrose. The pathway is also important during stress cycles when autophagic degradation of membrane lipids is necessary for cell survival [21,22]. While the role of this shunt has been appreciated for many years, the application of Arabidopsis genetics has recently provided exciting insights into the mechanism of fatty acid uptake into the glyoxysomes, the specific reactions involved, and the interplay of organelle and cytosolic metabolism. For a detailed recent review see [23].
599
an important role in detoxifying these fermentation intermediates. This decreased growth of the acs1 plants suggests an ongoing role for the ACS metabolic pathway in detoxifying fermentative intermediates even in a low stress environment. Two unanswered questions that arise from this research are what the role of ACS under normal conditions is, and what is the source of the acetate that ACS appears to detoxify. Two potential metabolic roles have been suggested (Fig. 2). There is substantial evidence for a PDHC-bypass in plants [28,29]. This ‘‘aerobic fermentation’’ pathway utilizes pyruvate produced by glycolysis, and oxidizes it first to acetaldehyde (catalyzed by pyruvate decarboxylase) and then to acetate (catalyzed by acetaldehyde dehydrogenase). ACS would then activate the acetate to acetyl-CoA as a substrate for fatty acid synthesis (Fig. 2A). This pathway appears to function in all tissues and organs of plants, although it is expressed at highest levels in flowers. It has been suggested that this pyruvate bypass pathway may be essential for providing high metabolic rates in tissues such as growing pollen tubes, which may be under hypoxic conditions [30]. Indeed acs1 plants have decreased male fertility [11] and the bypass pathway can partially alleviate blockages in PDHC [30].
2.4. Acetyl-CoA in the mitochondria In mitochondria acetyl-CoA is predominately converted to citrate, which is either further metabolized by the TCA cycle or exported to the cytosol. In addition, the mitochondria of barley and possibly all grasses appear to contain a multifunctional, homomeric acetyl-CoA carboxylase that can convert acetyl-CoA to malonyl-CoA [24]. The malonyl-CoA is predominately directed toward octanoate as a precursor for forming lipoic acid, an essential cofactor for glycine decarboxylase and a-keto acid dehydrogenase complexes in the matrix. Some longer fatty acids are also produced and may be used for repairing mitochondrial lipids. Dicots examined to date lack this enzyme and likely take in malonate for reaction with CoASH or acyl carrier protein to form the 3-ketoacyl carrier protein synthase substrate [25]. 3. Some unanswered questions about acetyl-CoA metabolism 3.1. The physiological function of acetate metabolism in plants The role of acetate metabolism in plants is still unclear. Exogenous acetate is readily converted to acetyl-CoA, the plastid ACS providing about 90% of the catalysis [11] and a peroxisomal acyl-CoA synthetase [26] supplying the rest. Surprisingly, Arabidopsis plants can grow with acetate as their major carbon source and this growth is blocked in the acs1 mutant [11]. Characterization of transgenic Arabidopsis plants expressing ACS ranging from 5% to 500% of wild-type levels shows that ACS levels control the rate of exogenous acetate and ethanol conversion to fatty acids [27]. These characterizations also indicate that acetate metabolism in vivo is not limited by the available ACS activity, but rather, by the availability of the substrate, acetate. Endogenous acetate levels have been estimated as ranging from 0.05 mM to 1.4 mM in plants [13 and cited references]. The ACS knockout mutant, acs1, has a modest morphological phenotype; plants that lack this enzyme are smaller and flower a few days later than wild-type [11]. The growth rate of acs1 plants is more sensitive to exogenous acetate, ethanol and acetaldehyde than wild-type, suggesting that ACS has
Fig. 2. Proposed roles for acetyl-CoA synthetase in plants. (A) The pyruvate dehydrogenase bypass where acetyl-CoA can be formed from pyruvate under conditions of limited PDHC activity. The plastidic PDCH reaction is in red and the bypass which resides in the mitochondria, cytosol, and plastid is in blue. (B) A proposed new role for acetyl-CoA metabolism in leaves. In this model ethanol and/ or acetaldehyde produced in roots in the presence of limited O2 is transported to shoots where it is either lost from the plant as volatile organic compounds or oxidized to acetate which can be converted into fatty acids.
600
D.J. Oliver et al. / Plant Science 176 (2009) 597–601
An alternative source for acetate (or its precursors) in the aerial portions of plants may be from organs such as roots and stem that are more prone to low oxygen stress. The fermentation intermediates are then translocated to the leaves via the transpirationstream [31]. Two-carbon fermentation intermediates, ethanol and acetaldehyde, are part of the collection of volatile organic compounds that plants emit into the environment. These compounds could either be transported to the leaves or be produced in the leaves by aerobic fermentation processes [32]. Precursor-feeding experiments suggest that these compounds are formed in the roots and delivered to the leaves. Imposing anaerobic conditions on the roots increases acetaldehyde release from the leaves, while inhibition of alcohol dehydrogenase activity in the leaves decreases acetaldehyde release [33]. These data suggest that ethanol produced in roots is transported to leaves where it is oxidized to acetaldehyde and acetate. Therefore, ACS may be part of a salvage and detoxification pathway that allows leaves to capture fermentative products produced in the roots, keeping their concentrations below toxic levels while recapturing some of the lost carbon (Fig. 2B). It is interesting to postulate that this pathway might partially contribute to increased survival in hypoxic conditions. Clearly the availability of mutants in these pathways will make it possible to test this hypothesis. Another interesting question is the source of acetyl-CoA for histone acetylation. Because acetylation impacts histone function in the nucleus, the methods by which acetylation occurs could be important for understanding histone behavior and its relation to the control of gene expression. 3.2. Triacylglycerols in leaves An under appreciated set of observations that may broaden our understanding of the metabolic role of acetyl-CoA in leaves is the finding that some plants accumulate triacylglycerol in leaves (see [34,35] for detailed reviews of this literature). Leaf mesophyll cells of about a quarter of the plants microscopically surveyed contained one or more lipid bodies [35]. Chemical analyses of the lipids present in leaves have documented that they are triacylglycerols and that the fatty acids associated with this lipidclass have shorter chain length and fewer double bonds than those in the membrane lipids [36,37]. The fatty acids were in fact more similar to those found in the seed triacylglycerol from these same plants than to the leaf membrane lipids. Interestingly, the amount of triacylglycerol in crabapple leaves was lower in the morning than in the evening suggesting that their levels were controlled by diurnal events [37]. It is possible that these leaf triacylglycerols might be diurnal energy and carbon stores, in part filling the role traditionally assigned to starch in these plants, although 14CO2 labeling in wheat suggested that this might not be the case [38]. In crabapple leaves calculations based on the differences in TAG levels during the day suggest that about 5% of the total photosynthate was incorporated into triacylglycerol in these leaves [37]. This is an intriguing idea and would provide a new role for lipids in leaves. Certainly this is an area that deserves additional study. Recent speculation [39] has focused on the possibility and advantages of producing sufficient oil in leaves to make them a commercially viable source of lipids for biodiesel formation. The observation that the expression of WRI1 transcription factor causes oil accumulation in seedling [40] hints at an approach that might produce crops with elevated TAG in leaves. It is possible that while TAG production in oil seeds is reaching practical maximal limits and thus it has been difficult to increase levels significantly, this may not be the case in leaves. Similarly leaves might be more flexible than seeds in accumulating TAG with unusual fatty acids a goal that has proven difficult to reach in seeds [41].
4. Metabolic engineering of acetyl-CoA metabolism for production of biorenewable chemicals The earth contains finite quantities of fossil-carbon and the price of these commodities is highly volatile. While it may be debatable whether there is a role for biologically derived replacements for petroleum as a liquid fuel, there is little debate that some form of biological carbon will ultimately be needed to replace oil as a precursor for common petrochemicals [42]. Commodity chemicals are currently produced from light petroleum distillates by first cracking them to ethylene and propylene and then using these platform chemicals along with benzene to synthesize most of the high volume chemical products needed. While a number of specialty chemicals including glycerol, lactate, and 1,3-propanediol are produced biologically, ultimately we will need to produce the same types of platform chemicals that will serve as feedstocks for the chemical industry. Given that more reduced-carbon intermediates of the polyketide biosynthesis pathways (e.g., fatty acids, stilbenes and chalcones) are more chemically flexible than the more oxidized carbohydrates, the polyketide intermediates are logical platform chemicals to be produced from biological sources [42]. The fatty acid biosynthesis pathway is a familiar standard for polyketide biosynthesis. In plastids acetyl-CoA acts both to prime the reactions of fatty acid synthesis and to extend the fatty acid chain following its conversion to malonyl-CoA and malonyl-ACP. The extension reactions initiate with a condensation that is catalyzed by a family of 3-ketoacyl-acyl carrier protein synthases (three isoforms with specificity for different carbon chain lengths are used). Intermediates of fatty acid biosynthesis are even numbered, four or more carbon atoms, along with their b-keto, bhydroxyl, and a,b-enoyl intermediates. The condensing enzymes, reductases, and dehydratases that catalyze these reactions are collectively referred to as fatty acid synthase, which more generally belong to a family of enzymes known as polyketide synthases. While the intermediates that could be released from this process are diverse, this chemical diversity could be further expanded by altering the range of condensing enzymes, thioester elongation substrates, and chain terminating reactions involved. For example, odd number and branched chain fatty acids are synthesized in gram positive bacteria, such as Bacillus, and in plants in the trichomes of some Solanaceae species where they are esterified to sugars. In this latter system these branched chain fatty acids appear to be produced by two mechanisms. Both are primed with branched chain a-keto acids. In some species the aketo acids are elongated via the fatty acid synthesis complex. In other species a one-carbon extension mechanism termed a-keto acid elongation (a-KAE) is used [43,44]. Such a process could increase the diversity of chemicals produced. Multiple branch points could be introduced into the acyl chain by using methylmalonyl-CoA as the elongation substrate instead of malonyl-CoA, such a system occurs in mycobacteria such as Mycobacterium tuberculosis var. bovis [45]. Acetyl-CoA is an important metabolic intermediate that is both produced and consumed within several subcellular compartments using different biocatalysts and control functions. Recently molecular, biochemical, and metabolic studies have identified many of the key reactions and their physiological roles. These discoveries have provided the basic knowledge that will be needed to open new avenues for exploring the metabolism associated with acetyl-CoA and to direct plant metabolism in new directions. Acknowledgements This research is funded by support from the U.S. Department of Energy (DE-FG02-01ER15170) and from the U.S. National Science
D.J. Oliver et al. / Plant Science 176 (2009) 597–601
Foundation Center for BioRenewable Chemicals (http://www.cbirc.iastate.edu/) and MCB-0416730.
[25]
References [26] [1] B.L. Fatland, J. Ke, M.D. Anderson, W.I. Mentzen, L.W. Cui, C.C. Allred, J.L. Johnston, B.J. Nikolau, E.S. Wurtele, Molecular characterization of a heteromeric ATP-citrate lyase that generates cytosolic acetyl-coenzyme A in Arabidopsis, Plant Phys. 130 (2002) 740–756. [2] B.L. Fatland, B.J. Nikolau, E.S. Wurtele, Reverse genetic characterization of cytosolic acetyl-CoA generation by ATP-citrate lyase in Arabidopsis, Plant Cell 17 (2005) 182–203. [3] B.P. Smirnov, The biosynthesis of higher acids from acetate in intact chloroplasts of Spinacea oleracea leaves, Biokhimii 25 (1960) 419–426. [4] P.G. Roughan, C.R. Slack, R. Holland, High rates of [1-14C] acetate incorporation into the lipid of isolated spinach chloroplasts, Biochem. J. 158 (1976) 593–601. [5] D.N. Kuhn, M. Knauf, P.K. Stumpf, Subcellular localization of acetyl-CoA synthetase in leaf protoplasts of Spinacia oleracea leaf cells, Arch. Biochem. Biophys. 209 (1981) 441–450. [6] E.E. Reid, P. Thompson, C.R. Lyttle, D.T. Dennis, Pyruvate dehydrogenase complex from higher plant mitochondria and proplastids, Plant Physiol. 59 (1977) 842– 848. [7] J. Ohlrogge, J. Browse, Lipid biosynthesis, Plant Cell 7 (1995) 957–970. [8] J. Ke, R.H. Behal, S. Yunker, B.J. Nikolau, E.S. Wurtele, D.J. Oliver, The role of pyruvate dehydrogenase and acetyl-CoA synthetase in fatty acid synthesis in developing Arabidopsis seeds, Plant Physiol. 123 (2000) 497–508. [9] W. Mentzen, E.S. Wurtele, Regulon organization of Arabidopsis, BMC Plant Biol. 8 (2008) 99 http://www.biomedcentral.com/1471-2229/8/99. [10] W. Mentzen, J.L. Peng, B.J. Nikolau, E.S. Wurtele, Articulation of core metabolic processes in Arabidopsis, BMC Plant Biol. 8 (2008) 76 http://www.biomedcentral.com/1471-2229/8/76. [11] M. Lin, D.J. Oliver, The role of acetyl-CoA synthetase in Arabidopsis, Plant Phys. 147 (2008) 1822–1829. [12] M. Lin, R.H. Behal, D.J. Oliver, Disruption of plE2, the gene for the E2 subunit of the plastid pyruvate dehydrogenase complex, in Arabidopsis causes an early embryo lethal phenotype, Plant Mol. Biol. 52 (2003) 865–872. [13] X. Bao, M. Focke, M. Pollard, J. Ohlrogge, Understanding the in vivo carbon precursor supply for fatty acid synthesis in leave tissue, Plant J. 22 (2000) 39–50. [14] J. Schwender, J.B. Ohlrogge, Y. Shachar-Hill, A flux model of glycolysis and the oxidative pentose phosphate pathway in developing Brassica napus embryos, J. Biol. Chem. 278 (2003) 29442–29453. [15] J. Schwender, Y. Shachar-Hill, J.B. Ohlrogge, Mitochondrial metabolism in developing embryos of Brassica napus, J. Biol. Chem. 281 (2006) 34040–34047. [16] J.D. Newman, J. Chappell, Isoprenoid biosynthesis in plants: carbon partitioning within the cytoplasmic pathway, Crit. Rev. Biochem. Mol. Biol. 34 (1999) 95–106. [17] S. Reumann, L. Babujee, C. Ma, S. Wienkoop, T. Siemsen, G.E. Antonicelli, N. Rasche, F. Luder, W. Weckwerth, O. Jahn, Proteome analysis of Arabidopsis leaf peroxisomes reveals novel targeting peptides, metabolic pathways, and defense mechanisms, Plant Cell 19 (2007) 3170–3193. [18] M. Sapir-Mir, A. Mett, E. Belausov, S. Tal-Meshulam, A. Frydman, D. Gidoni, Y. Eyal, Peroxisomal localization of Arabidopsis isopentenyl diphosphate isomerases suggest that part of the plant isoprenoids mevalonic acid pathway is compartmentalized to peroxisomes, Plant Physiol. 148 (2008) 1219–1228. [19] M.A. Phillips, J.C. D’Auria, J. Gershenzon, E. Pichersky, The Arabidopsis thaliana type I isopentenyl diphosphate isomerases are targeted to multiple subcellular compartments and have overlapping functions in isoprenoid biosynthesis, Plant Cell 20 (2008) 677–696. [20] J. Ohlrogge, M.E. Pollard, P.K. Stumpf, Studies on biosynthesis of waxes by developing jojoba seed tissue, Lipids 13 (1978) 203–210. [21] P.J. Eastmond, V. Germain, P.R. Lange, J.H. Bryce, S.M. Smith, I.A. Graham, Postgerminative growth and lipid catabolism in oilseeds lacking the glyoxylate cycle, Proc. Natl. Acad. Sci. U.S.A. 97 (2000) 5669–56674. [22] I. Ismail, L. De Bellis, A. Alpi, S.M. Smith, Expression of glyoxylate cycle genes in cucumber roots responds to sugar supply and can be activated by shading or defoliation of the shoot, Plant Mol. Biol. 35 (1997) 633–640. [23] I.A. Graham, Seed storage oil mobilization, Annu. Rev. Plant Biol. 59 (2008) 115–142. [24] M. Focke, E. Gieringer, S. Schwan, L. Jansch, S. Binder, H.-P. Brawn, Fatty acid biosynthesis in mitochondria of grasses: malonyl-coenzyme A is generated by a
[27] [28] [29]
[30]
[31] [32]
[33]
[34] [35]
[36]
[37] [38] [39] [40]
[41]
[42] [43]
[44]
[45]
[46]
[47] [48]
601
mitochondrial-localized acetyl-coenzyme A carboxylase, Plant Physiol. 133 (2003) 875–884. V. Gueguen, D. Macherel, J. Jaquinod, R. Douce, J. Bourguignon, Fatty acid and lipoic acid biosynthesis in higher plant mitochondria, J. Biol. Chem. 275 (2000) 5016–5025. J.E. Turner, K. Greville, E.C. Murphy, M.A. Hooks, Characterization of Arabidopsis fluoroacetate-resistant mutants reveals the principal mechanism of acetate activation for entry into the glyoxylate cycle, J. Biol. Chem. 280 (2005) 2780– 2787. R.H. Behal, M. Lin, S. Back, D.J. Oliver, Role of acetyl-CoA synthetase in the leaves of Arabidopsis thaliana, Arch. Biochem. Biophys. 402 (2002) 259–267. M. Tadege, I. Dupuis, C. Kuhlemeier, Ethanolic fermentation: new functions for an old pathway, Trends Plant Sci. 4 (1999) 320–325. S. Mellema, W. Eichenberger, A. Rawyler, M. Suter, M. Tadege, C. Kuhlemeier, The ethanolic fermentation pathway supports respiration and lipid biosynthesis in tobacco pollen, Plant J. 30 (2002) 329–336. N. Gass, T. Glagotskaia, S. Mellema, J. Stuurman, M. Barone, T. Mandel, U. Roessner-Tunali, C. Kuhlemeier, Pyruvate decarboxylase provides growing pollen tubes with a competitive advantage in petunia, Plant Cell 7 (2005) 2355–2368. J. Kreuzwieser, U. Scheerer, H. Rennenberg, Metabolic origin of acetaldehyde emitted by poplar (Populus tremula P. alba) trees, J. Exp. Bot. 50 (1999) 757–765. T. Karl, A.J. Curtis, T.N. Rosenstiel, R.K. Monson, R. Fal, Transient releases of acetaldehyde from tree leaves—products of a pyruvate overflow mechanism? Plant Cell Environ. 25 (2002) 1121–1131. J. Kreuzwieser, F.J.M. Harren, L.J.J. Lerhoven, I. Boamfa, S. Linten-Hekker, U. Scheerer, C. Huglin, H. Rennenberg, Acetaldehyde emission from the leaves of trees—correlation with physiological and environmental parameters, Physiol. Plant. 113 (2001) 41–49. D.J. Murphy, The biogenesis and function of lipid bodies in animals, plants, and microorganisms, Prog. Lipid Res. 40 (2001) 325–438. N.R. Lersten, A.R. Czlapinski, J.D. Curtis, R. Freckmann, H.T. Horner, Oil bodies in leaf mesophyll cells of angiosperms: overview and a selected survey, Am. J. Bot. 93 (2006) 1731–1739. M.L. Parker, G.J.P. Murphy, Oleosomes in flag leaves of wheat; their distribution, composition and fate during senescence and rust-infection, Planta 152 (1981) 36– 43. W. Lin, D.J. Oliver, Role of triacylglycerols in leaves, Plant Sci. 175 (2008) 233–237. G.J.P. Murphy, M.L. Parker, Lipid composition and carbon turnover of wheat leaf oleosomes, J. Exp. Bot. 35 (1984) 348–355. T.P. Durrett, C. Benning, J. Ohlrogge, Plant triacylglycerols as feedstocks for the production of biofuels, Plant J. 54 (2008) 593–607. A. Cernac, C. Benning, WRINKLED1 encodes an AP2/EREB domain protein involved in the control of storage compound biosynthesis in Arabidopsis, Plant J. 40 (2004) 575–585. J.M. Dyer, R.T. Mullen, Engineering plant oils as high-value industrial feedstocks for biorefining: the need for underpinning cell biology research, Physiol. Plant. 132 (2008) 11–22. B.J. Nikolau, M. Perera, L. Brachova, B. Shanks, Platform biochemicals for a biorenewable chemical industry, Plant J. 54 (2008) 536–545. A.B. Kroumova, G.J. Wagner, Different elongation pathways in the biosynthesis of acyl groups of trichome exudate sugar esters from various solanaceous plants, Planta 216 (2003) 1013–1021. A.B. Kroumova, Z. Xie, G.J. Wagner, A pathway for the biosynthesis of straight and branched, odd- and even-length, medium-chain fatty acids in plants, Proc. Natl. Acad. Sci. U.S.A. 91 (1994) 11437–11441. D.L. Rainwater, P.E. Kolattukudy, Fatty acid biosynthesis in Mycobacterium tuberculosis var. bovis Bacillus Calmette-Guerin. Purification and characterization of a novel fatty acid synthase, mycocerosic acid synthase, which elongates n-fatty acyl-CoA with methylmalonyl-CoA, J. Biol. Chem. 260 (1985) 616–623. K. Earley, R.J. Lawrence, O. Pontes, R. Reuther, A.J. Enciso, M. Silva, N. Neves, M. Gross, W. Viegas, C.S. Pikaard, Erasure of histone acetylation by Arabidopsis HDA6 mediates large-scale gene silencing in nucleolar dominance, Genes Dev. 20 (2006) 1283–1293. N. Geraldo, R. Abranches, Immunolocalization of histone modifications as a tool to visualize chromatin dynamics in plants, Microsc. Microanal. 14 (2008) 130–133. A.T. Martı´nez, J. Rencoret, G. Marques, A. Gutie´rrez, D. Ibarra, J. Jime´nez-Barbero, Jose´ C. del Rı´o, Monolignol acylation and lignin structure in some nonwoody plants: a 2D NMR study, Phytochemistry 69 (2008) 2831–2843.
Contents lists available at ScienceDirect
Plant Science journal homepage: www.elsevier.com/locate/plantsci
Review
Acetyl-CoA—Life at the metabolic nexus David J. Oliver a,*, Basil J. Nikolau b, Eve Syrkin Wurtele a a b
Department of Genetics, Development and Cell Biology, Iowa State University, Ames, IA 50011, USA Department of Biochemistry, Biophysics and Molecular Biology, Iowa State University, Ames, IA 50011, USA
A R T I C L E I N F O
A B S T R A C T
Article history: Received 27 October 2008 Received in revised form 10 February 2009 Accepted 10 February 2009 Available online 20 February 2009
As a key intermediate in a number of different metabolic pathways, acetyl-CoA is independently produced and consumed in the plastids, mitochondria, peroxisomes and cytosol. Recent studies have led to new understanding of the enzymes that generate these different acetyl-CoA pools, including the role of ATP-citrate lyase in producing acetyl-CoA in the cytosol and of the pyruvate dehydrogenase complex for making it in the plastids. The involvement of a second plastid enzyme, acetyl-CoA synthetase, is more speculative but appears to be important in protecting plants from the two-carbon intermediates of fermentation by allowing the conversion of ethanol and acetaldehyde to fatty acids. New roles have also recently been proposed for triacylglycerols in energy storage and oil production in leaves. While additional biological functions for acetyl-CoA are being discovered there are also new proposals for using the chemistry of acetyl-CoA and fatty acid synthesis as the foundation for a new biologically based source of commodity chemicals. ß 2009 Elsevier Ireland Ltd. All rights reserved.
Keywords: Anaerobic metabolism Fatty acids Metabolic engineering Triacylglycerols
Contents 1. 2.
3.
4.
Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Recent breakthroughs in acetyl-CoA metabolism. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. Acetyl-CoA in the plastid . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. Acetyl-CoA in the cytosol. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3. Acetyl-CoA in the glyoxysomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4. Acetyl-CoA in the mitochondria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Some unanswered questions about acetyl-CoA metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1. The physiological function of acetate metabolism in plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2. Triacylglycerols in leaves . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Metabolic engineering of acetyl-CoA metabolism for production of biorenewable chemicals. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
1. Introduction Acetyl-CoA is a metabolite at a key point connecting catabolic and anabolic metabolism. It is produced by the catabolism of carbohydrates, lipids, and amino acids. At the same time it is the precursor for the synthesis of oils, membrane lipids, cuticle, suberin, isoprenoids, flavonoids, stilbenoids, some amino acids, and sometimes sugars. Because of the unique central metabolic
* Corresponding author. Tel.: +1 515 294 4118; fax: +1 515 294 1303. E-mail address: [email protected] (D.J. Oliver). 0168-9452/$ – see front matter ß 2009 Elsevier Ireland Ltd. All rights reserved. doi:10.1016/j.plantsci.2009.02.005
597 598 598 598 599 599 599 599 600 600 600 601
position occupied by acetyl-CoA, flux through this intermediate ultimately controls the rate that carbon reserves in plants can be converted into many important chemicals. Plans for engineering plants for optimal production of proteins, lipids, and specialty chemicals must consider the systems that produce acetyl-CoA as a substrate for these products. It is surprising therefore that so many questions about the basic biochemistry of acetyl-CoA production and use in plant cells remain unanswered. The purpose of this review is threefold. First, we provide a brief overview of recent breakthroughs in the understanding of acetylCoA metabolism in plants. Second, we discuss some unanswered questions concerning acetate metabolism. Finally, we describe a
598
D.J. Oliver et al. / Plant Science 176 (2009) 597–601
vision of how to change acetyl-CoA metabolic pathways to turn plants into engines for producing commodity chemicals. 2. Recent breakthroughs in acetyl-CoA metabolism Acetyl-CoA metabolism occurs in many subcellular compartments in a plant cell. The chloroplasts, peroxisomes, mitochondria, cytosol, and nucleus are shown in Fig. 1 [1,2]. The origins and metabolic fates of acetyl-CoA are distinct in each subcellular compartment, with the plastid and cytosol acetyl-CoA pools being used primarily in anabolic processes, whereas the mitochondrial and peroxisomal pools are used in both anabolic and catabolic processes. Acetyl-CoA is produced in plastids from pyruvate and acetate and supplies precursors for the biosynthesis of fatty acids, amino acids, and glucosinolates. Cytosolic acetyl-CoA is largely produced from citrate and provides carbon for elongation of fatty acids, which are converted to a variety of chemicals and is condensed yielding other polyketides. Cytosolic acetyl-CoA has
also been implicated in the synthesis of secondary metabolites including flavonoids, stilbenoids, and isoprenoids. Peroxisomal acetyl-CoA is formed from fatty acid breakdown and is the substrate for the glyoxylate shunt. In mitochondria acetyl-CoA is made from pyruvate oxidation and is the substrate for the TCA cycle and in some cases fatty acid biosynthesis. Because acetyl-CoA cannot readily cross membranes, the subcellular organization can be rationalized on the basis of providing plants unique regulatory mechanisms to control acetyl-CoA metabolism for different metabolic purposes. 2.1. Acetyl-CoA in the plastid Plastidic acetyl-CoA is synthesized by two enzyme systems, the pyruvate dehydrogenase complex (PDHC) and acetyl-CoA synthetase (ACS). ACS was discovered first [3] and for years it was considered a primary source of acetyl-CoA for lipid synthesis, although the source of its acetate substrate was never clear [4,5]. The discovery of PDHC provided a more direct connection between acetyl-CoA formation and central metabolism [6], still there was little indication as to which enzyme was the primary source of acetyl-CoA for fatty acid formation in plastids [7]. Several independent approaches now indicate that PDHC is the major source for lipid synthesis. The expression of the mRNA for the subunits of PDHC correlates temporally and spatially with lipid accumulation in Arabidopsis embryos, while ACS mRNA does not [8]. Furthermore, analysis of microarray data indicates that plastid PDHC and the fatty acid biosynthesis genes are co-expressed in a single regulon [9,10]. Also, knocking out the ACS gene in Arabidopsis does not alter lipid levels in seeds [11], while disrupting the E2 subunit of PDHC results in an early embryo lethal mutation [12]. These molecular genetic experiments have been reinforced by metabolic measurements that first suggested there was insufficient acetate to account for the rates of fatty acid accumulation [13] and showed that the flux rate through plastid pyruvate was sufficient to account for fatty acid synthesis in that organelle [14,15]. This body of research established the role of PDHC in producing acetyl-CoA for plastidic fatty acid synthesis, but left no obvious function for ACS. 2.2. Acetyl-CoA in the cytosol
Fig. 1. Acetyl-CoA metabolism occurs in at least five subcellular compartments. Acetyl-CoA is produced and consumed in the plastid, mitochondria, peroxisome, and cytosol. Because of its limited permeability through biological membranes and the apparent lack of acetyl-CoA transporters in plants, different sources of acetylCoA are needed in each of these compartments [2]. Acetyl-CoA is also required in the nucleus for acetylation of histones [46,47], however, nuclei contain large pores and it is not known whether it is synthesized in the nucleus or translocated from the cytosol. Isoprenoids are likely produced from IPP synthesized in three compartments; in the cytosol and peroxisomes this IPP would be acetyl-CoAderived (i.e., the mevalonic acid pathway). Identification of the localization(s) of all the enzymes from acetyl-CoA to IPP, combined with understanding the mechanisms of the variable permeability of IPP across organelle membranes, will clarify the subcellular source of acetyl-CoA [16,18,19]. Acetylation and malonation occur in the cytosol. In addition (omitted from this figure for clarity) a wide variety of acetylated and malonated metabolites, such as polypeptides, lignin, and specialized plant products, are present in other subcellular/extracellular compartments [48]. Though not well studied, it is likely that a number of these are synthesized in situ from a local pool of acetyl-CoA.
Cytosolic acetyl-CoA is essential for elongating the plastidproduced fatty acids for deposition in suberin, the cuticle, and seed oils. This acetyl-CoA pool is also necessary for producing flavonoids, stilbenoids, a range of malonylated and acetylated metabolites. In addition, it probably supplies IPP substrate to support the synthesis of several classes of isoprenoids, including membrane sterols, brassinosteroids, sesquiterpenes, and to a lesser extent plastid isoprenoids. The extent to which cytosolic acetylCoA contributes to isoprenoids is not well understood, because IPP is variably permeable to membranes, thus IPP produced in one compartment can potentially contribute to isoprenoids made in another compartment. In addition, while the (acetyl-CoA-derived) mevalonic acid pathway for isoprenoid synthesis is usually considered to be cytosolic [16] an exclusively cytosolic subcellular localization was questioned by the discovery of acetoacetyl-CoA thiolase in peroxisomes in a proteomics experiment [17], localization studies [18] indicating a part of the isoprenoid pathway is peroxisomal, and biochemical and informatics evidence suggest that IPP isomerases are targeted to plastids, peroxisomes and the cytosol [19]. Cytosolic acetyl-CoA is synthesized by a a4b4 heteromeric ATPcitrate lyase in Arabidopsis [1]. No other major source of cytosolic acetyl CoA has been identified to date (although the rapid incorporation of acetate into fatty acids extended in the cytosol
D.J. Oliver et al. / Plant Science 176 (2009) 597–601
suggests another pathway may exist to move carbon in plastid acetyl-CoA to the cytosol [20]). Indeed, even modestly decreasing the ATP-citrate lyase activity by antisense RNA technology results in small ‘‘bonsai’’ plants having complex metabolic changes and stress responses [2]. Many of these phenotypes can be overcome by providing exogenous malonate. The ultimate source of carbon for cytosolic acetyl-CoA is mitochondrial, with a citrate/oxaloacetate shuttle envisioned that, together with citrate synthase in the mitochondrial matrix and ATP-citrate lyase in the cytosol, moves acetate-equivalents from the matrix to the cytosol. Metabolic flux experiments seem to support this model [14,15]. 2.3. Acetyl-CoA in the glyoxysomes Acetyl-CoA within the glyoxysomes is produced from fatty acid breakdown and feeds into the glyoxylate shunt. This pathway is most important during early germination and is an essential process in the efficient conversion of fatty acids to sucrose. The pathway is also important during stress cycles when autophagic degradation of membrane lipids is necessary for cell survival [21,22]. While the role of this shunt has been appreciated for many years, the application of Arabidopsis genetics has recently provided exciting insights into the mechanism of fatty acid uptake into the glyoxysomes, the specific reactions involved, and the interplay of organelle and cytosolic metabolism. For a detailed recent review see [23].
599
an important role in detoxifying these fermentation intermediates. This decreased growth of the acs1 plants suggests an ongoing role for the ACS metabolic pathway in detoxifying fermentative intermediates even in a low stress environment. Two unanswered questions that arise from this research are what the role of ACS under normal conditions is, and what is the source of the acetate that ACS appears to detoxify. Two potential metabolic roles have been suggested (Fig. 2). There is substantial evidence for a PDHC-bypass in plants [28,29]. This ‘‘aerobic fermentation’’ pathway utilizes pyruvate produced by glycolysis, and oxidizes it first to acetaldehyde (catalyzed by pyruvate decarboxylase) and then to acetate (catalyzed by acetaldehyde dehydrogenase). ACS would then activate the acetate to acetyl-CoA as a substrate for fatty acid synthesis (Fig. 2A). This pathway appears to function in all tissues and organs of plants, although it is expressed at highest levels in flowers. It has been suggested that this pyruvate bypass pathway may be essential for providing high metabolic rates in tissues such as growing pollen tubes, which may be under hypoxic conditions [30]. Indeed acs1 plants have decreased male fertility [11] and the bypass pathway can partially alleviate blockages in PDHC [30].
2.4. Acetyl-CoA in the mitochondria In mitochondria acetyl-CoA is predominately converted to citrate, which is either further metabolized by the TCA cycle or exported to the cytosol. In addition, the mitochondria of barley and possibly all grasses appear to contain a multifunctional, homomeric acetyl-CoA carboxylase that can convert acetyl-CoA to malonyl-CoA [24]. The malonyl-CoA is predominately directed toward octanoate as a precursor for forming lipoic acid, an essential cofactor for glycine decarboxylase and a-keto acid dehydrogenase complexes in the matrix. Some longer fatty acids are also produced and may be used for repairing mitochondrial lipids. Dicots examined to date lack this enzyme and likely take in malonate for reaction with CoASH or acyl carrier protein to form the 3-ketoacyl carrier protein synthase substrate [25]. 3. Some unanswered questions about acetyl-CoA metabolism 3.1. The physiological function of acetate metabolism in plants The role of acetate metabolism in plants is still unclear. Exogenous acetate is readily converted to acetyl-CoA, the plastid ACS providing about 90% of the catalysis [11] and a peroxisomal acyl-CoA synthetase [26] supplying the rest. Surprisingly, Arabidopsis plants can grow with acetate as their major carbon source and this growth is blocked in the acs1 mutant [11]. Characterization of transgenic Arabidopsis plants expressing ACS ranging from 5% to 500% of wild-type levels shows that ACS levels control the rate of exogenous acetate and ethanol conversion to fatty acids [27]. These characterizations also indicate that acetate metabolism in vivo is not limited by the available ACS activity, but rather, by the availability of the substrate, acetate. Endogenous acetate levels have been estimated as ranging from 0.05 mM to 1.4 mM in plants [13 and cited references]. The ACS knockout mutant, acs1, has a modest morphological phenotype; plants that lack this enzyme are smaller and flower a few days later than wild-type [11]. The growth rate of acs1 plants is more sensitive to exogenous acetate, ethanol and acetaldehyde than wild-type, suggesting that ACS has
Fig. 2. Proposed roles for acetyl-CoA synthetase in plants. (A) The pyruvate dehydrogenase bypass where acetyl-CoA can be formed from pyruvate under conditions of limited PDHC activity. The plastidic PDCH reaction is in red and the bypass which resides in the mitochondria, cytosol, and plastid is in blue. (B) A proposed new role for acetyl-CoA metabolism in leaves. In this model ethanol and/ or acetaldehyde produced in roots in the presence of limited O2 is transported to shoots where it is either lost from the plant as volatile organic compounds or oxidized to acetate which can be converted into fatty acids.
600
D.J. Oliver et al. / Plant Science 176 (2009) 597–601
An alternative source for acetate (or its precursors) in the aerial portions of plants may be from organs such as roots and stem that are more prone to low oxygen stress. The fermentation intermediates are then translocated to the leaves via the transpirationstream [31]. Two-carbon fermentation intermediates, ethanol and acetaldehyde, are part of the collection of volatile organic compounds that plants emit into the environment. These compounds could either be transported to the leaves or be produced in the leaves by aerobic fermentation processes [32]. Precursor-feeding experiments suggest that these compounds are formed in the roots and delivered to the leaves. Imposing anaerobic conditions on the roots increases acetaldehyde release from the leaves, while inhibition of alcohol dehydrogenase activity in the leaves decreases acetaldehyde release [33]. These data suggest that ethanol produced in roots is transported to leaves where it is oxidized to acetaldehyde and acetate. Therefore, ACS may be part of a salvage and detoxification pathway that allows leaves to capture fermentative products produced in the roots, keeping their concentrations below toxic levels while recapturing some of the lost carbon (Fig. 2B). It is interesting to postulate that this pathway might partially contribute to increased survival in hypoxic conditions. Clearly the availability of mutants in these pathways will make it possible to test this hypothesis. Another interesting question is the source of acetyl-CoA for histone acetylation. Because acetylation impacts histone function in the nucleus, the methods by which acetylation occurs could be important for understanding histone behavior and its relation to the control of gene expression. 3.2. Triacylglycerols in leaves An under appreciated set of observations that may broaden our understanding of the metabolic role of acetyl-CoA in leaves is the finding that some plants accumulate triacylglycerol in leaves (see [34,35] for detailed reviews of this literature). Leaf mesophyll cells of about a quarter of the plants microscopically surveyed contained one or more lipid bodies [35]. Chemical analyses of the lipids present in leaves have documented that they are triacylglycerols and that the fatty acids associated with this lipidclass have shorter chain length and fewer double bonds than those in the membrane lipids [36,37]. The fatty acids were in fact more similar to those found in the seed triacylglycerol from these same plants than to the leaf membrane lipids. Interestingly, the amount of triacylglycerol in crabapple leaves was lower in the morning than in the evening suggesting that their levels were controlled by diurnal events [37]. It is possible that these leaf triacylglycerols might be diurnal energy and carbon stores, in part filling the role traditionally assigned to starch in these plants, although 14CO2 labeling in wheat suggested that this might not be the case [38]. In crabapple leaves calculations based on the differences in TAG levels during the day suggest that about 5% of the total photosynthate was incorporated into triacylglycerol in these leaves [37]. This is an intriguing idea and would provide a new role for lipids in leaves. Certainly this is an area that deserves additional study. Recent speculation [39] has focused on the possibility and advantages of producing sufficient oil in leaves to make them a commercially viable source of lipids for biodiesel formation. The observation that the expression of WRI1 transcription factor causes oil accumulation in seedling [40] hints at an approach that might produce crops with elevated TAG in leaves. It is possible that while TAG production in oil seeds is reaching practical maximal limits and thus it has been difficult to increase levels significantly, this may not be the case in leaves. Similarly leaves might be more flexible than seeds in accumulating TAG with unusual fatty acids a goal that has proven difficult to reach in seeds [41].
4. Metabolic engineering of acetyl-CoA metabolism for production of biorenewable chemicals The earth contains finite quantities of fossil-carbon and the price of these commodities is highly volatile. While it may be debatable whether there is a role for biologically derived replacements for petroleum as a liquid fuel, there is little debate that some form of biological carbon will ultimately be needed to replace oil as a precursor for common petrochemicals [42]. Commodity chemicals are currently produced from light petroleum distillates by first cracking them to ethylene and propylene and then using these platform chemicals along with benzene to synthesize most of the high volume chemical products needed. While a number of specialty chemicals including glycerol, lactate, and 1,3-propanediol are produced biologically, ultimately we will need to produce the same types of platform chemicals that will serve as feedstocks for the chemical industry. Given that more reduced-carbon intermediates of the polyketide biosynthesis pathways (e.g., fatty acids, stilbenes and chalcones) are more chemically flexible than the more oxidized carbohydrates, the polyketide intermediates are logical platform chemicals to be produced from biological sources [42]. The fatty acid biosynthesis pathway is a familiar standard for polyketide biosynthesis. In plastids acetyl-CoA acts both to prime the reactions of fatty acid synthesis and to extend the fatty acid chain following its conversion to malonyl-CoA and malonyl-ACP. The extension reactions initiate with a condensation that is catalyzed by a family of 3-ketoacyl-acyl carrier protein synthases (three isoforms with specificity for different carbon chain lengths are used). Intermediates of fatty acid biosynthesis are even numbered, four or more carbon atoms, along with their b-keto, bhydroxyl, and a,b-enoyl intermediates. The condensing enzymes, reductases, and dehydratases that catalyze these reactions are collectively referred to as fatty acid synthase, which more generally belong to a family of enzymes known as polyketide synthases. While the intermediates that could be released from this process are diverse, this chemical diversity could be further expanded by altering the range of condensing enzymes, thioester elongation substrates, and chain terminating reactions involved. For example, odd number and branched chain fatty acids are synthesized in gram positive bacteria, such as Bacillus, and in plants in the trichomes of some Solanaceae species where they are esterified to sugars. In this latter system these branched chain fatty acids appear to be produced by two mechanisms. Both are primed with branched chain a-keto acids. In some species the aketo acids are elongated via the fatty acid synthesis complex. In other species a one-carbon extension mechanism termed a-keto acid elongation (a-KAE) is used [43,44]. Such a process could increase the diversity of chemicals produced. Multiple branch points could be introduced into the acyl chain by using methylmalonyl-CoA as the elongation substrate instead of malonyl-CoA, such a system occurs in mycobacteria such as Mycobacterium tuberculosis var. bovis [45]. Acetyl-CoA is an important metabolic intermediate that is both produced and consumed within several subcellular compartments using different biocatalysts and control functions. Recently molecular, biochemical, and metabolic studies have identified many of the key reactions and their physiological roles. These discoveries have provided the basic knowledge that will be needed to open new avenues for exploring the metabolism associated with acetyl-CoA and to direct plant metabolism in new directions. Acknowledgements This research is funded by support from the U.S. Department of Energy (DE-FG02-01ER15170) and from the U.S. National Science
D.J. Oliver et al. / Plant Science 176 (2009) 597–601
Foundation Center for BioRenewable Chemicals (http://www.cbirc.iastate.edu/) and MCB-0416730.
[25]
References [26] [1] B.L. Fatland, J. Ke, M.D. Anderson, W.I. Mentzen, L.W. Cui, C.C. Allred, J.L. Johnston, B.J. Nikolau, E.S. Wurtele, Molecular characterization of a heteromeric ATP-citrate lyase that generates cytosolic acetyl-coenzyme A in Arabidopsis, Plant Phys. 130 (2002) 740–756. [2] B.L. Fatland, B.J. Nikolau, E.S. Wurtele, Reverse genetic characterization of cytosolic acetyl-CoA generation by ATP-citrate lyase in Arabidopsis, Plant Cell 17 (2005) 182–203. [3] B.P. Smirnov, The biosynthesis of higher acids from acetate in intact chloroplasts of Spinacea oleracea leaves, Biokhimii 25 (1960) 419–426. [4] P.G. Roughan, C.R. Slack, R. Holland, High rates of [1-14C] acetate incorporation into the lipid of isolated spinach chloroplasts, Biochem. J. 158 (1976) 593–601. [5] D.N. Kuhn, M. Knauf, P.K. Stumpf, Subcellular localization of acetyl-CoA synthetase in leaf protoplasts of Spinacia oleracea leaf cells, Arch. Biochem. Biophys. 209 (1981) 441–450. [6] E.E. Reid, P. Thompson, C.R. Lyttle, D.T. Dennis, Pyruvate dehydrogenase complex from higher plant mitochondria and proplastids, Plant Physiol. 59 (1977) 842– 848. [7] J. Ohlrogge, J. Browse, Lipid biosynthesis, Plant Cell 7 (1995) 957–970. [8] J. Ke, R.H. Behal, S. Yunker, B.J. Nikolau, E.S. Wurtele, D.J. Oliver, The role of pyruvate dehydrogenase and acetyl-CoA synthetase in fatty acid synthesis in developing Arabidopsis seeds, Plant Physiol. 123 (2000) 497–508. [9] W. Mentzen, E.S. Wurtele, Regulon organization of Arabidopsis, BMC Plant Biol. 8 (2008) 99 http://www.biomedcentral.com/1471-2229/8/99. [10] W. Mentzen, J.L. Peng, B.J. Nikolau, E.S. Wurtele, Articulation of core metabolic processes in Arabidopsis, BMC Plant Biol. 8 (2008) 76 http://www.biomedcentral.com/1471-2229/8/76. [11] M. Lin, D.J. Oliver, The role of acetyl-CoA synthetase in Arabidopsis, Plant Phys. 147 (2008) 1822–1829. [12] M. Lin, R.H. Behal, D.J. Oliver, Disruption of plE2, the gene for the E2 subunit of the plastid pyruvate dehydrogenase complex, in Arabidopsis causes an early embryo lethal phenotype, Plant Mol. Biol. 52 (2003) 865–872. [13] X. Bao, M. Focke, M. Pollard, J. Ohlrogge, Understanding the in vivo carbon precursor supply for fatty acid synthesis in leave tissue, Plant J. 22 (2000) 39–50. [14] J. Schwender, J.B. Ohlrogge, Y. Shachar-Hill, A flux model of glycolysis and the oxidative pentose phosphate pathway in developing Brassica napus embryos, J. Biol. Chem. 278 (2003) 29442–29453. [15] J. Schwender, Y. Shachar-Hill, J.B. Ohlrogge, Mitochondrial metabolism in developing embryos of Brassica napus, J. Biol. Chem. 281 (2006) 34040–34047. [16] J.D. Newman, J. Chappell, Isoprenoid biosynthesis in plants: carbon partitioning within the cytoplasmic pathway, Crit. Rev. Biochem. Mol. Biol. 34 (1999) 95–106. [17] S. Reumann, L. Babujee, C. Ma, S. Wienkoop, T. Siemsen, G.E. Antonicelli, N. Rasche, F. Luder, W. Weckwerth, O. Jahn, Proteome analysis of Arabidopsis leaf peroxisomes reveals novel targeting peptides, metabolic pathways, and defense mechanisms, Plant Cell 19 (2007) 3170–3193. [18] M. Sapir-Mir, A. Mett, E. Belausov, S. Tal-Meshulam, A. Frydman, D. Gidoni, Y. Eyal, Peroxisomal localization of Arabidopsis isopentenyl diphosphate isomerases suggest that part of the plant isoprenoids mevalonic acid pathway is compartmentalized to peroxisomes, Plant Physiol. 148 (2008) 1219–1228. [19] M.A. Phillips, J.C. D’Auria, J. Gershenzon, E. Pichersky, The Arabidopsis thaliana type I isopentenyl diphosphate isomerases are targeted to multiple subcellular compartments and have overlapping functions in isoprenoid biosynthesis, Plant Cell 20 (2008) 677–696. [20] J. Ohlrogge, M.E. Pollard, P.K. Stumpf, Studies on biosynthesis of waxes by developing jojoba seed tissue, Lipids 13 (1978) 203–210. [21] P.J. Eastmond, V. Germain, P.R. Lange, J.H. Bryce, S.M. Smith, I.A. Graham, Postgerminative growth and lipid catabolism in oilseeds lacking the glyoxylate cycle, Proc. Natl. Acad. Sci. U.S.A. 97 (2000) 5669–56674. [22] I. Ismail, L. De Bellis, A. Alpi, S.M. Smith, Expression of glyoxylate cycle genes in cucumber roots responds to sugar supply and can be activated by shading or defoliation of the shoot, Plant Mol. Biol. 35 (1997) 633–640. [23] I.A. Graham, Seed storage oil mobilization, Annu. Rev. Plant Biol. 59 (2008) 115–142. [24] M. Focke, E. Gieringer, S. Schwan, L. Jansch, S. Binder, H.-P. Brawn, Fatty acid biosynthesis in mitochondria of grasses: malonyl-coenzyme A is generated by a
[27] [28] [29]
[30]
[31] [32]
[33]
[34] [35]
[36]
[37] [38] [39] [40]
[41]
[42] [43]
[44]
[45]
[46]
[47] [48]
601
mitochondrial-localized acetyl-coenzyme A carboxylase, Plant Physiol. 133 (2003) 875–884. V. Gueguen, D. Macherel, J. Jaquinod, R. Douce, J. Bourguignon, Fatty acid and lipoic acid biosynthesis in higher plant mitochondria, J. Biol. Chem. 275 (2000) 5016–5025. J.E. Turner, K. Greville, E.C. Murphy, M.A. Hooks, Characterization of Arabidopsis fluoroacetate-resistant mutants reveals the principal mechanism of acetate activation for entry into the glyoxylate cycle, J. Biol. Chem. 280 (2005) 2780– 2787. R.H. Behal, M. Lin, S. Back, D.J. Oliver, Role of acetyl-CoA synthetase in the leaves of Arabidopsis thaliana, Arch. Biochem. Biophys. 402 (2002) 259–267. M. Tadege, I. Dupuis, C. Kuhlemeier, Ethanolic fermentation: new functions for an old pathway, Trends Plant Sci. 4 (1999) 320–325. S. Mellema, W. Eichenberger, A. Rawyler, M. Suter, M. Tadege, C. Kuhlemeier, The ethanolic fermentation pathway supports respiration and lipid biosynthesis in tobacco pollen, Plant J. 30 (2002) 329–336. N. Gass, T. Glagotskaia, S. Mellema, J. Stuurman, M. Barone, T. Mandel, U. Roessner-Tunali, C. Kuhlemeier, Pyruvate decarboxylase provides growing pollen tubes with a competitive advantage in petunia, Plant Cell 7 (2005) 2355–2368. J. Kreuzwieser, U. Scheerer, H. Rennenberg, Metabolic origin of acetaldehyde emitted by poplar (Populus tremula P. alba) trees, J. Exp. Bot. 50 (1999) 757–765. T. Karl, A.J. Curtis, T.N. Rosenstiel, R.K. Monson, R. Fal, Transient releases of acetaldehyde from tree leaves—products of a pyruvate overflow mechanism? Plant Cell Environ. 25 (2002) 1121–1131. J. Kreuzwieser, F.J.M. Harren, L.J.J. Lerhoven, I. Boamfa, S. Linten-Hekker, U. Scheerer, C. Huglin, H. Rennenberg, Acetaldehyde emission from the leaves of trees—correlation with physiological and environmental parameters, Physiol. Plant. 113 (2001) 41–49. D.J. Murphy, The biogenesis and function of lipid bodies in animals, plants, and microorganisms, Prog. Lipid Res. 40 (2001) 325–438. N.R. Lersten, A.R. Czlapinski, J.D. Curtis, R. Freckmann, H.T. Horner, Oil bodies in leaf mesophyll cells of angiosperms: overview and a selected survey, Am. J. Bot. 93 (2006) 1731–1739. M.L. Parker, G.J.P. Murphy, Oleosomes in flag leaves of wheat; their distribution, composition and fate during senescence and rust-infection, Planta 152 (1981) 36– 43. W. Lin, D.J. Oliver, Role of triacylglycerols in leaves, Plant Sci. 175 (2008) 233–237. G.J.P. Murphy, M.L. Parker, Lipid composition and carbon turnover of wheat leaf oleosomes, J. Exp. Bot. 35 (1984) 348–355. T.P. Durrett, C. Benning, J. Ohlrogge, Plant triacylglycerols as feedstocks for the production of biofuels, Plant J. 54 (2008) 593–607. A. Cernac, C. Benning, WRINKLED1 encodes an AP2/EREB domain protein involved in the control of storage compound biosynthesis in Arabidopsis, Plant J. 40 (2004) 575–585. J.M. Dyer, R.T. Mullen, Engineering plant oils as high-value industrial feedstocks for biorefining: the need for underpinning cell biology research, Physiol. Plant. 132 (2008) 11–22. B.J. Nikolau, M. Perera, L. Brachova, B. Shanks, Platform biochemicals for a biorenewable chemical industry, Plant J. 54 (2008) 536–545. A.B. Kroumova, G.J. Wagner, Different elongation pathways in the biosynthesis of acyl groups of trichome exudate sugar esters from various solanaceous plants, Planta 216 (2003) 1013–1021. A.B. Kroumova, Z. Xie, G.J. Wagner, A pathway for the biosynthesis of straight and branched, odd- and even-length, medium-chain fatty acids in plants, Proc. Natl. Acad. Sci. U.S.A. 91 (1994) 11437–11441. D.L. Rainwater, P.E. Kolattukudy, Fatty acid biosynthesis in Mycobacterium tuberculosis var. bovis Bacillus Calmette-Guerin. Purification and characterization of a novel fatty acid synthase, mycocerosic acid synthase, which elongates n-fatty acyl-CoA with methylmalonyl-CoA, J. Biol. Chem. 260 (1985) 616–623. K. Earley, R.J. Lawrence, O. Pontes, R. Reuther, A.J. Enciso, M. Silva, N. Neves, M. Gross, W. Viegas, C.S. Pikaard, Erasure of histone acetylation by Arabidopsis HDA6 mediates large-scale gene silencing in nucleolar dominance, Genes Dev. 20 (2006) 1283–1293. N. Geraldo, R. Abranches, Immunolocalization of histone modifications as a tool to visualize chromatin dynamics in plants, Microsc. Microanal. 14 (2008) 130–133. A.T. Martı´nez, J. Rencoret, G. Marques, A. Gutie´rrez, D. Ibarra, J. Jime´nez-Barbero, Jose´ C. del Rı´o, Monolignol acylation and lignin structure in some nonwoody plants: a 2D NMR study, Phytochemistry 69 (2008) 2831–2843.