Yarrowia lipolytica as a model for bio-oil production

Progress in Lipid Research 48 (2009) 375–387

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Progress in Lipid Research journal homepage: www.elsevier.com/locate/plipres

Review

Yarrowia lipolytica as a model for bio-oil production Athanasios Beopoulos a, Julien Cescut b, Ramdane Haddouche a, Jean-Louis Uribelarrea b, Carole Molina-Jouve b, Jean-Marc Nicaud a,* a b

Microbiology and Molecular Genetic Laboratory, CNRS UMR2585, INRA UMR1238, AgroParisTech, INRA centre de Versailles-Grignon BP 01, F-78850 Thiverval-Grignon, France Laboratoire d’Ingénierie des Systèmes Biologiques et des Procédés, CNRS UMR5504, INRA UMR792, INSA, 135 Avenue de Rangueil, F-31077 Toulouse, France

a r t i c l e

i n f o

Article history: Received 24 July 2009 Received in revised form 18 August 2009 Accepted 20 August 2009

Keywords: Lipid Yeast Yarrowia lipolytica Triacylglycerol Lipid particle b-oxidation Fermentation

a b s t r a c t The yeast Yarrowia lipolytica has developed very efficient mechanisms for breaking down and using hydrophobic substrates. It is considered an oleaginous yeast, based on its ability to accumulate large amounts of lipids. Completion of the sequencing of the Y. lipolytica genome and the existence of suitable tools for genetic manipulation have made it possible to use the metabolic function of this species for biotechnological applications. In this review, we describe the coordinated pathways of lipid metabolism, storage and mobilization in this yeast, focusing in particular on the roles and regulation of the various enzymes and organelles involved in these processes. The physiological responses of Y. lipolytica to hydrophobic substrates include surface-mediated and direct interfacial transport processes, the production of biosurfactants, hydrophobization of the cytoplasmic membrane and the formation of protrusions. We also discuss culture conditions, including the mode of culture control and the culture medium, as these conditions can be modified to enhance the accumulation of lipids with a specific composition and to identify links between various biological processes occurring in the cells of this yeast. Examples are presented demonstrating the potential use of Y. lipolytica in fatty-acid bioconversion, substrate valorization and single-cell oil production. Finally, this review also discusses recent progress in our understanding of the metabolic fate of hydrophobic compounds within the cell: their terminal oxidation, further degradation or accumulation in the form of intracellular lipid bodies. Ó 2009 Elsevier Ltd. All rights reserved.

Contents 1. 2.

3. 4.

5.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Lipid synthesis and accumulation factors in oleaginous microorganisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. Oleaginous yeasts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. Lipid accumulation pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3. The de novo lipid synthesis pathway and non-polar lipid synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4. Biochemistry and regulation of lipid accumulation potential in oleaginous yeast . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Factors affecting lipid accumulation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Modes of culture for ensuring high levels of lipid accumulation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1. Batch mode . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2. Continuous mode . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3. Fed-batch mode. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mastering lipid production . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1. The ex novo lipid accumulation pathway. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2. The b-oxidation degradation pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.3. Growth conditions and genetic modifications favoring lipid production . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

376 376 376 377 377 379 380 381 381 382 382 383 383 383 384

Abbreviations: AA, arachidonic acid; ACAT, acyl-CoA:cholesterol acyltransferase; ACC, acyl-CoA carboxylase; ACL, ATP citrate lyase; AMP, adenosine monophosphate; ATP, adenosine triphosphate; DAG, diacylglycerol; DHAP, dihydroxyacetone phosphate; FFA, free fatty acids; GLA, c-linolenic acid; G-3-P, glycerol-3-phosphate; HS, hydrophobic substrates; IMP, inosine 50 -monophosphate; LB, lipid body; MAG, monoacylglycerol; ME, malic enzyme; NADPH, nicotinamide adenine dinucleotide phosphate; PUFA, polyunsaturated fatty acids; SCO, single-cell oil; SE, steryl esters; TAG, triacylglycerols (triglycerides). * Corresponding author. E-mail address: [email protected] (J.-M. Nicaud). 0163-7827/$ - see front matter Ó 2009 Elsevier Ltd. All rights reserved. doi:10.1016/j.plipres.2009.08.005

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6. 7.

Potential applications. . . . . . . . . . . . . . . . . . . . Summary, conclusions and future directions . Acknowledgements . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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1. Introduction The yeast Yarrowia lipolytica is often found in environments rich in hydrophobic substrates, such as alkanes or lipids, and has developed sophisticated mechanisms for the efficient use of hydrophobic substrates (HS) as the sole carbon source [1,2]. One of the most striking features of this yeast is the presence in its genome of several multigene families involved in these metabolic pathways. The complexity and multiplicity of these genes enable Y. lipolytica to use and valorize a wide range of hydrophobic substrates (HS). Using these mechanisms, this yeast can accumulate lipids to levels exceeding 50% of cell dry weight [3]. Y. lipolytica may therefore be considered an oleaginous yeast. Lipid accumulation is probably enhanced by the many protrusions on the cell surface, facilitating HS uptake from the medium [4]. The internalized aliphatic chains are then broken down to meet needs for growth, or accumulate in an unchanged or modified form. These lipids form the storage lipid fraction, which consists mostly of triacylglycerols (triglycerides) (TAG) and steryl esters (SE). In addition to direct substrate assimilation from the medium, de novo TAG biosynthesis is another energy storage process providing fatty acids for membrane phospholipid formation. SE formation and mobilization provide the sterols required for membrane proliferation. Storage molecules accumulate in a specialized compartment of the cell known as the lipid body (LB). Yeast lipid bodies consist of a lipid core encased in a phospholipid monolayer, within which many proteins with diverse biochemical activities are embedded [5–7]. Several of these proteins metabolize lipids and the LB therefore probably plays a key role not only in lipid storage, but also in lipid biosynthesis, metabolism, degradation and substrate trafficking [6]. LB formation and function are tightly linked to the synthesis of TAG and SE. A recently identified lipid-binding protein in Y. lipolytica LB [8,9] has been implicated in lipid trafficking between the cytoplasm and LB, suggesting that free (non-esterified) fatty acids (FFA) probably accumulate in lipid bodies too [4,8,10,11]. A few models have been developed for the study of lipid metabolism. These models include Saccharomyces cerevisiae, which has long been used as a genetic model in studies that have greatly improved our understanding of lipid metabolism [12]. The enzymes involved in TAG biosynthesis, storage and degradation are very similar between species, and particularly between yeasts, but S.

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384 385 386 386

cerevisiae is not an oleaginous yeast and accumulates only moderate amounts of lipids (less than 15% of its biomass). Furthermore, unlike S. cerevisiae, which produces similar amounts of TAG and SE, Y. lipolytica stores mostly TAGs (>90%). This yeast is also unusual in accumulating significant quantities of FFA within the cell. The unique features of Y. lipolytica, together with the availability of efficient genetic tools for this species, have stimulated interest in the use of this yeast as a model for bio-oil production, with great potential for biotechnological applications. Several technologies, including various fermentation configurations, have been already used for single-cell oil (SCO) production by strains of Y. lipolytica grown on various agro-industrial by-products or waste [2,11]. The potential applications of these processes include the production of reserve lipids with particular structures (e.g. oils enriched in essential polyunsaturated fatty acids) and the production of nonspecific oils for use as renewable starting materials for the synthesis of bio-fuels. This review aims to provide insight into the routes of biosynthesis and degradation leading to the formation of oils and an overview of recent advances in the physiology and genetics of Y. lipolytica relating to the assimilation of HS.

2. Lipid synthesis and accumulation factors in oleaginous microorganisms 2.1. Oleaginous yeasts Few microorganisms are known to accumulate lipids to a significant level. Those species able to do so to a level corresponding to more than 20% of their biomass are described as oleaginous. Fewer than 30 of the 600 species of microorganisms investigated in one study were found to be oleaginous [13–15]. The best known oleaginous yeasts include genus of Candida, Cryptococcus, Rhodotorula, Rhizopus, Trichosporon and Yarrowia. On average, these yeasts accumulate lipids to a level corresponding to 40% of their biomass. However, in conditions of nutrient limitation, they may accumulate lipids to levels exceeding 70 % of their biomass (Table 1). Nevertheless, lipid content and profile differ between species. For instance, Cryptococcus curvatus and Cryptococcus albidus accumulate lipids to equivalent levels (58% and 65%, respectively), but their fatty acid profiles differ significantly. C. curvatus accumulates

Table 1 Lipid contents and fatty acid profiles of selected oleaginous yeasts [16]. Lipid contents are expressed, in terms of mass, as a fraction of dry cell mass (% ½glip g1 X , weight/dry weight).

a b

Species

% Lipid (glip gX1)

Major fatty acid residues (relative% w/w) C16:0

C16:1

C18:0

C18:1

C18:2

C18:3

Cryptococcus curvatus Cryptococcus albidus Candida sp. 107 Lipomyces starkeyi Rhodotorula glutinis Rhodotorula graminis Rhizopus arrhizus Trichosporon pullulans Yarrowia lipolytica

58 65 42 63 72 36 57 65 36

25 12 44 34 37 30 18 15 11

Ta 1 5 6 1 2 0 0 6

10 3 8 5 3 12 6 2 1

57 73 31 51 47 36 22 57 28

7 12 9 3 8 15 10 24 51

0b 0 1 0 0 4 12 1 1

T means trace. 0 means none detected.

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large amounts of palmitic acid, whereas oleic acid is the principal fatty acid accumulating in C. albidus. By contrast, in Rhodotorula species, lipid content diverges significantly (Rhodotorula glutinis and R. graminis accumulate lipids at levels corresponding to 72% and 36% of their biomass, respectively), but fatty acid composition remains similar. Y. lipolytica accumulates lipids to lower levels than some other oleaginous species, but it is the only yeast known to be able to accumulate such a high proportion of linoleic acid (more than 50% of the fatty acid residues present – Table 1). 2.2. Lipid accumulation pathways Lipids may accumulate via two different pathways: (1) de novo synthesis, involving the production, in defined conditions, of fatty acid precursors, such as acetyl and malonyl-CoA and their integration into the storage lipid biosynthetic pathway (the Kennedy pathway, see below) and (2) the ex novo accumulation pathway, involving the uptake of fatty acids, oils and triacylglycerols (TAG) from the culture medium and their accumulation in an unchanged or modified form within the cell. This pathway requires hydrolysis of the hydrophobic substrate (HS), transport of the released fatty acids within the cell, their re-assembly in the TAG and the steryl ester (SE) fractions and their accumulation within the LB. The main enzymes involved in these pathways are summarized in Table 2. 2.3. The de novo lipid synthesis pathway and non-polar lipid synthesis Non-polar lipid synthesis in yeasts requires a constant supply of coenzyme A (CoA)- activated FA for acylation of the glycerol backbone to synthesize TAG or the esterification of sterols to produce steryl esters (SE). The first two carbon atoms for de novo FA synthesis are provided by cytosolic acetyl-CoA, through citrate cleavage by ATP-citrate lyase (ACL) in the TCA cycle. The fatty acid chain

then grows through the addition of units of malonyl-CoA (endoplasmic reticulum) or acetyl-CoA (mitochondria). Mitochondrial acetyl-CoA for elongation is supplied by the breakdown of the lipids accumulated via the b-oxidation pathway. Malonyl-CoA is generated by acyl-CoA carboxylase (ACC) and is the principal source of carbon atoms for de novo FA synthesis. For each step in the elongation of the growing FA acyl chain, two molecules of NADPH are required. This NADPH is generated principally by malic enzyme (ME). These three enzymes (ACL, ACC and ME) are believed to play a crucial role in determining the potential for lipid accumulation and in regulating this process (see below). TAG synthesis generally follows the Kennedy pathway [17]. During the first step of TAG assembly, glycerol-3-phosphate (G-3-P) is acylated by G-3-P acyltranferase (SCT1) to generate lysophosphatidic acid (LPA), which is further acylated by lysophosphatidic acid acyltransferase (SLC1) to generate phosphatidic acid (PA). Upon dephosphorylation by phosphatidic acid phosphohydrolase (PAP), diacylglycerol (DAG) is released from PA. A gene encoding PAP, YALI0D27016g, has just been identified in the genome of Y. lipolytica (Table 1). This gene is 39% identical to the PAH1 gene of S. cerevisiae. Pah1p belongs to the PAP1 enzyme family, the members of which require Mg2+ [18] as a cofactor for catalytic activity. The activity of this enzyme in S. cerevisiae is regulated by lipids, nucleotides and phosphorylation [19,20]. In the final step of TAG synthesis, DAG is acylated in the sn-3 position, via an acyl-CoA-dependent or acyl-CoA-independent reaction (Fig. 1). The acyl-CoA-dependent reaction is catalyzed by three enzymes: Dga1p, Are1p and Are2p. DGA1 encodes an acylCoA:diacylglycerol acyltransferase (ACAT), but is unrelated to the DGAT1 subfamily encoding enzymes homologous to the acylCoA:cholesterol acyltransferases identified in plants and mammals [21]. Instead, it belongs to the same family as the mammalian DGAT2 gene. In Y. lipolytica, Dga1p seems to be the major enzyme

Table 2 Genes involved in fatty acid metabolism in Y. lipolytica and S. cerevisiae.a Gene

SC name

EC number

YL ortholog YL name

Function

GUT1 GPD1 GPD2 GUT2 PAP SCT1 GPT2 SLC1 DGA1 LRO1 TGL3 TGL4 TGL5 ARE1 ARE2sc ARE2yl TGL1 POX1 POX2 POX3 POX4 POX5 POX6 MFE1 POT1 ACL1 ACL2 MAE1 ACC1

YHL032c YDL022w YDL059w YIL155c YMR165c YBL011w YKR067w YDL052c YOR245c YNR008w YMR313c YKR089c YOR081c YCR048w YNR019w

EC EC EC EC EC EC EC EC EC EC EC EC EC EC EC EC EC EC

YALI0F00484g YALI0B02948g

Glycerol kinase Glycerol-3-phosphate dehydrogenase (NAD(+)) Glycerol-3-phosphate dehydrogenase (NAD(+)) Glycerol-3-phosphate dehydrogenase Phosphatidate phosphatase Glycerol-3-phosphate acyl transferase Glycerol-3-phosphate acyl transferase 1-acyl-sn-glycerol-3-phosphate acyltransferase Diacylglycerol acyltransferase Phospholipid:diacylglycerol acyltransferase Triacylglycerol lipase Triacylglycerol lipase Triacylglycerol lipase Acyl-CoA:sterol acyltransferase Acyl-CoA:sterol acyltransferase Acyl-CoA:sterol/Diacylglycerol acyltransferase Cholesterol esterase Acyl-coenzyme A oxidase Acyl-coenzyme A oxidase Acyl-coenzyme A oxidase Acyl-coenzyme A oxidase Acyl-coenzyme A oxidase Acyl-coenzyme A oxidase Multifunctional beta-oxidation protein Peroxisomal oxoacyl thiolase ATP-citrate lyase, subunit a ATP-citrate lyase, subunit b Malic enzyme Acetyl-CoA carboxylase

YKL140w YGL205w

YKR009c YIL160c NP NP YKL029c YNR016C

2.7.1.30 1.1.1.18 1.1.1.18 1.1.99.5 3.1.3.4 2.3.1.15 2.3.1.15 2.3.1.51 2.3.1.20 2.3.1.158 3.1.1.3 3.1.1.3 3.1.1.3 2.3.1.26 2.3.1.26 2.3.1.26 3.1.1.13 6.2.1.3

EC 4.2.1.74 EC 2.3.1.16

EC 1.1.1.38 EC 6.4.1.2

YALI0B13970g YALI0D27016g YALI0C00209g YALI0E18964g YALI0E32769g YALI0E16797g YALI0D17534g YALI0F10010g YALI0F06578g YALI0D07986g YALI0E32035g YALI0E32835g YALI0F10857g YALI0D24750g YALI0E27654g YALI0C23859g YALI0E06567g YALI0E15378g YALI0E18568g YALI0E34793g YALI0D24431g YALI0E18634g YALI0C11407g

Bioinformatic data were obtained from the Saccharomyces Genome Database (http://www.yeastgenome.org/) and the Genolevures database (http://cbi.labri.fr/Genolevures/). NP; not present in this microorganism. a Genes, corresponding S. cerevisiae gene name and EC number, Y. lipolytica ortholog (gene name), and corresponding function. EC number: enzyme commission number.

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Fatty acid synthesis Acetyl-CoA

Malonyl-CoA

Sterol

Glucose

DHAP

Elongation cycle

GUT2

GPD1 Glycerol

G-3-P

Acyl-CoA

GUT1

SCT1 LPA SLC1 PA

ARE1 ARE2 PL

SE

DAG

LRO1

Acyl-CoA DGA1, ARE1, ARE2

TAG

TGL1 Sterol

Neutral lipid synthesis

PAP

FFA Acyl-CoA

TGL3, TGL4 Glycerol

Mobilization Degradation

POX 1-6

β-oxydation

MFE1 THIO1

Acetyl-CoA Fig. 1. Overview of the various pathways involved in fatty acid synthesis and in the storage and degradation of non-polar lipids. Synthesis of non-polar lipids (SE and TAG) required sterol, acyl-CoA and glycerol-3-phosphate (G-3-P). Synthesis of fatty acids (Acyl-CoA) is catalyzed by the fatty acid synthase from the basic blocks acetyl-CoA and malonyl-CoA through elongation cycles. G-3-P can be produced either from glycerol by the glycerol kinase encoded by GUT1 or from glucose via conversion of dihydroxyacetone DHAP by the glycerol-3-phosphate dehydrogenase encoded by GPD1 gene. G-3-P can be oxidized to DHAP by the glycerol-3-phosphate dehydrogenase encoded by GUT2 gene. The synthesis of SE is catalyzed by SE synthases encoded by ARE1 and ARE2. For the synthesis of TAG, three acyls are added to the G-3-P backbone through enzymatic steps: first, an acyl is added at the sn-1 position of G-3-P by a G-3-P acyltransferase to produce LPA (SCT1 gene), then a second acyl is added at the sn-2 position by a 1-acyl G-3-P acyltransferase (SLC1 gene) to produce phosphatidic acid (PA), which is then dephosphorylated by a phosphatidate phosphatase (PAP) yielding DAG. Finally, the third acyl can be added at sn-3 position either by the acetyl-CoA-dependent pathway (directly from Acyl-CoA) by acyl-CoA:diacylglycerol acyltransferase (DGA1) and by acyl-CoA:diacylglycerol acyltransferase/acyl-CoA:cholesterol acyltransferase (ARE1, ARE2) or by the acetyl-CoA-independent pathway (from a glycerophospholipid, PL) by the phospholipid:diacylglycerol acyltransferase (LRO1). Homologs to S. cerevisiae TAG lipases TGL1, TGL3 and TGL4 involved in SE and TAG mobilization have been identified (Table 2). The FFA can then be degraded in the b-oxidation pathway which involved acyl-CoA oxidase (POX), multifonctional beta-oxidation protein (MFE1) and the thiolase (THOI1). In Y. lipolytica, six genes (POX1–POX6) coding for acyl-CoA oxidases are involved in the second step of the b-oxidation.

involved in TAG synthesis, accounting for 45% of DAG acylation (Beopoulos et al., manuscript in preparation). In vivo essays have shown that this enzyme preferentially makes use of oleic and palmitic acid as precursors for CoA-mediated acylation. The contribution of Dga1p in Y. lipolytica seems to increase during the growth cycle, being greatest in the stationary phase. The tagging of Dga1p with the fluorescent protein EYFP showed Dga1p to be located mostly at the surface of LB (Beopoulos et al., manuscript in preparation). Two steryl ester synthases, encoded by the ARE1 and ARE2 genes, have been identified as orthologs of the genes found in S. cerevisiae, and have been shown to contribute to DAG acylation by acting as acyl transferases in an acyl-CoA-dependent mechanism [22,12]. YlAre1p has an amino-acid sequence about 30% identical to that of its two orthologs in S. cerevisiae, whereas YlAre2p has a lower level of sequence identity to Ylare1p (17%) and is more similar to the diacylglycerol O-acyl transferase (DGAT) from the plant Perilla frutescens (22%), human DGAT1 (28%) and DGAT1 from the plant A. thaliana (25%) [23]. All these enzymes belong to the acyl-CoA:cholesterol acyltransferase (ACAT) family, but their genes display higher levels of sequence identity to the members of the mammalian DGAT1 gene family [24]. Despite the differences between their sequences and that of Dga1p, Are proteins display DGAT activity and are the major enzymes required for TAG synthesis during the exponential growth phase of Y. lipolytica. The ob-

served DGAT activity may result from the evolution of ARE genes from an ancestral DGAT1 gene duplication in yeast. In vivo studies have suggested that Are1p prefers saturated acyl-CoA substrates, whereas Are2p prefers unsaturated acyl-CoAs and, more specifically, incorporates oleic acid into TAG (Beopoulos et al., manuscript in preparation). In yeasts, the acyl-CoA-independent reaction is carried out by lro1p, a protein with a sequence 27% identical to that of the human lecithin cholesterol acyl-transferase (LCAT) [25]. This enzyme has both phospholipase and acyltransferase functions. Unlike its human ortholog, yeast Lro1p cannot synthesize sterols and functions as a phospholipid: diacyl glycerol acyl transferase (PDAT) [26]. The acylation of DAG is an esterification reaction involving the sn-2 group of glycerophospholipids, preferentially PtdCho and PtdEtn. The deletion of LRO1 in Y. lipolytica results in a 35% decrease in TAG levels in vivo, with Lro1p making a minor contribution during the exponential phase and a much greater contribution during the stationary phase (Beopoulos et al., manuscript in preparation). SE formation involves the reaction of a fatty acid molecule with the hydroxyl group of sterols [27]. The SE fraction accounts for 50% of storage lipids in S. cerevisiae [28], whereas only small quantities of SE are synthesized (2–5%) in Y. lipolytica [29]. In S. cerevisiae, the acyl-CoA:sterol acyltransferases Are1p and Are2p are the only SE synthases involved in sterol esterification, as a double mutant lacking both ARE genes cannot synthesize SE [30]. Similar results have

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been obtained for Y. lipolytica (Beopoulos et al., manuscript in preparation). In vivo measurements of SE accumulation in Y. lipolytica have suggested that the Are proteins act in synergy and that their relative contribution is probably greater in the exponential growth phase. Fatty acid (Acyl-CoA) synthesis is catalyzed by the fatty acid synthase complex, with the basic acetyl-CoA and malonyl-CoA building blocks. The acyl-CoA may be stored as sterol ester (SE) or triacylglycerol (TAG). SE synthesis is catalyzed by an SE synthase homologous to the human acyl-CoA:cholesterol acyltransferase (ACAT) and SE mobilization is catalyzed by SE hydrolases, which release sterol and FFA. TAG synthesis requires acyl-CoA and glycerol-3-phosphate (G-3-P). G-3-P may be produced from glycerol or from dihydroxyacetone (DHAP). GUT1 encodes a glycerol kinase that converts glycerol to G-3-P in the cytosol. The G-3-P produced is then oxidized to DHAP by the glycerol-3-phosphate dehydrogenase encoded by the GUT2 gene and this DHAP may enter glycolysis or gluconeogenesis. G-3-P may also be used as a skeleton for triacylglycerol synthesis. Three acyl groups are added to the G-3P backbone to generate TAG, and this process involves four enzyme-catalyzed steps: (1) an acyl group is added at the sn-1 position of G-3-P by a G-3-P acyltransferase to produce LPA; (2) a second acyl group is added at the sn-2 position by a 1-acyl G3-P acyltransferase (AGAT) to produce PA; (3) the PA is then dephosphorylated by a phosphatidate phosphatase (PAP), yielding DAG and (4) the third acyl group is added at the sn-3 position via the acyl-CoA-dependent pathway or the acyl-CoA-independent pathway. In the acyl CoA-independent pathway, this third acyl group is supplied by a glycerophospholipid, whereas, in the acylCoA-dependent pathway, it is supplied by acyl-CoA. TAG can be mobilized by conversion to FFA and DAG upon hydrolysis by TAG lipase. The FFA generated may then be degraded by the b-oxidation pathway which involved POX, MFE and THIO genes. This pathway involves four enzyme-catalyzed steps. In Y. lipolytica, six genes (POX1 to POX6) encoding acyl-CoA oxidases involved in the second step of b-oxidation have been identified [31,2]. 2.4. Biochemistry and regulation of lipid accumulation potential in oleaginous yeast Oleaginous microorganisms begin to accumulate lipids when an element in the medium becomes limiting and the carbon source (such as glucose) is present in excess. Many elements can induce lipid accumulation. Nitrogen limitation is generally used in lipid accumulation studies in microorganisms. Nitrogen limitation is the easiest condition to control and is generally the most efficient type of limitation for inducing lipid accumulation. During the growth phase, the carbon flux is distributed between the four macromolecular pools (carbohydrate, lipid, nucleic acid, protein). Nitrogen is essential for the protein and nucleic acid synthesis required for cellular proliferation. This process is therefore slowed by nitrogen limitation. However, in conditions of nitrogen limitation, the catalytic growth rate slows down rapidly, whereas the rate of carbon assimilation slows more gradually. This results in the preferential channeling of carbon flux toward lipid synthesis, leading to an accumulation of triacylglycerols within discrete lipid bodies in the cells. If non-oleaginous microorganisms are placed in the same nutrient-limiting medium, further cell proliferation tends to cease, with carbon flux into the cell maintained but, in this case, the carbon is converted into various polysaccharides, including glycogen and various glucans and mannans. During the transition between the growth phase (growth with the production of catalytic biomass) and the lipid accumulation phase (decrease in growth rate due to nutrient limitation and the diversion of excess carbon to lipid production), some pathways are repressed (nucleic acid and protein synthesis), whereas others

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are induced (fatty acid and triacylglycerol synthesis). This transition is induced by the establishment of nitrogen limitation (see below). In addition, during the accumulation phase, precursors (acetyl-CoA, malonyl-CoA and glycerol) and energy (ATP, NADPH) are required for lipid synthesis. We will describe here the role of the key enzymes involved in regulating lipid accumulation potential. AMP deaminase is activated by the exhaustion of nitrogen in the medium during the growth of an oleaginous microorganism. AMP deaminase catalyzes the following reaction [32]:

AMP ! IMP þ NHþ4 The activation of AMP deaminase decreases mitochondrial AMP concentration and increases cellular ammonium concentration. The decrease in AMP concentration inhibits isocitrate dehydrogenase, blocking the citric acid cycle at the isocitrate level. Aconitase mediates the accumulation of citrate in mitrochondria, with exit from the mitochondria mediated by the citrate/malate cycle [32,33]. This reaction provides large amounts of acetyl-CoA for fatty acid synthesis. Acetyl-CoA is provided by the cleavage of citrate coming from the mitochondria by ATP-citrate lyase (ACL) in the cytosol. ACL cleaves the citrate to give oxaloacetate and acetyl-CoA.

Citrate þ HS-CoA þ ATP ! acetyl-CoA þ oxaloacetate þ ADP þ Pi This enzyme is absent from non-oleaginous yeasts, such as S. cerevisiae, but has been shown to be present in Y. lipolytica. Whereas the human ACL consists of a single protein encoded by a single gene, the ACL enzymes of both Y. lipolytica and Neurospora crassa consist of two subunits, Aclap and Aclbp, encoded by ACL1 (YALI0E34793g) and ACL2 (YALI0D24431g), respectively (Table 2). This enzyme requires an ammonium ion for activation and is dependent on adenosine mono- and diphosphate [16,34]. However, ammonium ions are scarce in the absence of nitrogen, due to the induction of AMP deaminase [32,33]. In addition to acetyl-CoA, fatty acid synthesis requires a continuous supply of malonyl-CoA and NADPH. Malonyl-CoA can also be generated from acetyl-CoA, in a reaction catalyzed by acetyl-CoA carboxylase (Acc1p) [35].

Acetyl-CoA þ HCO3 þ ATP ! malonyl-CoA þ ADP þ P i In mammalian cells, this enzyme is activated in the presence of tricarboxylic acid intermediates, such as citrate [36]. In yeast, however, Acc1p undergoes allosteric activation as a function of citrate concentration [37]. In Y. lipolytica, this enzyme is encoded by the ACC1 gene, and is known as YALI0C11407 g (Table 2). NADPH is required for the function of the fatty acid synthase (FAS). It is thought that NADPH concentration is controlled by the activity of malic enzyme (ME). This enzyme catalyzes the following reaction:

Malate þ NADPþ ! pyruvate þ NADPH The first evidence of ME involvement in lipid accumulation was provided by the inhibition of this enzyme by sesamol in Mucor circinelloides, resulting in a decrease in lipid accumulation from 25% to 2% of cell biomass [38]. Ratledge et al. subsequently demonstrated a direct correlation between decreasing ME activity during the lipid accumulation phase and the extent of lipid accumulation [39]. ME overproduction in M. circinelloides, through the expression of the gene under the control of the strong constitutive promoter of the glyceraldehyde-3-phosphate dehydrogenase gene (gpd1), was recently shown to increase lipid accumulation by a factor of 2.5 [40]. In Y. lipolytica, this enzyme is encoded by the MAE1 gene

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(YALI0E18634 g – Table 2). However, preliminary results for ME overproduction in Y. lipolytica indicate that NADPH concentration is not limiting for lipid accumulation in this yeast.

3. Factors affecting lipid accumulation Lipid accumulation depends primarily on microorganism physiology, nutrient limitation and environmental conditions, such as temperature and pH. It is also affected by the production of secondary metabolites, such as citrate and ethanol. The transition from catalytic growth to lipid accumulation generally occurs when excess carbon in the medium is associated with a nutrient limitation affecting biomass production. Y. lipolytica and S. cerevisiae have followed different courses of physiological evolution. S. cerevisiae catabolizes glucose efficiently, glycerol to a lesser extent, and fatty acids poorly. S. cerevisiae does not use fatty acids very efficiently. Indeed, the doubling time of cell populations on this substrate is about 4 h. It is widely accepted that this poor lipid utilization efficiency results from a limited capacity for b-oxidation (breakdown of FA to generate acetyl-CoA). S. cerevisiae cannot use oils (mixture of TAG and FA) due to its inability to secrete lipases. For S. cerevisiae (Fig. 2A), the presence of excess carbon (more glucose taken up than required for biomass production) induces a metabolic shift from oxidative to oxido-reductive metabolism with ethanol production. In aerobic batch culture, S. cerevisiae produces ethanol, with a glucose-to-ethanol conversion yield of 1 0:548 Cmoletoh Cmolglc , a maximum specific growth rate of 0.43 h1 and an average ethanol volumic productivity of 3.3 g l1 h1 [41]. Y. lipolytica grows as well on glucose (lmax = 0.26 h1, [41,42]) as on oleic acid (lmax = 0.33 h1 [43]). In conditions of nutrient limitation in the presence of excess carbon, Y. lipolytica produces large amounts of TCA-cycle intermediates, such as citric (CA), isocitric acid (ICA), 2-ketoglutaric acid and pyruvic acid (for review see [2]). It also converts excess carbon into TAG. By contrast to S. cerevisiae, Y. lipolytica (Fig. 2B) is a strict aerobic yeast unable to produce ethanol. Depending on C/N ratio, different governing metabolisms can be observed: pure growth, organic acid production or conversion of excess carbon into lipids (triacylglycerols and sterol esters). By monitoring growth, carbon excess in the

S. cerevisiae

A

anabolism of Y. lipolytica can be either oriented towards organic acids production or lipids production (TAG, SE). Y. lipolytica is an oleaginous yeast able to accumulate lipids to levels exceeding 50% of cell dry weight. Lipid accumulation, in terms of lipid profile, amount, productivity and conversion yield, is influenced by various operating conditions, such as the nature of nutrient limitation, pH, aeration and temperature conditions. Published values for lipid accumulation by the yeasts Candida sp. 107, R. glutinis and Y. lipolytica are presented in Table 3 as a function of the nature of nutrient limitation: nitrogen, magnesium, zinc, iron or phosphorus. However, nitrogen limitation is most commonly used to induce lipid accumulation and gives the best conversion yield with glucose, reaching 0:22 glip g1 glu [44,45]. We will deal here specifically with nitrogen limitation, which remains the most efficient form of nutrient limitation for the induction of lipid accumulation. The culture strategy involves two phases, as a function of yeast metabolism. The first is a growth phase [48,49]. The growth of the yeast is then slowed by nitrogen limitation and the lipid accumulation phase begins. The global conversion yield of glucose into lipids in batch culture depends on the

Table 3 Lipid content and lipid yield as a function of the nature of nutrient limitation for Candida sp. 107 (a) Rhodotorula glutinis (b) and Yarrowia lipolytica (c). Limitation

Yeast Culture mode

% Lipid ðglip g1 X Þ

Maximum yield ðglip g1 glu Þ

References

Nitrogen Nitrogen Nitrogen Nitrogen Nitrogen Carbon Phosphorus Phosphorus and nitrogen Magnesium Zinc Iron

a c c c b a a a

Batch Continuous Fed-batch Batch Fed-batch Batch Batch Batch

37% 28% 38.6% 11% 72% 14% 31% 35%

0.22 0.11 0.22 0.017 0.255 0.07 0.15 0.052

[37] [46] [42] [47] [41] [37] [37] [37]

a b b

Batch Batch Batch

32.3% 34% 45%

0.10 0.22 0.12

[37] [44] [44]

Y. lipolytica

B

Glucose

Glucose

Fatty Acid transport and β -oxydation

No limitation

Alkane Fatty acid

Fatty acid

Oil Glycerol glycerol

Ethanol

Ethanol

Metabolite

Lipid

Respiration capacity

Biomass Metabolite

Biomass

Fig. 2. Schematic comparison of S. cerevisiae and Y. lipolytica metabolisms. S. cerevisiae catabolizes preferentially sugars (glucose, saccharose) rather than fatty acids. S. cerevisiae cannot catabolize oils, because it does not secrete lipase. In the presence of excess sugar, S. cerevisiae produces ethanol. Y. lipolytica catabolizes alkanes, glucose, fatty acids and glycerol. This yeast secretes lipases and is therefore able to grow on oils. In the presence of excess carbon, Y. lipolytica either produces metabolites or stores lipids.

-1

0.45 0.4 0.35 0.3 0.25 0.2 100

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temperature. For example, R. glutinis produces 0:44 gAGI g1 AG unsaturated lipids at 15 °C and 0:27 gAGI g1 AG at 30 °C [44]. This modulation of the lipid profile probably results from an increase in desaturase stability at low temperatures, with no such increase in stability observed for the other enzymes. However, low temperatures do not favor lipid production, because they also lead to large decreases in cellular activity and metabolism. There are few alternatives to genetic modification for the modulation of fatty acid profile. One of these alternatives is the use of growth inhibitors, such as cerulenin [58], or natural antimicrobial compounds, such as Teucrium polium extracts [46].

0.5 [Cmollip.Cmolglc ]

conversion of glucose to lipids yield

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150

200 250 300 -1 initial ratio C/N [Cmolglc.molN ]

350

400

Fig. 3. Changes in conversion yield for the conversion of glucose into lipids plotted against C/N ratio for batches of R. glutinis [44].

duration of the growth phase and the transition to the accumulation phase. The duration of the growth phase depends on the C/N ratio. The total substrate-to-lipid conversion yield therefore depends on the initial C/N ratio of the batch culture. Glucose-to-lipid 1 conversion yield increases from 0.25 to 0.40 ðCmollip Cmolglu Þ as C/ 1 N ratio increases from 150 to 350 ðCmolglu molN Þ for the oleaginous yeast R. glutinis (Fig. 3) [44]. However, a C/N ratio exceeding 350 gC g1 N (data not shown) creates a severe nitrogen deficiency, leading to a rapid decrease in cell viability before the cells are able to enter the lipid accumulation stage. The immediate precursor of cellular lipid accumulation in oleaginous microorganisms is citric acid [15,50–52]. In Y. lipolytica, if the initial C/N molar ratio is high [80–120 Cmol Nmol1], cell growth may be followed by significant citric acid production, with low levels of lipid accumulation within cells. Papanikolaou et al. assumed that ATP-citrate lyase was inactive in the presence of excess glucose, resulting in low levels of lipid accumulation [53]. However, if a cosubstrate was used, optimal C/N ratios of 35 Cmol Nmol1 for lipid production from glucose and 180 Cmol Nmol1 for lipid production from glycerol and glucose were obtained for batch cultures [54,55]. In continuous culture conditions, Aggelis and Komaitis used a medium with a C/N ratio of 66 Cmol Nmol1 [46]. In general, the maximum C/N ratio suitable for lipid accumulaY 1 ½CmolX Cmolsub  , where q is tion can be estimated from the ratio X=S q 1 the nitrogen content of biomass [Nmol Cmol ], YX/S is the ratio of theoretical biomass production to substrate, expressed as Cmol of biomass/Cmol of substrate in conditions of carbon limitation in the absence of by-product production [56]. If carbon is not limiting (present in large excess), the uptake of carbon is limited only by the substrate transport system of the cell. In this case, limiting concentrations of nitrogen in the medium lead to the induction of lipid accumulation. The critical nitrogen concentration for lipid induction in Y. lipolytica has been found to be about 103 mol l1[42]. It is important for nitrogen concentration to exceed this threshold value to prevent the production of secondary metabolites (citric acid) that will otherwise affect lipid accumulation. Conversely, if the extracellular carbon supply is exhausted, stored lipids may be mobilized. Thus, lipid accumulation is always dependent on the influx of carbon substrates. This complex regulation makes it difficult to achieve high rates of lipid accumulation in batch culture. In such conditions, lipid accumulation is always followed by citric acid production [47]. These findings highlight the need to manage both carbon and nitrogen flows to optimize lipid accumulation and reduce citric acid production (see below). Lipid profile can be modified by adjusting the temperature [57]. Fatty acid composition is dependent on culture temperature, because the degree of saturation generally decreases with decreasing

4. Modes of culture for ensuring high levels of lipid accumulation Bioprocesses for lipid production may be designed on the basis of our knowledge of Y. lipolytica metabolism, taking into account the ability of this yeast to produce large amounts of intermediates and to accumulate large amounts of lipids and to break them down by b-oxidation, even on a glucose substrate [42]. Four metabolic states can be defined as a function of differences in C/N flux ratio for a constant nitrogen flux (Fig. 4). The first corresponds to a carbon flux lower than that required for growth. In this case, if the cells have storage lipids, they make up the carbon deficit by mobilizing these storage lipids (Fig. 4, state a). The second state corresponds to the maximum growth rate obtained with optimal carbon influx from the medium, at a defined nitrogen flux rate. This state results in maximal biomass production (Fig. 4, state b). The third state corresponds to an influx of excess carbon from the medium, which has a high C/N ratio, resulting in a decrease in biomass production and high levels of lipid production (Fig. 4, state c). The fourth state results from a further increase in C/N ratio (Fig. 4, state d), leading to the repression of lipid accumulation in favor of secondary metabolite production. If the desired end product of the process is lipid, then the process must be designed so as to ensure the maximal conversion of the carbon taken up into lipids, by minimizing by-product (citric acid) production and maximizing lipid synthesis, holding the cells in state c. Three different modes of culture are commonly used: batch, fed-batch and continuous mode. Most of the processes described in previous publications relate to batch mode [45,47,59]. 4.1. Batch mode In batch cultures, minerals and carbon substrates are initially mixed in the bioreactor, with a high initial C/N ratio to boost lipid accumulation. As nitrogen is actively consumed right from the start of culture, the rC/rN ratio (residual carbon to residual nitrogen ratio) continually increases, tending to infinity. In this mode, growth remains exponential whilst nitrogen is not limiting (Fig. 4, state b). Granger showed that, in R. glutinis grown at 30 °C with glucose, nitrogen limitation led to a decrease in the rate of substrate consumption by a factor of four and an increase in lipid production rate by a factor of two to three [44]. Microbial metabolism then shifts into phase c (Fig. 4). Nevertheless, citric acid production is induced as a function of rC/rN ratio, resulting in a shift of microbial metabolism into phase d (Fig. 4). Citric acid production decreases the total conversion yield for the production of lipids from carbon substrate. Thus, this conversion yield depends mostly on the ratio of biomass constituted during the growth phase to lipids accumulated during the accumulation phase in batch culture. Control of the ratio of carbon consumption to nitrogen consumption is therefore essential to prevent citric acid secretion, hence the importance of monitoring rC/rN ratio in continuous and fed-batch cultures.

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Carbon flow/growth carbon flow need

382

250% 200%

a

b

c

d

150% 100% 50% 0%

carbon substrate

nitrogen

catalytic biomass

lipids

citric acid

Fig. 4. Yeast activity as a function of carbon flow rate for a fixed nitrogen flow rate. State a: pure growth and mobilization of stored lipids; state b: pure growth; state c: growth with lipid accumulation; state d: growth, lipid accumulation and citric acid production. The size of arrow is proportional to flow. The carbon substrate and nitrogen arrows correspond to specific influx into cells. Catalytic biomass, lipids and citric acid arrows correspond to specific production rates (consumption for lipids, state a).

4.2. Continuous mode Lipid production in continuous culture has been modeled by Ykema et al. [56]. In a continuous culture, the C/N ratio in the culture medium and rC/rN are constant for a given dilution rate. For low dilution rates, with intermediate C/N ratios promoting lipid accumulation ð40 gC g1 N Þ, the lipid and biomass concentrations obtained are higher than those obtained for higher dilutions. Indeed, at similar rC/rN ratios, low specific growth rates promote lipid accumulation. The optimization of the process therefore involves determining the optimal dilution rate with an optimal intermediate C/N ratio (see Fig. 5).

as the substrate goes through three phases: a pure growth phase (i), a transition phase (ii) and a lipid accumulation phase (iii). During the growth phase, yeast metabolism results in the balanced distribution of carbon between the four main macromolecular pools (carbohydrate, lipid, nucleic acid, protein), with catalytic biomass production (biomass without the accumulation of a macromolecular compound). This phase corresponds to state b in Fig. 4.

4.3. Fed-batch mode The production of lipid by yeast may ensure that lipids are efficiently and reproducibly produced [60]. If lipid production is to be controlled, regulation of the environmental variables is require to maximize the stability of the metabolic state. This stability can be achieved through the precise control of nutrient flow rate. Greater accuracy in the control of nutrient flow rate is associated with greater control of metabolic state and more optimal production, which is the case for protein production processes [61] (for review see [62]). In fed-batch culture, nitrogen and carbon flows are monitored to control the specific growth rate and the rC/rN ratio. As shown in Fig. 6, a fed-batch culture of Y. lipolytica at 28 °C with glucose

Fig. 5. Volumetric productivity of Y. lipolytica biomass P(X) and lipid P(lip) plotted against dilution rate on industrial glycerol. Cells were grown in single-stage continuous cultures, with a fixed C/N ratio at pH 6 ± 0.05; T = 28 °C and aeration rate = 1.8 VVM. [55].

383

glucose, biomass, lipid, citric acid concentration [arbitrary unit]

A. Beopoulos et al. / Progress in Lipid Research 48 (2009) 375–387

transition

lipid accumulation nitrogen concentration [arbitrary unit]

growth

305

20

300

18

295

16 14

290

12

285

10

280

8

275

6

270

4

265

2

260

0

255

-2

0

10

20

30

40

50

60

70

80

time [h] glucose [g]

NH4 [g]

lipids [g]

biomass [g]

citric acid [g]

Fig. 6. Modeling and prediction for a fed-batch culture of Y. lipolytica. The first part of the growth (from 0 h to 27 h) corresponds to a pure growth phase, with a C/N flux ratio equal to catalytic biomass production requirements. The second phase (transition from 27 to 40 h) corresponds to the establishment of nutrient limitation (nitrogen), followed by the establishment of lipid accumulation. The last step (from 40 to 70 h) corresponds to the lipid accumulation phase, during which nutrient limitation is 1 controlled by optimizing the C/N ratio [around 20 Cmol molN to favor lipid accumulation.

The transition phase corresponds to the establishment of nitrogen limitation, with excess carbon leading to the accumulation of lipids. This phase corresponds to the transition between state b and state c. Lipid accumulation phase is the most extensive, corresponding to the establishment lipid production under constant nitrogen limitation conditions, with a C/N ratio of about 20 Cmol Nmol1, preventing citric acid production. 5. Mastering lipid production Lipid accumulation in oleaginous species is upregulated in culture media containing fatty acids or oils. Y. lipolytica has developed sophisticated mechanisms for taking up and assimilating hydrophobic substrates. However, this process is counterbalanced by beta oxidation, which mobilizes the accumulated lipids. Nevertheless, with a suitable culture medium and appropriate genetic modifications, high levels of lipid accumulation can be achieved, exceeding 60% of cell dry weight in some cases, with modification of the profile of accumulated fatty acids. 5.1. The ex novo lipid accumulation pathway Y. lipolytica has evolved elaborate strategies and adaptation mechanisms for the efficient use of hydrophobic substrates, such as n-alkanes, fatty acids and triacylglycerols. It has adapted to the use of these substrates by evolving genes encoding surfactants for their solubilization (liposan), by modifying its cell surface to facilitate adhesion of hydrophobic droplets (cell surface protrusions), thereby maximizing cell-substrate contact and by developing complex transport mechanisms for the incorporation of these compounds into the cell (for reviews see [2], [63], [64]). This adaptation of Y. lipolytica may have involved genome amplification and successive evolution of the genes required for the utilization of diverse hydrophobic substrates (with respect to the chain lengths of alkanes and fatty acids). Indeed, Y. lipolytica has at least 13 families of genes involved in hydrophobic substrate utilization, reflecting the vast expansion of its genome with respect to those of other ascomycetous yeasts [65,66]. For example, the lipase gene family of this yeast has 16 members and the acyl-CoA oxidase family has six members.

A fine example of the complex mechanisms developed by Y. lipolytica is in the hydrolysis and incorporation of oil substrates. The yeast secretes an extracellular lipase called lip2p, encoded by the LIP2 gene. This gene encodes a precursor pre-pro-mature protein with a Lys-Arg (KR) cleavage site [67]. It simultaneously produces other intracellular lipases, such as Lip7p and Lip8p, which may be released into the medium, depending on the substrate. These lipases have different chain-length specificities, with Lip2, Lip7 and Lip8 displaying maximal activity with oleate (C18), caproate (C6) and caprate (C10), respectively [68,69]. The released fatty acids must then be transported into the cell. Kholwein et al. carried out the first study of FA transport in Y. lipolytica. They showed that an energy-free transporter was required below a threshold of 10 lM, whereas at higher concentrations, lauric or oleic acid diffused freely [70]. This model also suggested that Y. lipolytica had two different chain length-selective transporters. Papanikolaou and Aggelis analyzed fatty acid uptake in Y. lipolytica during growth on lipid-containing media with different substrate compositions [55]. Various mixtures of stearin and hydrolyzed rapeseed oil were used for growth, and fatty acid uptake and profiles were analyzed. Regardless of the external fatty acid composition, all fatty acids displayed similar incorporation constants for uptake (0.023 h1), with the exception of C16:0 and C18:0, which had lower constants (0.01 and 0.013 h1, respectively) (Fig. 4). However, if Y. lipolytica is grown on saturated fatty acids alone, the incorporation constants for C16:0 and C18:0 were significantly higher (0.018 h1) [71] (see Fig. 7). 5.2. The b-oxidation degradation pathway In Y. lipolytica, FA are broken down in peroxisomes, via the fourstep b-oxidation pathway. Six different acyl-CoA oxidases (Aox1-6, encoded by the POX1 to POX6 genes) catalyze the first and rate-limiting step of b-oxidation. These acyl-CoA oxidases (Aox) have different activities and substrate specificities, as shown by gene disruption in Y. lipolytica [31] and by expression and purification in E. coli. Aox2p is very active and highly specific for long-chain fatty acids, whereas Aox3p preferentially acts on short-chain fatty acids. Thevenieau et al. [63] recently showed that Aox1p and Aox6p are involved in breaking down dicarboxylic acids (DCA).

A. Beopoulos et al. / Progress in Lipid Research 48 (2009) 375–387

Incorporation constant (h-1)

384

0.030 C12:0

0.025

C18:2

C14:0

C18:3

C18:1

0.020 0.015

C16:0 C18:0

0.010 0.005 0.000

Fatty acid Fig. 7. Fatty acid uptake in Y. lipolytica. Incorporation constant (k, h1) of fatty acids during the growth of Y. lipolytica on lipid-containing medium (mixture of hydrolyzed rapeseed oil and stearin). (Adapted from [71]).

Gene disruption analysis have also shown that the mechanism of peroxisome entry depends on POX genotype and that these enzymes have no peroxisome targeting sequence (PTS). By contrast, the acyl-CoA oxidase (Aox) complex of Y. lipolytica has been shown to be assembled in the cytosol before its import into peroxisomes as a heteropentameric, cofactor-containing complex, with Aox2p and Aox3p playing a key role in this process [72]. Aox proteins have also been shown to play a major role in peroxisomal division. During peroxisome maturation, a membrane-bound pool of Aox interacts with a membrane-associated peroxin, Pex16p. Pex16p downregulates membrane fission, thus preventing the excessive proliferation of immature peroxisomal vesicles. The Dpox4 and Dpox5 mutants form giant peroxisomes [73]. The second and third steps of b-oxidation are catalyzed by the multifunctional enzyme encoded by the MFE gene, which displays hydratase and dehydrogenase activities (Fig. 1). The fourth step is catalyzed by the 3-ketoacyl-CoA-thiolase encoded by the POT1 gene. A decane-inducible peroxisomal acetoacetyl-CoA thiolase (encoded by PAT1) has recently been identified and, together with the thiolase encoded by POT1, this enzyme is thought to catalyze the last step of b-oxidation. Changes in b-oxidation flux or the complete abolition of this process might increase lipid accumulation (see below). 5.3. Growth conditions and genetic modifications favoring lipid production The ability of Y. lipolytica to utilize and degrade HS has led to its use in bioremediation processes and in fermentation techniques for the production of intermediate metabolites, enzymes and added-value oils. The capacity of this yeast to break down HS has made it possible to decrease chemical oxygen demand (COD) significantly in oil mill wastewater containing fats, sugars, phosphate, phenols and metals [74]. Similar methods have been used for selective medium degradation [75] and fat separating processes in wastewater purification. The production of intermediate metabolites was highlighted by Papanikolaou et al. [53], who used Y. lipolytica to produce citric acid from raw glycerol, the major byproduct of biodiesel production units. A high C/N ratio, combined with a buffered pH, resulted in rates of citric acid production of 35 g.l1, and low rates of lipid accumulation, probably due to the downregulation of ATP-citrate lyase in the experimental conditions used. Similar results were obtained by Stottmeister et al. [76], who used sunflower oil or alkanes as a substrate for the production of up to 250 g l1 citric acid, in fed-batch cultures. Marty

et al. developed a mineral medium fulfilling the nutritional requirements of Y. lipolytica for the production of the extracellular lipase Lip2 which is known to have a high enantiomeric resolution capacity and catalyzes re-esterification reactions [77], yielding over 60,000 U l1 Lip2. The ability of Y. lipolytica to utilize and accumulate HS has led to the use of this yeast for the production of specific added-value oils from raw materials. Lipid composition and accumulation capacity in Y. lipolytica depend strongly on growth and culture conditions, providing a broad range of choice for substrate/final product combinations. For example, replacing glucose by oleic acid as the carbon source increases lipid accumulation capacity and the size of lipid particles and also modifies the composition of lipids and proteins [78]. Depending on the composition of the substrate used, this yeast selectively removes or assimilates FA from the substrate to produce fats with a predetermined composition [43]. Papanikolaou et al. [79] used these properties to produce cocoa butter-like substances from mixtures of inexpensive substrates, such as saturated fat and raw glycerol. In a similar procedure, Schrader et al. obtained high yields of c-lactone aroma from castor oil [80]. Several procedures for increasing lipid production in Y. lipolytica have been proposed. Fine adjustments of culture conditions can be used to upregulate lipid metabolism: Bati et al. reported a major effect of dissolved oxygen, nitrogen/carbon ratio, pH and amount of oil substrate on lipid accumulation, resulting in yeasts containing 37–70% lipid [81]. Aggelis et al. obtained yeasts that accumulated lipids to 43% of dry biomass, with a volumetric productivity of 0.12 g lipid l1 h1, using industrial fats or glycerol – both of which are cheap industrial carbon side-products – as the substrate [46]. In addition to the regulation of nutrient levels and culture conditions, genetic engineering approaches have been developed, based on the potential of yeasts as cell factories for the production of large amounts of oil with a particular lipid composition and added-value metabolites. Redirection of carbon flux toward TAG assembly mediated by deletion of the glycerol-3-phospate dehydrogenase gene (GUT2), an enzyme situated at the crossroad of lipid metabolism and DHAP production for glycolysis, increases lipid accumulation to levels three times those observed in wild-type strains [22]. The additional deletion of the POX1 to POX6 genes encoding the acyl-CoA oxidases, completely abolishes b-oxidation, preventing lipid mobilization. The resulting mutant strain had lipid levels four times those of the wild-type strain. In addition, the POX mutant genotypes affected lipid profile, as each POX gene has a different substrate specificity. POX2 deletion blocks the use of longchain fatty acids, whereas POX3 deletion decreases the uptake of short-chain fatty acids [82]. In mutants producing only Aox4p, or Aox1p and Aox6p, only a few, small intracellular LB are formed, resulting in a so-called ‘‘slim yeast” phenotype. Slim yeasts cannot accumulate the substrate, probably due to the feedback regulation of oxidation, affecting HS transporters and downregulating the diffusion process. By contrast, strains producing either Aox2p alone or both Aox2p and Aox4p form fewer, but larger, LB than the wild type. The overexpression of Aox2p in the Dpox2–5 quadruple mutant restores lipid accumulation, by increasing non-polar lipid storage in the LB, resulting in an ‘‘obese yeast” phenotype. These findings suggest a regulatory mechanism preventing or enabling lipid storage as a function of the ability of the yeast to assimilate the substrate from the medium. The LB phenotype also depends on the acyltransferase profile of mutant strains, with strains lacking ARE2 sterol-acyltransferase being able to form large LB. 6. Potential applications A potential application of Y. lipolytica or of oleaginous microorganisms in general is the production of single-cell oils (SCO).

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Single-cell oils may be defined as edible oils obtained from microorganisms and of a similar type and composition to the oils and fats obtained from plants or animals. SCO are now accepted as biotechnological products fulfilling key roles in the supply of major polyunsaturated fatty acids (PUFA), which are known to be essential for human nutrition and development. The commercial niche targeted by SCO is that of dietary supplements enriched in docosahexaenoic acid (DHA), arachidonic acid (AA) and c-linolenic acid (GLA) [3]. Picataggio et al. (pending US patent application Ser. No. 10/840,579) recently demonstrated the feasibility of engineering Y. lipolytica for the production of x-3 and x-6 fatty acids (e.g. 18:3, x-3, ALA, a-linoleic acid; C18:3, x-6, GLA, c-linoleic acid; 18:4, x-3, STA, stearidonic acid; 20:3, x-3, ETrA, eicosatrienoic acid and x-6, DGLA, dihomo-c-linoleic acid; 20:4, x-6, ARA, arachidonic acid; 20:5, x-3, EPA, eicosapentaenoic acid and 22:6, x-3, DHA, docosahexaenoic acid), by introducing and expressing heterologous genes encoding the proteins of the x-3/x-6 biosynthetic pathway in the oleaginous host. Damude et al. expressed in Y. lipolytica a bifunctional D12/x3 desaturase from Fusarium moniliformis. This strain produced a-linolenic acid (ALA, 18:3 D9,12,15) to levels corresponding to 28% of its cellular dry mass [83]. Similarly, GLA production was obtained by overproducing the D6 and D12 desaturases from Mortierella alpina in Y. lipolytica under the control of a strong constitutive promoter [84]. Furthermore, Dupont de Nemours have identified and isolated genes encoding acytransferases suitable for the transfer of these newly synthesized PUFA into TAG. These modified strains have been patented (US Patents 7267976, 7238482, 7256033) for the marketing of dietary supplements based on PUFA to protect against cardiovascular disease. One potential application of a Y. lipolytica strain with an altered POX genotype grown on a medium with a specific composition would be the production of dicarboxylic acids (DCA) or diacids [2]. Y. lipolytica strains are also regarded as very promising agents for the treatment of mineral oil pollution and plant oil waste. Oil mill wastewater (OMW), in particular, is a major source of water pollution as it contains fats, sugars, phosphate, phenols and metals. Scioli and Volaro [74] reported an 80% decrease in the COD of OMW after treatment for 24 h treatment with Y. lipolytica, whereas Oswal [85] found this yeast to be capable of decreasing COD by 90% in palm oil mill wastewater. Efforts are currently being made to generate strains with extreme oil accumulation capacities and specific fatty acid compositions. These studies are based on the regulation of substrate transport mechanisms, the overproduction or elimination of desaturases involved in TAG synthesis and the strict control of genes involved in fatty acid metabolism. The preliminary results obtained are very promising and potential applications have been identified in the areas of SCO and biodiesel production. Re-engineering microbial metabolism to favor oil production for fuel use seems to be the way forward to a third generation of biofuels: oils will be produced from more appropriate materials (such as cellulose, glycerol, or even oil waste) in better ways. The capacity of Y. lipolytica to produce long-chain molecules and to adjust its metabolism for the production of specific oils could lead to the production of more energy-efficient fuels. 7. Summary, conclusions and future directions Studies of lipid accumulation are currently focusing on maximizing oil production in unicellular organisms, such as yeast or algae, and in plants, for the generation of bio-fuels, including biodiesel. Like fossil hydrocarbons, TAG oils are highly concentrated stores of saturated hydrocarbons that can be oxidized to generate energy. The metabolism of Y. lipolytica, oriented toward the accumulation of lipids, and the unique capacity of this yeast to use HS efficiently, make this microorganism a prime candidate

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for use in the production of bio-oils. The use of this yeast as a model organism for investigating the mechanisms underlying these metabolic pathways has already provided insight into substrate transport processes, the function and use of the diverse organelles in metabolic processes and the identification and regulation of genes involved in these processes. However, although research on yeast has generated a number of important results concerning TAG and SE synthesis, many questions remain unanswered. One of these questions concerns the redundancy of the enzymes involved in lipid metabolism. The multiple functions of these proteins, which may act as acyltransferases or hydrolases, generating similar products from similar substrates, raises questions about the existence of multiple copies of enzymes to serve as a back-up for important metabolic pathways. These enzymes may be subject to multiple regulation procedures, including classical regulation at the transcriptional and translational levels, post-translational modifications of enzymes, and additional coordination at the organelle level. In addition, certain enzymes may be involved in protein–protein interactions or subject to differential regulation by auxiliary proteins, through processes that remain to be clarified [86]. For example, LB-associated proteins characteristically have PAT domains in their primary sequences. The PAT domain is defined by a conserved amino-acid sequence present in perilipin, adipophilin (also named adipocyte differentiation-related protein, ADRP), and the tail-interacting protein of 47 kDa (TIP47) [87]. The PAT proteins (perilipin, adipophilin/adipose differentiation-related protein, TIP47, and other related proteins) have structural and regulatory functions and are thought to target lipid bodies through different mechanisms [88]. Some PAT proteins are found exclusively on lipid droplets (e.g. adipophilin), whereas others are found both in the cytosol and on lipid droplets. The structural properties underlying the binding of PAT proteins to lipid droplets provide a large number of possibilities for regulating this distribution. Indeed, the localization of several proteins, such as TIP47, to lipid droplets is highly regulated and can be induced by adding fatty acids to the medium to trigger droplet formation [89]. However, the mechanism underlying this regulation remains largely unclear. It has been suggested that lipase access to the core of the lipid droplet is regulated in part by PAT proteins, and the phosphorylation of perilipin may indeed be crucial for the regulation of lipase access to substrates [90]. It has been estimated that PAT proteins cover 15–20% of the droplet surface, possibly resulting in steric hindrance, restricting lipase access. However, the hydrolysis and mobilization of sterol esters is less well understood. In S. cerevisiae, sterol esters and TAG are hydrolyzed by three consecutive steps (Yehl, Yeh2, and Tgl1 for SE, Tgl3, Tgl4, and Tgl5 for TAG) catalyzed by enzymes present on the surface of lipid bodies [91,92]. Little is known about the regulation of the activities of these enzymes, but PAT proteins may be involved in the limiting step of hydrolysis. In addition to the small number of lipid droplet-specific proteins identified, such as PAT proteins, many cellular proteins with known functions have been identified in lipid body fractions in proteomic studies. Some of these proteins are involved in dynamic cellular processes, suggesting a possible similar role in lipid droplet biology [78,89,93]. However, both the localization of most of these proteins to droplets and their functional roles remain to be validated. Another unanswered question relating to lipid metabolism concerns LB biogenesis. Insight into this process might also provide clues to the role of auxiliary proteins in the early steps of LP formation at the molecular level. In conclusion, too few detailed studies of the functional and structural properties of enzymes involved in lipid metabolism have been carried out. In one such study, a polymorphism in DGAT was recently identified as responsible for a quantitative trait locus controlling high levels of oil production in maize. A single

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amino-acid substitution boosted oil content by up to 41% and oleic acid content to 107% [94]. An understanding of the molecular processes governing oil storage in lipid bodies might lead to novel approaches to the engineering of unicellular microorganisms. Improvements in our understanding of the regulatory and structural properties of the enzymes involved in lipid metabolism and the use of genetic techniques for the engineering of these organisms would make it possible to maximize cellular lipid storage and, therefore, oil production per cell. A combination of modifications to the culture conditions and the regulated expression of key enzymes (native or indigenous) could be used to produce specific lipid compounds with added value and of potential interest in the oleochemical and petrochemical field. The study of de novo lipid accumulation as a result of the conversion of carbohydrate substrates, such as glucose, provides insight into the regulation of the entire lipid synthesis pathway. The accumulation of lipids produced from these substrates is triggered by nutrient limitation. Nitrogen limitation is the principal type of limitation studied and the C/N ratio is the key factor governing lipid accumulation. Most ex novo lipid accumulation studies have been carried out in batch mode. However, the fed-batch mode seems to be the best culture system, because carbon and nitrogen flows can be perfectly controlled, making it possible to dissociate growth and lipid accumulation and the absence of by-product secretion. Novel approaches combining lipidomic, metabolomic and genetic approaches and making use of fed-batch cultures will undoubtedly provide a wealth of information about the regulation of lipid metabolism. Acknowledgements A. Beopoulos was supported by the Lipicaero French national research program (ANR-PNRB) No. 0701C0089. J. Cescut was supported by CNRS and Airbus. R. Haddouche was supported by the European research program (LIPOYEAST). This work was financially supported in part by Aerospace Valley. We thank Julie Sappa of Alex Edelman and Associates for her excellent help in correcting the English version of the manuscript. References [1] Barth G, Gaillardin C. Yarrowia lipolytica. In: Wolf K, editor. Nonconventional yeasts in biotechnology. Berlin, Heidelberg, New York: Springer-Verlag; 1996. p. 313–88. [2] Fickers P, Benetti PH, Wache Y, Marty A, Mauersberger S, Smit MS. Hydrophobic substrate utilisation by the yeast Yarrowia lipolytica, and its potential applications. FEMS Yeast Res 2005;5:527–43. [3] Ratledge C. Single cell oils for the 21st century. In: Cohen, Ratlege, editors. Single cell oils. Champaign: AOCS Press; 2005. p. 1–20. [4] Mlickova K, Roux E, Athenstaedt K, d’Andrea S, Daum G, Chardot T, et al. Lipid accumulation, lipid body formation, and acyl coenzyme A oxidases of the yeast Yarrowia lipolytica. Appl Environ Microbiol 2004;70:3918–24. [5] Zweytick D, Athenstaedt K, Daum G. Intracellular lipid particles of eukaryotic cells. Biochim Biophys Acta 2000;1469:101–20. [6] Brown DA. Lipid droplets: proteins floating on a pool of fat. Curr Biol 2001;11:446–9. [7] Fujimoto T, Ohsaki Y, Cheng J, Suzuki M, Shinohara Y. Lipid droplets: a classic organelle with new outfits. Histochem Cell Biol 2008;130:263–79. [8] Athenstaedt K, Jolivet P, Boulard C, Zivy M, Negroni L, Nicaud JM, et al. Lipid particle composition of the yeast Yarrowia lipolytica depends on the carbon source. Proteomics 2006;6:1450–9. [9] Ferreyra RG, Burgardt NI, Milikowski D, Melen G, Kornblihtt AR, Dell’ Angelica EC, et al. A yeast sterol carrier protein with fatty-acid and fatty-acyl-CoA binding activity. Arch Biochem Biophys 2006;453:197–206. [10] Beckman M. Cell biology. Great balls of fat. Science 2006;311:1232–4. [11] Papanikolaou S, Chevalot I, Komaitis M, Marc I, Aggelis G. Single cell oil production by Yarrowia lipolytica growing on an industrial derivative of animal fat in batch cultures. Appl Microbiol Biotechnol 2002;58:308–12. [12] Czabany T, Athenstaedt K, Daum G. Synthesis, storage and degradation of neutral lipids in yeast. Biochim Biophys Acta 2007;1771:299–309. [13] Ratledge C. In: Ratledge C, Wilkinson SG, editors. Microbial lipids, vol. 2. London: Academic Press; 1988. [14] Rattray JBM. In: Ratledge C, Wilkinson SG, editors. Microbial lipids, vol. 1. London: Academic Press; 1988. p. 552–687.

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