Biotechnological applications of Yarrowia lipolytica ...

JBA-06962; No of Pages 25 Biotechnology Advances xxx (2015) xxx–xxx

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Research review paper

Biotechnological applications of Yarrowia lipolytica: Past, present and future Hu-Hu Liu, Xiao-Jun Ji ⁎, He Huang ⁎ State Key Laboratory of Materials-oriented Chemical Engineering, College of Biotechnology and Pharmaceutical Engineering, Nanjing Tech University, No. 30 South Puzhu Road, Nanjing 211816, People's Republic of China

a r t i c l e

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Article history: Received 21 August 2014 Received in revised form 13 July 2015 Accepted 29 July 2015 Available online xxxx Keywords: Yarrowia lipolytica Characteristics Physiology Metabolism Applications

a b s t r a c t Non-conventional yeasts have attracted increasing interest due to their biochemical characteristics and potential applications. Yarrowia lipolytica is a non-conventional yeast with specific characteristics and physiology. The potential physiological and metabolic capabilities of Y. lipolytica, which can assimilate many different carbon sources, including typical hydrophilic and hydrophobic materials, are reviewed in this paper. Concerning the uptake and metabolism substrates, this review focuses particularly on low-cost raw materials, such as glycerol. Moreover, this review presents the results of safety studies of Y. lipolytica. Finally, the wide applications of Y. lipolytica, such as functional enzyme production, metabolite synthesis and environmental bioremediation, are reviewed in this paper. Recently, with the development of system biology and synthetic biology, it was concluded that these technologies will provide new opportunities for potential applications of Y. lipolytica in the future. © 2015 Elsevier Inc. All rights reserved.

Contents 1. 2.

Introduction . . . . . . . . . . . . . . . . Characteristics and physiology of Y. lipolytica . 2.1. Characteristics . . . . . . . . . . . 2.2. Carbon sources . . . . . . . . . . . 2.2.1. Hydrophilic materials . . . . 2.2.2. Hydrophobic materials . . . 2.3. Safety studies on products of Y. lipolytica 3. Applications of Y. lipolytica . . . . . . . . . 3.1. Secretion of proteins . . . . . . . . . 3.2. Organic acids . . . . . . . . . . . . 3.3. Single cell oils . . . . . . . . . . . . 3.4. Bioremediation . . . . . . . . . . . 3.5. Aromatic compounds . . . . . . . . 3.6. Others . . . . . . . . . . . . . . . 4. Conclusions and prospects . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . .

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1. Introduction With economic development and the energy crisis, microbial-based biotechnology has been a promising alternative route for the production ⁎ Corresponding authors. E-mail addresses: [email protected] (X.-J. Ji), [email protected] (H. Huang).

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of valuable chemicals. Among these species, non-conventional yeasts, compared with conventional yeasts such as Saccharomyces cerevisiae, have been deemed important microorganisms based on their biochemical characteristics and potential applications. As early as the 1960s, Yarrowia lipolytica, one of the non-conventional yeasts with potential biotechnological applications, has been studied for single-cell protein (SCP) production. Historically, this yeast was originally isolated from

http://dx.doi.org/10.1016/j.biotechadv.2015.07.010 0734-9750/© 2015 Elsevier Inc. All rights reserved.

Please cite this article as: Liu, H.-H., et al., Biotechnological applications of Yarrowia lipolytica: Past, present and future, Biotechnol Adv (2015), http://dx.doi.org/10.1016/j.biotechadv.2015.07.010

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H.-H. Liu et al. / Biotechnology Advances xxx (2015) xxx–xxx

lipid-rich materials such as rancid butter; therefore, it was given the species name “lipolytica”. As an ascomycetous yeast, it was first identified as Candida lipolytica in the late 1960s and was then reclassified as Endomycopsis lipolytica, Saccharomycopsis lipolytica and finally Yarrowia lipolytica (Barth and Gaillardin, 1996). Y. lipolytica, due to its potential of producing lipases and proteases, can be isolated from lipid-rich or protein-rich environments such as dairy products, cheese, yogurt, meat, poultry and olive oil. Generally, Y. lipolytica can secrete many endogenous enzymes with functional applications. Moreover, compared with other well-characterized yeasts such as S. cerevisiae and Pichia pastoris, Y. lipolytica has particular genetic advantages, as follows. First, protein is secreted primarily by the co-transcription pathway (Domínguez et al., 1998); second, it has a high secretion capacity and low glycosylation modification; third, it is a non-pathogenic yeast; and lastly, as an obligate aerobe, Y. lipolytica is fit for cultivation via a high cell-density fermentation mode. Based on its mentioned genetic advantages, Y. lipolytica is considered a suitable host for protein expression. To date, many articles have reported the specific physiological and genomic characteristics as well as metabolic and biological applications of Y. lipolytica (Coelho et al., 2010; Gonçalves et al., 2014; Madzak et al., 2004; Nicaud, 2012; Titorenko et al., 2000; Zinjarde, 2014; Zinjarde et al., 2014). In particular, Madzak et al. (2004) presented a comprehensive review of the genetic and molecular tools of Y. lipolytica for heterologous protein production. Coelho et al. (2010) showed a review of the specific physiological features and industrial applications of Y. lipolytica. Papanikolaou and Aggelis (2011a, 2011b) presented various models on the accumulation and degradation of the microbial lipids of various oleaginous yeasts (including Y. lipolytica) with the potential of assimilating and utilizing various carbon sources (such as industrial fats, glycerol and glucose, etc.). Significantly, many potential metabolites, such as organic acids and microbial lipids, could be produced by Y. lipolytica when raw materials were used as carbon sources. Zinjarde (2014) and Zinjarde et al. (2014) reviewed the comprehensive environment-related and food-related biological applications of Y. lipolytica. Moreover, books that summarize special physiological features and present biotechnological applications and cutting-edge research on Y. lipolytica have also been published (Barth, 2013a, 2013b; Harzevili, 2014). Although significant progress has been made on Y. lipolytica, some unresolved problems remain for further research. One of these problems is that some characteristics are strain dependent. For example, André et al. (2009) showed that mannitol, a principal metabolic product, is synthesized by Y. lipolytica LFMB 19 and LFMB 20 grown on crude glycerol (initial concentration of 30 g/L) as a carbon source in nitrogen-limited culture. In contrast, citric acid, a principal organic acid, was produced primarily by Y. lipolytica ACA-YC 5033 under the same culture conditions. Moreover, the mechanisms of many enzymes involved in different biochemical reactions remain unclear. With the development of biotechnology, particularly the sequencing of the genome of Y. lipolytica and synthetic biology and combinational engineering, this article will provide more potential new ideas for researchers committed to this field. Therefore, the purpose of this article is to summarize and provide comprehensive information about the characteristics, physiology, results of safety studies and important applications of Y. lipolytica. This review will serve as a meaningful resource for increasing numbers of researchers working in this field. 2. Characteristics and physiology of Y. lipolytica 2.1. Characteristics Y. lipolytica has six chromosomes with high G + C contents of 49.6%– 51.7% and a genome size of approximately 12.7 to 22.1 Mb (Barth and Gaillardin, 1996). With the development of genome sequencing, the genomes of three strains, Y. lipolytica CLIB122, WSH-Z06 and Po1f, are available on the NCBI website (http://www.ncbi.nlm.nih.gov/genome/

genomes/194) (Liu and Alper, 2014). Moreover, the genome of Y. lipolytica CLIB89 was partially sequenced and made available by the Genolevures Consortium (http://www.genolevures.org/) (Dujon et al., 2004). In addition, the sequence of the mitochondrial genome with a size of 47.9 kb from Y. lipolytica was reported (Kerscher et al., 2001). Y. lipolytica is a typical heterothallic yeast that has two mating types, Mat A and Mat B. Most of the natural isolated strains are haploid. A diploid strain will be produced only if two types mate. However, the mating frequency is very low. Both the haploid and diploid states are stable under laboratory conditions (Thevenieau et al., 2009). To date, several inbred lines of Y. lipolytica have been obtained by different groups originating from the German (H222), French (W29) and American (CBS6214-2) strains (Nicaud, 2012). Y. lipolytica can assimilate and utilize different water-soluble or water-insoluble carbon sources. As an obligate aerobe, Y. lipolytica is very sensitive to oxygen concentration, which is deemed an important factor for the lifecycle growth and metabolism. Temperatures suitable for growth must not exceed 32–34 °C (Beopoulos et al., 2010). Naturally, the wild-type strains have various colony shapes, from smooth and glistening to heavily convoluted and matte. Y. lipolytica is non-pathogenic and is generally regarded as safe (GRAS) by the Food and Drug Administration (FDA). As a dimorphic yeast, Y. lipolytica has different cellular growth forms, such as yeast cells and mycelium. To date, little is known regarding the genetic regulation of the process that leads to dimorphism (Casaregola et al., 2000). Domínguez et al. (2000) determined that the transition of Y. lipolytica is related to the environmental conditions and genetic regulatory mechanism. There are two opposite signal mechanisms in Y. lipolytica, including the nitrogen-activated protein kinase pathway and the cyclic AMP-dependent protein kinase pathway, which play important functions during yeast shape transition. Domínguez et al. (2000) showed that N-acetyl glucosamine or serum acts on the same signal pathways during the transition (Domínguez et al., 2000). PérezCampo and Domínguez (2001) revealed that cell growth is primarily like that of mycelia when Glc-NAc or serum is used as a carbon source, whereas it is yeast-like in the shape of Y. lipolytica when grown on glucose as a carbon source. In this research, the transition was induced at a lesser extent with a medium containing bovine albumin. GuevaraOlvera et al. (1993) reported that polyamines are synthesized by mycelial cells rather than by yeast cells. In this research, it was concluded that polyamines play an important role in cell growth during the physiological transition. When hydrophobic palm-oil mill effluent was used as a carbon source, Y. lipolytica NCIM 3589 formed a true mycelium morphology transition (Oswal et al., 2002). Papanikolaou et al. (2007) observed that the yeast-true mycelia transition occurs in Y. lipolytica strain ACA-DC 50109 when tallow is used as the sole carbon source. Zinjarde et al. (2008) reported that the mycelial formation is observed in Y. lipolytica NCIM 3589 when various crude oils, such as coconut oil and palm kernel oil containing lauric and myristic acids, were used as carbon sources. However, a reverse transition to yeast cells was observed when saturated n-dodecane was used as a substrate. In addition, other environmental factors including pH, nitrogen starvation and temperature also have effects on the physiological transition of Y. lipolytica. Ruiz-Herrera and Sentandreu (2002) reported the effects of various environmental factors (including pH, carbon and nitrogen sources) on mycelium formation in Y. lipolytica W29 and Po1a. In particular, this research showed that pH, carbon and nitrogen sources (usually glucose and ammonium) are important factors for mycelium formation. Bellou et al. (2014) demonstrated that the dissolved oxygen concentration is a major factor that affects yeast morphology. Morín et al. (2007) analyzed the proteome of the wild-type and mutant Y. lipolytica during the yeast-to-hyphae transition using a two-dimensional gel electrophoresis (2-DE) and mass spectrometry method. In their research, 45 different proteins were detected, and some of the identified proteins were determined to be involved in metabolic pathways. Moreover, M. Li et al. (2014) determined that three Ras proteins in Y. lipolytica (YlRas1p, YlRas2p and YlRas3p) are critical for dimorphic transition. Among these



Ras proteins, YlRas2p played a major role in the regulation of dimorphic transition. Moreover, Y. Q. Li et al. (2014) demonstrated that the Rap GTPase YlRsr1p also plays an important role in the morphogenesis of Y. lipolytica. Although there are no natural plasmids in Y. lipolytica, a series of molecular tools have been developed for Y. lipolytica (Juretzek et al., 2001; Madzak et al., 2004; Nicaud et al., 2002; Spencer et al., 2002). Moreover, Fickers et al. (2003) developed a new method for rapid gene disruption, which combined the sticky-end PCR approach and the Cre–lox recombination system for Y. lipolytica. To date, the genetic tools for Y. lipolytica have been summarized (Abghari and Chen, 2014; Madzak et al., 2004). The vectors used for transformation, such as shuttle vectors including replicative (episomal) vectors and integrative vectors, are typically composed of a selection marker, an expression or secretion gene cassette and a target element. The typical components of vectors for Y. lipolytica are presented in Fig. 1. Generally, the replicative vectors contain a centromere (CEN) and an origin of replication (ORI). To date, some autonomously replicating sequences (ARS), such as ARS18 and ARS68, which carry a CEN and chromosomal ORI, have been identified as having replicative functions and can be cloned in Y. lipolytica. However, compared with integrative vectors, the copy numbers of ARS plasmids were low (only 1–3 copies/ cell) (Fournier et al., 1993; Vernis et al., 1997). Recently, L. Liu et al. (2014) developed a set of engineered CEN plasmids via altering its centromere function to improve the copy number and gene expression at the plasmid level of Y. lipolytica. Meanwhile, integrative vectors that carry functional target elements have been developed for integration into the chromosome of Y. lipolytica via homologous or nonhomologous recombination. The functional target elements in integrative vectors generally include one copy of tandem repeat sequences of rDNA, a single-copy gene such as XPR2, or one copy of dispersed repeated sequences such as the “zeta” sequence of Ylt1 (Barth and Gaillardin, 1996; Le Dall et al., 1994). Notably, a total of more than 200 copies of rDNA clusters was identified in Y. lipolytica. Moreover, Ylt1, a repetitive retrotransposon flanked by a long terminal repeat named the “zeta” sequence, was detected in the yeast genome (Schmid-Berger et al., 1994). It was shown that Ylt1-carrying Y. lipolytica presented at least 100 potential “zeta” sequence integration sites. Therefore, the expression cassette flanked by these functional sequences can be integrated into the chromosome of Y. lipolytica via homologous recombination. However, Ylt1 was not present in all wild-type strains. It was obviously present

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in the American strain CBS6214-2, but it was absent in the German H222 and French W29 wild isolates (Beopoulos et al., 2010). Moreover, the long terminal repeat-retrotransposon elements were confirmed to be absent in the strain Y. lipolytica Po1f (Liu and Alper, 2014). Notably, “zeta” sequences were able to direct random integration into Y. lipolytica strains devoid of the Ylt1 sequence via non-homologous recombination. Recently, Kretzschmar et al. (2013) developed an effective method for increasing the rate of heterologous recombination by deleting YlKU70 and YlKU80 in Y. lipolytica. Using “zeta” regions as a docking platform in Y. lipolytica, many efficient expression systems have been developed for tuning protein expression levels (Cambon et al., 2010; Pignède et al., 2000). Bordes et al. (2007) developed a new recombinant protein expression system by constructing a strain containing a “zeta” docking platform for transformation. Based on error-prone PCR, Bordes et al. (2011) presented a library of lipase Lip2p variants in the Y. lipolytica strain JMY 1212 using the “zeta” sequence as target sites. The transformation methods for Y. lipolytica include the heat shock method and the electroporation method (Chen et al., 1997; Davidow et al., 1985; Fournier et al., 1993; Wang et al., 2011). Chen et al. (1997) described an efficient one-step transformation protocol with higher transformation efficiency via the heat shock treatment for Y. lipolytica. Recently, Wang et al. (2011) developed a modified electroporation method that combined with lithium acetate and dithiothreitol pretreatment to enhance the transformation frequency for Y. lipolytica Po1g. However, the transformation efficiency of a Ylt1-carrying or Ylt1-free strain with a “zeta”-based vector carrying a defective selection marker is lower than a classical integrative vector. The markers, including auxotrophy markers, such as LEU2 and URA3, and dominant markers, such as hph-encoding hygromycin B, can be used for strain screening (Madzak et al., 2004). Usually, ura3d4, a defective version of the URA3 marker, was designed in different vectors for selecting multiple integrations. The gene cassette, an important element, is comprised of a promoter, a functional gene and a terminator. Promoters play an important role in gene expression. Based on reporter protein activity assays such as β-galactosidase (EC 3.2.1.23) and β-glucuronidase (EC 3.2.1.31), many suitable promoters, including constitutive and inducible promoters (such as pPOX2, pG3P, pTEF and pFBA1IN) have been selected (Hong et al., 2011a; Juretzek et al., 2000). In addition, a strong hybrid promoter, hp4d, was constructed; it has wide potential for driving protein expression (Madzak et al., 2000). With the advent of synthetic biology, tuning a gene expression via promoter engineering has received

Fig. 1. The typical components of expression or secretion vector for Yarrowia lipolytica.


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more attention. For a Y. lipolytica expression system, a promoter library was constructed via hybrid promoter engineering. A series of highstrength hybrid promoters, such as UAS1B1-32-Leum, UAS1B8-TEF and UAS1B16-TEF, which were composed of activating regions (upstream activation sequences, UAS) and core promoter regions, were selected based on reporter protein activity assays and quantitative reverse transcription-PCR assays (Blazeck et al., 2011). Blazeck et al. (2013b) generated more constitutive hybrid promoters with the potential to precisely tune functional gene expression; they were constructed with a newly isolated UAS element from a hybrid promoter library for Y. lipolytica. Terminators are also important elements in the transcription process. To date, terminator engineering has been applied in the expression systems of E. coli and S. cerevisiae (Cambray et al., 2013; Curran et al., 2013). Recently, Curran et al. (2015) reported that several of the designed synthetic terminators are confirmed to be highly functional in Y. lipolytica. In addition, suitable secretion signals are very important for protein expression and secretion. Using Y. lipolytica as a host, some homologous or heterologous secretion signals, such as the XPR2 prepro or LIP2 prepro region, have been used for protein expression and secretion. Hofmeyer et al. (2014) established a strategy for protein production and purification using Lip2p as a fusion protein in Y. lipolytica. Recently, a yeast expression kit for Y. lipolytica has been developed and can be purchased from Yeastern Biotech Co., Ltd (http://www. yeastern.com/). Moreover, the techniques of surface display on Y. lipolytica have attracted growing interest. Because Y. lipolytica has many advantages as an expression host, some cell wall proteins, including YlCWP1p, YlYWP1p and YlPir1p, have been identified. Based on the glycosylphosphatidylinositol (GPI) anchor domain of YlCWP1p, a new surface display vector, pINA1317-YlCWP110p, was developed for Y. lipolytica (Yue et al., 2008). Yang et al. (2009) constructed a novel surfacedisplay system for Y. lipolytica using the cell-wall anchor protein Flo1p from S. cerevisiae. In this study, a surface-displayed mannanase fused with the C-terminus of Flo1p with the potential to display high activity was identified. Recently, Bulani et al. (2012) developed a novel rDNAbased plasmid used to display heterologous proteins on the cell surface of Y. lipolytica using the C-terminal end of GPI-anchored YlCWP1p. To date, the surface-display system has been used to display many functional proteins (Liu et al., 2009, 2010; Ni et al., 2009; Yu et al., 2010; Song et al., 2011; Wang et al., 2009, 2012b). Through fusing GPIanchoring motifs derived from YlCWP1p and YlYWP1p, Moon et al. (2013) described the successful surface display of α-1, 2-mannosidase (EC 3.2.1.130) from Aspergillus saitoi on the cell surface of Y. lipolytica. Duquesne et al. (2014) reported the display activity of xylanase TxXYN (EC 3.2.1.8) from Thermobacillus xylanilyticus on the surface of Y. lipolytica JMY1212 with three different fusion proteins (YlCWP110p, YlPir100p and YlCBM87p). Based on the determination of xylanase activity, this research represented that YlPir100p is an efficient protein anchor among these display systems. 2.2. Carbon sources With non-renewable resource consumption and environmental deterioration, the development of an alternative biotechnological pathway for producing important chemicals has attracted growing interest. As a nonconventional yeast, Y. lipolytica can assimilate and ferment different carbon sources, such as hydrophilic materials (glucose, glycerol, alcohols and acetate) and hydrophobic substrates (fatty acids, triacylglycerols and alkanes), and produce important metabolites. Huang et al. (2013) presented a review of the use of low-cost raw materials such as molasses, wastewaters, industrial fat and vegetable oils as carbon sources for industrial applications. It is a fact that either hydrophilic or hydrophobic materials can be assimilated and metabolized through intrinsic assimilation pathways. In this review, the metabolic processes of Y. lipolytica grown on typical carbon sources such as glucose, glycerol, ethanol, acetate, fatty acids, alkanes and triacylglycerols are illustrated in Fig. 2.

2.2.1. Hydrophilic materials Sugars, as efficient carbon substrates, including hexoses (such as glucose, fructose, mannose and galactose) and other sugars (such as lactose), can be assimilated by yeasts. Flores et al. (2000) reviewed the metabolism of different carbon sources and energy-yielding pathways in non-conventional yeasts. With Y. lipolytica, this yeast is different from S. cerevisiae and other yeasts and can assimilate several sugars, such as glucose and fructose, as carbon sources only. For glucose metabolism in Y. lipolytica, there is a transport system with two components that are independent of the glucose concentration (Flores et al., 2000). Among these efficient water-soluble sugars, sucrose cannot be taken up directly because Y. lipolytica lacks a gene encoding secreted invertase (EC 3.2.1.26). Based on the development of genetic tools, the genetically modified (GM) strain Y. lipolytica was constructed for producing invertase to metabolize sucrose (Nicaud et al., 1989). Moreover, Lazar et al. (2011, 2013) revealed that Y. lipolytica has a preference of glucose consumption over fructose when grown on a mixture of glucose and fructose. Recently, Lazar et al. (2014) reported that fructose uptake can be successfully improved by overexpressing hexokinase. Glycerol is a byproduct of biodiesel production that can be assimilated and fermented as a carbon source by many microorganisms to produce functional chemicals. Naturally, pure glycerol with high purity (glycerol at least 98%) can be used as an important industrial feedstock for producing many important chemicals. However, raw glycerol, differing from pure glycerol, has low purity (glycerol in the range of 75–83%) and is heavily contaminated with various impurities, such as salts, methanol and residual fatty acids plus many other undesirable materials. Therefore, raw glycerol cannot be used as a carbon source for any products that will be used as a food or feed. Chatzifragkou and Papanikolaou (2012) reviewed how biodiesel-derived crude glycerol can be utilized by prokaryotic and eukaryotic microorganisms for various important chemical production under anaerobic and aerobic cultivation conditions. Generally, glycerol is assimilated and metabolized via the phosphorylation pathway for the synthesis of biotechnological products (Makri et al., 2010). As a potential raw material, the assimilation mechanism and applications of glycerol have been clearly reviewed (Amaral et al., 2009; Da Silva et al., 2009; Rywińska et al., 2013a). Using pure glycerol as a carbon source, a one-step feeding process for the Y. lipolytica strain was developed and the higher activity of α-amylase (up to 88 U/mL) was obtained in recombinant Y. lipolytica YLASIn (Kim et al., 2000). Meanwhile, raw glycerol, a primary byproduct of the biodiesel production process, also can be used as a carbon source for microbial lipid and citric acid production (Papanikolaou and Aggelis, 2002; Papanikolaou et al., 2002b). Notably, when grown on a mixture of glucose and biodiesel-derived raw glycerol, the wild strain Y. lipolytica LGAM S(7)1 favors glycerol for citric acid production (Papanikolaou and Aggelis, 2002). Recently, Workman et al. (2013) also showed that glycerol is metabolized and depleted before glucose utilization in batch fermentation by Y. lipolytica IBT 446. In addition, Y. lipolytica can assimilate and ferment ethanol due to the presence of endogenous functional genes encoding alcohol dehydrogenases (EC 1.1.1.1) and aldehyde dehydrogenases (EC 1.2.1.3). Barth and Gaillardin (1996) showed in a review that Y. lipolytica is able to survive in a medium with an ethanol concentration of up to 3% (v/v). However, the higher concentrations of ethanol are not beneficial for the growth of Y. lipolytica because they will disrupt the cell membrane and inactivate some functional proteins. Notably, these functional enzyme activities are remarkably repressed in Y. lipolytica when glucose is used as a carbon source. Moreover, Y. lipolytica also can efficiently utilize acetate as the sole carbon source in the fermentation process. It was reported that concentrations up to 0.4% (v/v) of sodium acetate were well tolerated for assimilation and metabolism in Y. lipolytica, higher concentrations reduced the growth rate and concentrations above 1.0% (v/v) inhibited growth (Barth and Gaillardin, 1996). This happened because higher concentrations of sodium acetate could increase pH after acetate consumption, and the high pH could stop cell growth. Venter et al. (2004) developed a novel strategy that could improve the yield



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Fig. 2. The metabolic pathways of Yarrowia lipolytica using typical carbon sources. Abbreviations: ER: endoplasmic reticulum; PER: peroxisome; MIT: mitochondria; TCA cycle: tricarboxylic-acid cycle; LP: lipid body; DHAP: dihydroxyacetone phosphate; GA3P: glycerol-3-phosphate; CA: citric acid; OAA: oxaloacetate; PA: phosphatidic acid; LPA: lysophosphatidic acid; DAG: diacylglycerol; TAG: triacylglycerol; DCAs: dicarboxylic acids; PHAs: polyhydroxyalkanoates; HFAs: hydroxylated fatty acids; WEs: wax esters.

of citric acid from Y. lipolytica UOFS Y-1701 via adding acetate to the medium. Moreover, Fontanille et al. (2012) demonstrated that volatile fatty acids, such as acetic acid and propionic acid, can be used as carbon sources to increase biomass and lipid concentration through an effective two-stage fed-batch strategy.

2.2.2. Hydrophobic materials Hydrophobic materials, such as alkanes, fatty acids and triacylglycerols, can be assimilated and metabolized by Y. lipolytica to provide energy and produce functional chemicals. Papanikolaou and Aggelis (2011a) reviewed the utilization of hydrophilic and hydrophobic substrates


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and the accumulation and degradation of microbial lipids. It was stated that the lipid accumulation of oleaginous yeasts (including Y. lipolytica) occurred via an ex novo lipid synthesis pathway through the use of various hydrophobic substrates (such as triacylglycerols and fatty acid ethyl-esters) as carbon sources. Moreover, Papanikolaou et al. (2003, 2006) observed that the microbial lipids can be produced via a de novo pathway in Y. lipolytica grown on a mixture of stearin (a substrate composed by saturated free-fatty acids), glucose or raw glycerol. Notably, Papanikolaou et al. (2003) revealed that in the presence of stearin, higher glycerol uptake can increase citric acid production from Y. lipolytica ACA-DC 50109 compared with the cultures with glucose as a co-substrate. Moreover, it was speculated that the presence of lipids could affect the carrier of glucose but not that of glycerol. The assimilation and metabolism of hydrophobic materials in Y. lipolytica have been previously reviewed (Fickers et al., 2005a; Fukuda, 2013; Thevenieau et al., 2010). However, the transport mechanisms of these substrates are not clear. Generally, it is supposed that the uptake pathway of hydrophobic materials consists of a sequence of steps as follows: first, membrane-mediated assimilation; second, substrate transportation; third, substrate catabolism in different organelles; and last, new compound synthesis. When grown on hydrophobic carbon sources, Y. lipolytica secretes functional chemicals with the potential of degrading the substrate and reducing the size of substrate droplets. For example, TAGs are degraded into fatty acids and glycerol by lipase for further assimilation. In some cases, the yeast surface forms protrusions, which can attach small hydrophobic substrate droplets onto the cell surface. Then, these droplets are transported into the endoplasmic reticulum (ER) via the membrane-mediated transport and export system. It is a fact that different typical carbon substrates are degraded and synthesized via various metabolic pathways located in different cell compartments. Of the different cell compartments, the ER and peroxisome play important roles in metabolic processes. In particular, long-chain hydrophilic substrates are energy-rich but carbonlow materials that, unlike sugars of carbon-rich but energy-low materials, are degraded via β-oxidation reaction for providing latent energy in the peroxisome of Y. lipolytica. Naturally, the degradation reaction is performed by linking their metabolism to the production of FADH2, which is re-oxidized to FAD with the release of H2O2 and is then cleaved by catalase with the release of heat. The mechanism of the transport of fatty acids is not well understood. Usually, selective uptake occurs in Y. lipolytica during assimilating mixtures of various fatty acids as carbon sources. Aggelis et al. (1997) revealed that C. lipolytica incorporated C18 fatty acids in the order of C18:3 N C18:2 N C18:1 N C18:0 when grown on the evening primrose oil that was rich in the common C18 fatty acids, such as stearic acid (C18:0), oleic acid (C18 :1), linoleic acid (C18:2) and γ-linolenic acid (C18:3). Moreover, it was revealed that the oleic and linoleic acids were more rapidly incorporated than the saturated fatty acids for growth needs by the strain Y. lipolytica LGAM S(7)1 cultivated on mixtures of industrial fats containing stearic, oleic, linoleic and palmitic acid (Papanikolaou et al., 2001). In this research, it was observed that the saturated fatty acids were incorporated more slowly. Moreover, Papanikolaou and Aggelis (2003b) reported that various fatty acids are assimilated selectively by Y. lipolytica ACA-DC 50109 grown on mixtures of free fatty acid as substrates. It was observed that the unsaturated fatty acids (C18:3, C18:2 and C18:1) and shorter aliphatic chain fatty acids (C12:0 and C14:0) were more rapidly incorporated in a cell than the saturated fatty acids (C16:0 and C18:0). In 1984, it was suggested that two fatty acid carrier transporter systems, one specific for C12 and C14 fatty acids and the other for C16 and C18 unsaturated or saturated fatty acid uptake might exist in S. lipolytica CBS 2075 (Kohlwein and Paltauf, 1984). Meanwhile, similar observations of preferential assimilation of unsaturated fatty acids have been reported in other strains. For example, Lee et al. (1993) reported that the oleaginous yeast Apiotrichum curvatum ATCC 20509 favors the mixtures of oleic acid and linoleic acid over the mixtures of palmitic acid and stearic acid for triacylglycerol production. However,

shorter-chain fatty acids, such as octanoic acid (C8:0) and decanoic acid (C10:0), cannot be assimilated efficiently. It is possible that shorter-chain fatty acids may have a negative effect on the cell membrane. The transport mechanism and assimilation mechanism of alkanes remain unclear. Generally, two models that include a passive diffusion process and an active and energy-dependent mechanism occur in Y. lipolytica. A chain length-dependent alkane uptake mechanism that discriminates between short carbon-chain alkanes (C10 or C12) and long carbon-chain alkanes (C14 or C16) was revealed in Y. lipolytica (Thevenieau et al., 2007). Usually, shorter-chain alkanes from C5 to C10 cannot be assimilated because they are cytotoxic and simply disrupt the cell membrane. Fukuda (2013) presented a comprehensive review on the metabolism of n-alkanes and its transcriptional control in Y. lipolytica. It is generally supposed that alkanes are first converted into the corresponding fatty alcohols of the same chain length by the alkane monooxygenase system (AMOS system), which is composed of a cytochrome P450 (belonging to the CYP52 family) and NADPHdependent cytochrome P450 reductase (EC 1.6.2.4). It was revealed that there was a single gene coding for cytochrome P450 reductase, whereas there were 12 genes (ALK1–ALK12) coding for cytochrome P450 isoforms (Thevenieau et al., 2010). Recently, Opi1 family members, such as Yas1p, Yas2p and Yas3p, were demonstrated to be proteins that act as transcription factors and have potential roles in n-alkane assimilation and degradation in Y. lipolytica (Endoh-Yamagami et al., 2007; Hirakawa et al., 2009; Yamagami et al., 2004). Then, long-chain fatty alcohols are converted into the corresponding fatty aldehydes of the same chain length by long-chain-alcohol oxidase (EC 1.1.3.20) in the ER, and the corresponding fatty aldehydes are converted into the fatty acids of the same chain length by fatty-aldehyde dehydrogenase (EC 1.2.1.3) in the ER. However, short-chain fatty alcohols in the ER can be transported directly into the peroxisome and are finally converted into corresponding fatty acids of the same chain length. Meanwhile, fatty alcohols located in the ER can be converted into ω-hydroxy-fatty acids, which are precursors of corresponding dicarboxylic acid. For fatty acid metabolism, short-chain fatty acids are converted into fatty acyl-CoAs in the peroxisome by ACS II (fatty acyl-CoA synthetase II), whereas long-chain fatty acids are first converted into fatty acyl-CoAs in the cytosol by ACS I (fatty acyl-CoA synthetase I) and then are either transported into the peroxisome or used as substrates for lipid synthesis. For Y. lipolytica, at least two acyl-CoA synthetases, including ACS I in the cytosol and ACS II in the peroxisome, have been identified (Kamiryo et al., 1977). Notably, Y. lipolytica can assimilate and utilize long-chain alcohols and long-chain aldehyde as carbon substrates directly from a culture medium. However, due to their cytotoxicity, shorter-chain alcohols and aldehyde, such as decanol and decanal, cannot be assimilated. In the cytosol, long-chain fatty acids can be assimilated for lipid synthesis directly. The longer carbon-chain polyunsaturated fatty acids (PUFAs) are proposed to be synthesized via elongation and desaturation in the ER. Finally, fatty acyl-CoAs are converted into acetyl-CoA or propionyl-CoA (in the case of odd-chain fatty acids or alkanes) via β-oxidation in the peroxisome. Acetyl-CoA, which is produced from β-oxidation reaction, also can be transported into the mitochondria for cell metabolism. Generally, βoxidation is composed of four reaction steps. The first and limiting step is catalyzed by acyl-CoA oxidases (Aox, EC 1.3.3.6), which include Aox16 coded by the POX1 to POX 6 genes, respectively (Mlícková et al., 2004a, 2004b; Wang et al., 1998, 1999). The second and third steps are catalyzed by a multifunctional enzyme (MFE) coded by an MFE gene. The fourth step is catalyzed by the POT1 gene encoding a 3-ketoacylCoA thiolase (EC 2.3.1.16). Yamagami et al. (2001) demonstrated that the PAT1 gene encoding an acetoacetyl-CoA thiolase (EC 2.3.1.9) is essential for n-decane utilization in Y. lipolytica. 2.3. Safety studies on products of Y. lipolytica Metabolites produced from Y. lipolytica can be transformed into various products that can be used for humans or animals. Therefore,



performing safety tests on products of Y. lipolytica is an important and meaningful work. Holzschu et al. (1979) reported that C. lipolytica is considered non-pathogenic. Although some strains can be isolated from patients, opportunistic infections from Y. lipolytica occur very rarely. Groenewald et al. (2014) reported that opportunistic Y. lipolytica infections can be treated effectively with antifungals such as amphotericin B or may resolve themselves without any treatment. Y. lipolytica, a non-conventional yeast, has potential applications such as in the maturation of cheese and sausage or in improving food tastes due to its lipolytic and proteolytic activities (Ferreira and Viljoen, 2003; Hébert et al., 2013; Patrignani et al., 2011; Sørensena et al., 2011). Although some strains isolated from spoiled food can secrete unfavorable compounds such as brown pigments or off-flavors, no cases of adverse effects from Y. lipolytica on health has been found. The industrial use of a wild-type strain Y. lipolytica was pioneered by British Petroleum (BP) in 1957. Using n-alkanes as a carbon substrate, the SCP product called Toprina G was produced by the wild-type strain. Although the production of Toprina G was later terminated due to its high prices of raw material compared to soybean meal as an animal feed material, it was concluded that there were no negative effects of SCP on animals through longterm safety tests on the SCP products. Based on safety and efficacy studies, BP obtained authorizations to sell their products as ingredients in the diet of animals in many countries (Groenewald et al., 2014). In addition, a GRAS notice has been submitted by Baolingbao Biology Co., Ltd (Shandong, China) for erythritol, a food ingredient that is produced from mutant strain Y. lipolytica CGMCC No. 1431 via fermentation using glucose as a carbon substrate (Groenewald et al., 2014). Moreover, safety tests on products from GM Y. lipolytica have been reported. Through clinical and anatomic pathology observations in rats with repeated-dose oral studies for 28 and 90 days, no adverse effects of eicosapentaenoic acid (EPA, C22:4, ω-3) oil produced from the GM strain have been found. Therefore, the EPA oil produced from GM Y. lipolytica was supported as a safe source of oil for food supplements (Belcher et al., 2011; MacKenzie et al., 2010). Hatlen et al. (2012) reported safety tests on the biomass from GM Y. lipolytica which contained 6% EPA and 20% oil. In this research, Atlantic salmon (Salmo salar) were fed with heat-killed biomass for 95 days. Compared with the control group fed with either rapeseed oil or a mix of rapeseed and fish oil, no case of adverse effects was demonstrated. Meanwhile, the results suggested that there had been some conversion of the EPA from GM Y. lipolytica into docosapentaenoic acid (DPA, C22:5, ω-3) and docosahexaenoic acid (DHA, C22:6, ω-3) in fish. These safety tests were all financed by the DuPont Applied Bioscience Division. Based on the conclusions from these safety tests, the EPA oil and biomass from GM strains were approved as safe diet additives. Recently, DuPont submitted a GRAS notice for its EPA-rich oil derived from Y. lipolytica to the FDA and has begun selling its products on the market. In addition, Grenfell-Lee et al. (2014) showed safety tests on carotenoids that were produced by metabolic pathway modification in Y. lipolytica. No adverse effects were revealed through the genotoxicity and subchronic toxicity assays in this study. Compared with the safety profile of current commercial products, it was shown that there were no significant differences in the carotenoid production from Y. lipolytica. Recently, at Microbia (now part of DSM), several carotenoid production strains were brought to a pilot scale, and a GRAS self-affirmation was prepared for β-carotene production by Y. lipolytica (Groenewald et al., 2014). Therefore, based on these safety studies, Y. lipolytica is considered a “safe-to-use” yeast under the criteria of a decision tree (Pariza and Johnson, 2001). 3. Applications of Y. lipolytica 3.1. Secretion of proteins The wild-type Y. lipolytica strain can produce and secrete many endogenous enzymes such as alkaline protease, extracellular protease,

7

RNase, phosphatase, lipase and esterase. Moreover, Y. lipolytica is regarded as an ideal expression host for heterologous protein production such as laccase and epoxide hydrolase. Madzak and Beckerich (2013) showed in a review that various functional heterogenous proteins can be synthesized from Y. lipolytica. In this review, only recent applied examples of these functional proteins are discussed. Based on these functional applications, it is concluded that Y. lipolytica can be used as a potential cell factory for further applications. Lipase (EC 3.1.1.3), including intracellular and extracellular lipase, in contrast to esterase (EC 3.1.1.1), is an acylhydrolase that can hydrolyze TAGs into free fatty acids and glycerol. For example, Casas-Godoy et al. (2014) reported that the functional enzyme Lip2p from Y. lipolytica has the potential capability for DHA purification using ω-3 esters, and particularly DHA ethyl ester, as substrates. As an important enzyme, lipase can be used in many fields such as the food industry, bioremediation, and the production of fine chemicals and pharmaceuticals. Fickers et al. (2011) reviewed the characterization and biotechnological applications of lipases from Y. lipolytica. Naturally, lipase productivity and activity are affected by genetic coding genes and various environmental factors. From its genome sequence, the lipase family contains at least 16 different genes encoding lipase. Of the members of the lipase family, substantial information regarding lipases including Lip2p, Lip7p and Lip8p, is known (Fickers et al., 2005b, 2005c). For example, Guieysse et al. (2004) showed that the lipase Lip2p can be used as a selective catalyst to resolve a racemic mixture of 2-bromo-aryl acetic acid esters. Based on safety data, it was also revealed that the extracellular lipase Lip2p from Y. lipolytica can be used as a pharmaceutical (Turki et al., 2010b). Turki et al. (2010c) reported that the lipase Lip2p can be used as a therapeutic tool for exocrine pancreatic insufficiency treatment. Recently, Fickers et al. (2011) reported that the lipase Lip2p is under investigation by Laboratoire Mayoly Spindler, a French pharmaceutical company (http://www.mayoly-spindler.com/). Gonçalves et al. (2014) reviewed the effects of different factors (carbon source, nitrogen source, mineral elements, temperature, pH, oxygenation, water activity) on lipase and lipid production. For example, Nicaud et al. (2002) showed that the high production level of lipase (11,500 U/mL in batch and 90,500 U/mL in fed-batch, respectively) is produced by Y. lipolytica YPL280 with glucose as a carbon source. Turki et al. (2010a) reported that the production level of lipase activity (over 10,000 U/mL) is reached by Y. lipolytica LgX64.81 grown on glucose via a stepwise fedbatch strategy. Recently, lipase production with raw material as carbon substrate has received increasing interest. Using rapeseed oil as a carbon source, Kamzolova et al. (2005) reported that the highest production level of lipase activity (2760 U/mL) is produced by Y. lipolytica 704 under optimized conditions. Papanikolaou et al. (2007) presented that the solid fat tallow, which is primarily composed of saturated free-fatty acids (C18:0), can be used as a potential carbon source for lipase and biomass production in Y. lipolytica ACA-DC 50109. In this research, the selective uptake of fatty acids was observed during the yeast growth process. Mafakher et al. (2010) reported the potential lipase-production ability of two wild-type strains (Y. lipolytica M1 and M2 isolated from agroindustrial wastes). In their research, compared with carbon sources such as glycerol, kerosene, paraffin and crude oil, the maximum levels of lipase (11 and 8.3 U/mL, respectively) and the high levels of CA (27 and 8 g/L, respectively) were produced by Y. lipolytica M1 and M2 cultivated on olive oil. Kamzolova et al. (2011b) showed that a higher level of lipase activity is produced by Y. lipolytica VKM Y-2373 grown on rapeseed oil compared with that grown on glycerol and oleic acid under optimized fermentation conditions. Galvagno et al. (2011) demonstrated that the maximum production level of lipase activity at 12 U/mL is produced by Y. lipolytica NRRL Y-1095 grown on glycerol in a shake-flask culture. Darvishi et al. (2011) showed that methyl oleate, a good alternative to olive oil, can be used as a carbon substrate for lipase production. In this study, the maximum level of lipase with 356 U/mL was produced by the UV mutant Y. lipolytica U6, which was higher than


8


the production level of lipase produced from the wild-type Y. lipolytica DSM3286. Moftah et al. (2013) reported that the lipase yield of Y. lipolytica NRRL Y-1095 increased when olive oil-processing wastes such as olive mill wastewater and crude olive oil cake were used as substrates. In addition, it was demonstrated that alkaline treatment of olive oil cake appeared to remarkably improve lipase production via the solid state fermentation mode in the study. Imandi et al. (2013) reported that mustard oil cake can also be used as raw substrate for lipase production. Under optimized conditions, high lipase activity was observed in Y. lipolytica NCIM 3589 via solid state fermentation. Moreover, the LIP2 gene from Y. lipolytica was cloned and expressed in other yeasts, such as S. cerevisiae and P. pastoris. For example, Yu et al. (2007) showed that the lipase activity of 12,500 U/mL is obtained by cloning and expressing the lipase coding gene LIP2 from Y. lipolytica in P. pastoris X-33 under a fed-batch fermentation mode when methanol feeding was followed after initial glycerol culture exhaustion. Darvishi (2012) cloned the lipase coding gene LIP2 from the wild-type Y. lipolytica DSM3286 and its mutant Y. lipolytica U6, respectively, and expressed it in S. cerevisiae. Using olive oil as a carbon source, the recombinant S. cerevisiae FDS101 (with mutant LIP2 gene) could produce 1.5-fold higher levels of lipase than S. cerevisiae FDS100 (with the wild-type LIP2 gene). N-linked glycosylated protein, due to its special structure and properties, is viewed as functional protein-based therapeutics for the biopharmaceutical industry. Generally, yeasts and mammalian cells have different modification methods of glycosylation (Cereghino and Cregg, 2000). Usually, the yeast and mammalian cells share the initial stages during the N-glycosylation process. However, the glycosylation pathway of yeast cells is different from that of mammalian cells when trimming the core. Compared with the glycosylation patterns of mammalian cells and yeasts such as S. cerevisiae, the pattern of Y. lipolytica presents less hyperglycosylation, which is deemed more adapted to therapeutic protein production. Madzak and Beckerich (2013) presented a review on the glycosylation of proteins produced by “humanized” Y. lipolytica. With increasing interest, some progressive results on glycosylation of protein production have been reported. For example, Park et al. (2011a, 2011b) identified a functional gene YlMPO1 which was homologous to S. cerevisiae MNN4 and relevant to the mannosyl phosphorylation of N-linked and O-linked glycans in Y. lipolytica. However, it was indicated that YlKTR genes (YlKTR1, YlKTR2, YlKTR3 and YlKTR4) showing significant sequence homology with S. cerevisiae MNN6 homologues were not notably involved in the mannosyl phosphorylation of N-linked oligosaccharides in this research. Man3GlcNAc2, a common core of all types of eukaryotic N-glycans, was synthesized by a combinational strategy that included the removal of the ALG3 gene disruption, the overexpression of the ALG6 gene, the removal of capping glucoses and the overexpression of ER-targeted α-1,2-mannosidase (De Pourcq et al., 2012a). Meanwhile, De Pourcq et al. (2012b) reported that Man8GlcNAc2 and Man5GlcNAc2, which are deemed homogeneous human high-mannose type glycans, can also be produced by “humanized” Y. lipolytica. Moreover, glyco-engineered Y. lipolytica strains have been developed for the Pompe disease treatment (Tiels et al., 2012). Groenewald et al. (2014) showed in a review that Oxyrane, a biopharmaceutical company, has developed and optimized Y. lipolytica as a platform for the production of lysosomes of recombinant enzymes devoted to lysosomal storage disease therapy. Inulin is a linear chain of β-(2,1)-linked D-fructofuranose polymer (Pandey et al., 1999). It generally occurs in the roots and tubers of plants such as Jerusalem artichoke, dahlia tubers and chicory root. Before its assimilation, inulin must be hydrolyzed by inulinase (EC 3.2.1.7), such as exo-inulinase or endo-inulinase. Therefore, inulin, a reserve carbohydrate, is used as a renewable raw material for industrial applications. Chi et al. (2009) presented a review on inulinase-producing microorganisms and applications of inulinase. Recently,Cui et al. (2011) expressed the INU1 gene encoding exo-inulinase from Kluyveromyces marxianus CBS 6556 in Y. lipolytica SWJ-1b. In Cui's research, when 4.0% inulin was added to the medium, the crude protein in the cells and cell mass

reached 47.5% and 20.1 g/L, respectively, in engineered Y. lipolytica under optimized conditions. Moreover, using 8.0% of Jerusalem artichoke tuber as a carbon source, 53.7% crude protein and a cell mass of 20.8 g/L were produced, respectively, within 80 h. Li et al. (2012) cloned the endo-inulinase coding gene (EnIA) from Arthrobacter sp. S37, ligated it into the expression vector pINA1317 and overexpressed it in Y. lipolytica Po1h. It was reported that the production level of endoinulinase activity and the specific activity of endo-inulinase produced from the transformant 1317-EnIA were 16.7 U/mL and 93.4 U/mg, respectively. Moreover, the purified recombinant rEnIA could actively convert inulin into disaccharides under optimized conditions. Laccase (EC 1.10.3.2) is an important enzyme that carries out multicopper oxidation and has potential applications. Laccase can be naturally produced by different organisms. Y. lipolytica, due to its physiological characteristics, was used as a host for laccase expression (Madzak et al., 2005). Jolivalt et al. (2005) expressed the laccase IIIb cDNA gene from Trametes versicolor in Y. lipolytica strain Po1g. In this study, an estimated laccase yield of 2.5 mg/L was produced. Theerachat et al. (2012) revealed lcc1 cDNA gene expression, which encoded a laccase from T. versicolor DSM 11269 in Y. lipolytica JMY1212. In this study, a production level of laccase activity (0.25 U/mL) was yielded using a single-copy vector. Meanwhile, using a multi-copy vector for expression, a laccase activity of 1 U/mL was produced. In addition, compared with the wildtype strain, the mutant L185P/Q214K (rM4A) exhibited high enzyme activity and catalytic efficiency. Moreover, other functional heterologous proteins have been produced from Y. lipolytica. Linoleic acid hydroperoxide is an important substrate that can be transformed into aromatic compounds by hydroperoxide lyase. However, high concentrations of linoleic acid hydroperoxide may have toxic effects on the strain. To reduce its toxicity, Bourel et al. (2004) cloned the hydroperoxide lyase coding gene from green bell pepper and expressed it in Y. lipolytica Po1d. Using olive oil as a carbon source, the highest hydroperoxide lyase activity was obtained from the recombinant strain. Cytochrome P450 plays an important role in the oxidative and reductive metabolism. Nthangeni et al. (2004) successfully obtained the human cytochrome P450 CYP1A1 via expressing its functional coding gene in Y. lipolytica. Homoeriodictyol, one of the important flavonoid compounds with many potential applications, can be synthesized by chemical methods. However, in an alternative biotechnological pathway, it can be synthesized through the transfer of one methyl group of S-adenosyl-L-methionine to eriodictyol by 3′-Omethyltransferase (EC 2.1.1.42). To improve the production of homoeriodictyol, Liu et al. (2013a) synthesized the ROMT-9 gene encoding a 3′-O-methyltransferase from rice, cloned it into the multicopy integrative vector pINA1297 and expressed it in Y. lipolytica Po1h. Under optimized conditions, the maximal transformation ratio reached 52.4% when this methyltransferase was used as a catalyst. Ricinoleic acid (12-hydroxy-octadec-cis-9-enoic acid, C18:1-OH, RA), due to its specific chemical structure, has potential industrial applications. Beopoulos et al. (2014) developed the strategy of expressing heterologous FAH12 genes encoding hydroxylases from Ricinus communis and Claviceps purpurea to improve RA production in Y. lipolytica. To further increase the RA content, the phospholipid: diacylglycerol acyltransferase (PDAT, EC 2.3.1.158) Lro1p from Y. lipolytica was overexpressed. In this study, an RA yield of up to 43% of its total lipids accumulated in engineered Y. lipolytica. In addition, Zhao et al. (2013) successfully expressed an antibacterial peptide coding gene with important functions in Y. lipolytica SWJ-1b. Yu et al. (2013) effectively expressed an acid protease coding gene from Saccharomycopsis fibuligera A11 in the mutant Y. lipolytica strain T3-13-10. In this study, a highly specific acid protease activity of 46.7 U/mg was produced by a transformant of strain 43. 3.2. Organic acids Organic acids, such as citric acid, isocitric acid, α-ketoglutaric acid and succinic acid, are important building-block chemicals with potential



applications. Generally, organic acids with potential applications can be produced by different microorganisms, such as bacteria, molds and fungi. Finogenova et al. (2005) showed in a review that organic acids, including citric acid, isocitric acid, α-ketoglutaric acid and pyruvic acid, can be produced by Y. lipolytica. Due to its special biochemical characteristics, Y. lipolytica has received more attention for organic acid production. For example, a Y. lipolytica-based fermentation process has been used (or is still possibly being used) for citric acid production by Archer Daniels Midland (ADM) (Fickers et al., 2005a). Normally, the yield of organic acids produced by Y. lipolytica is affected primarily

9

by the genetic mechanism and various environmental factors, such as the carbon source, nitrogen source, temperature, pH, iron concentration and dissolved oxygen. Different fermentation modes also have important effects on organic acid production. A table of recent applications of organic acid produced from Y. lipolytica grown on typical carbon sources (such as glycerol, glucose, rapeseed oil, ethanol, etc.) is summarized in Table 1. Due to its chemical structure, citric acid, an important metabolic intermediate of tricarboxylic-acid cycle (TCA cycle), is of interest for industrial applications. The filamentous fungus Aspergillus niger has been

Table 1 Fermentative production of organic acids by Yarrowia lipolytica. Strains

Carbon sources

Organic acids

Production (g/L)

References

VKM Y-2373 N-1 N-1 VKM Y-2412 VKM Y-2412 VKM Y-2412 VKM Y-2412 NRRL-Y-1095 H222-S4 (p67ICL1)-T5 H181 (DSM 7806) VKM Y-2373 N15 EH59 187/1 VKM Y-2373 VKM Y-2373 704-UV4-A/NG50 H355 VKM Y-2412 VKM Y-2412 N-1 ACA-YC 5028 Wratislavia 1.31 NRRL YB-423 ACA-YC 5033 A-101-1.22 A-101-B56-5 Wratislavia 1.31 NCYC3825 JMY1203 NG40/UV7 T1 WSH-Z06 H222 WSH-Z06 WSH-Z06-RoPY2 WSH-Z06-ACL WSH-Z06 Y3314 374/4 WSH-Z06 H222-MH1 WSH-Z06-CON ACA-DC 50109 Wratislavia 1.31 NCIM 3589 LGAM S(7)1 Wratislavia 1.31 ACA-YC 5033 Wratislavia AWG7 Wratislavia AWG7 Wratislavia AWG7 H355A(FUM1-P)T4 H355(PYC1-IDP1)T5 Wratislavia K1 Wratislavia AWG7 Wratislavia AWG7 N15 NG40/UV7 LFMB 19 LFMB 20

Ethanol Ethanol Ethanol Ethanol Ethanol Ethanol Ethanol n-Paraffins Sucrose Sunflower oil Sunflower oil Sunflower oil Sunflower oil Rapeseed oil Rapeseed oil Rapeseed oil Rapeseed oil Rapeseed oil Rapeseed oil Rapeseed oil Petrolatum OMW Glucose Glycerol Glycerol Glycerol Glycerol Glycerol Glycerol Glycerol Glycerol Glycerol Glycerol Glycerol Glycerol Glycerol Glycerol Glycerol Glycerol Glycerol Glycerol Glycerol Glycerol Raw glycerol Raw glycerol Raw glycerol Raw glycerol Raw glycerol Raw glycerol Raw glycerol Raw glycerol Raw glycerol Raw glycerol Raw glycerol Pure glycerol Pure glycerol Pure glycerol Pure glycerol Pure glycerol Raw glycerol Raw glycerol

CA CA KGA KGA KGA SA SA CA CA CA CA CA ICA CA CA CA ICA KGA KGA SA CA CA CA CA CA CA CA CA CA CA CA CA KGA KGA KGA KGA KGA KGA SA PA PA PA PA CA CA CA CA CA CA CA CA ICA KGA KGA CA CA CA CA CA AA AA

105.4 120 49 172 88.7 63.4 71.7 42 140 198 68 150 93 135 77.1 22.4 86 134 102.5 69.0 217 18.9 76.4 21.6 50.1 124.2 57.15 92.8 59 57.7 115 43.3 39.2 97 66.2 62.5 56.5 66.2 45.5 61.3 16.8 66 35.1 62.5 124.5 77 35 124.5 50.1 139 197 4.3 138 186.0 89.0 82.9 157.5 98 115 29.2 10.2

Arzumanov et al. (2000) Kamzolova et al. (2003) Chernyavskaya et al. (2000) Kamzolova et al. (2012a) Kamzolova et al. (2012b) Kamzolova et al. (2009) Kamzolova et al. (2012b) Crolla and Kennedy (2004) Förster et al. (2007a) Aurich et al. (2003) Kamzolova et al. (2008) Kamzolova et al. (2008) Heretsch et al. (2008) Kamzolova et al. (2005) Kamzolova et al. (2007) Kamzolova et al. (2013) Kamzolova et al. (2013) Förster et al. (2006) Kamzolova and Morgunov (2013) Kamzolova et al. (2014a) Finogenova et al. (2005) Sarris et al. (2011) Rywińska et al. (2010b) Levinson et al. (2007) André et al. (2009) Rymowicz et al. (2010) Lazar et al. (2011) Rywińska et al. (2012) Celińska and Grajek (2013) Papanikolaou et al. (2013) Morgunov et al. (2013) Guo et al. (2014) Zhou et al. (2010) Holz et al. (2011) Yu et al. (2012) Yin et al. (2012) Zhou et al. (2012) Yu et al. (2012) Yuzbashev et al. (2010) Morgunov et al. (2004) Zhou et al. (2010) Holz et al. (2011) Zhou et al. (2012) Papanikolaou et al. (2008a) Rymowicz et al. (2006) Imandi et al. (2007) Papanikolaou et al. (2002b) Rymowicz et al. (2006) André et al. (2009) Rywińska et al. (2009) Rywińska and Rymowicz (2010) Rymowicz et al. (2006) Otto et al. (2012) Yovkova et al. (2014) Rywińska et al. (2009) Rywińska et al. (2010b) Rywińska et al. (2010a) Kamzolova et al. (2011a) Morgunov et al. (2013) Chatzifragkou et al. (2011a) Chatzifragkou et al. (2011b)

Abbreviations: OMW: olive-mill wastewater; CA: citric acid; KGA: α-ketoglutaric acid; ICA: isocitric acid; SA: succinic acid; PA: pyruvic acid; AA: acetic acid.


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used for citric acid production on a large scale. Due to its special potential, Y. lipolytica is viewed as an alternative producer to A. niger. Soccol et al. (2006) showed in a review that citric acid can be produced from various microorganisms. As early as the 1960s, it was reported that one wild-type strain, Y. lipolytica ATCC 20114, could assimilate and ferment n-paraffins as a carbon source for citric acid production (Akiyama et al., 1973). Goncalves et al. (2014) reviewed the influences of various factors, including substrates, growing conditions and culture modes, on citric acid production from wild-type or GM Y. lipolytica. To date, many different carbon substrates, such as ethanol, petrolatum, n-paraffin, glucose, glycerol, rapeseed oil, sunflower oil, olive oil and olive mill wastewater, have been used for citric acid accumulation from Y. lipolytica (Arzumanov et al., 2000; Aurich et al., 2003; Crolla and Kennedy, 2001; Darvishi et al., 2009; Fickers et al., 2005a; Finogenova et al., 2002, 2005; Kamzolova et al., 2003, 2005, 2008; Moresi, 1994; Papanikolaou et al., 2006, 2008b, 2009; Rymowicz et al., 2006; Sarris et al., 2011; Souza et al., 2014). Notably, using hydrophilic materials as carbon sources, nutrient limitation (particularly nitrogen) is considered to be an important factor for increasing citric acid production in Y. lipolytica. Under a condition of an excess of carbon (such as glucose and glycerol) and limited nitrogen condition, it will induce a rapid drop in the AMP concentration, which ultimately leads to isocitrate dehydrogenase inactivation; then, the high amount of citric acid is synthesized and transported into the cytosol. Moresi (1994) showed that the citric acid production model can be divided into three different phases (including the trophophase, the citric acid-lag phase and the citric acid-production phase) in Y. lipolytica ATCC 20346 grown on a growth medium and a production medium that contain glucose. Arzumanov et al. (2000) presented that the high production of citric acid with 105.4 g/L is achieved by mutant Y. lipolytica grown on ethanol as a carbon source in repeat-batch cultivation. Using glucose as the sole carbon substrate, 42.9 g/L of citric acid was produced by Y. lipolytica ACA-DC 50109 in nitrogen-limited cultures (Papanikolaou et al., 2006). Papanikolaou et al. (2009) reported the citric acid production from wild-type or GM strains cultivated in the presence of different glucose concentrations. Under nitrogen depletion condition, a maximum total citric acid of up to 49 g/L was produced from Y. lipolytica W29 with an initial glucose concentration of 60 g/L as a carbon source. Notably, the yeast–mycelia transition was observed in some of the strains grown in a high initial glucose concentration. However, citric acid production is not affected by nitrogen limitation in Y. lipolytica when hydrophobic materials are used as carbon sources. Kamzolova et al. (2005) reported the effect of rapeseed oil on citric acid production. In this study, under nitrogen-limited conditions when a suitable concentration of rapeseed oil was used as a fermentation substrate, a maximum citric acid concentration of 135 g/L was obtained from Y. lipolytica 187/1. Meanwhile, lipase activity of up to approximately 1200–2040 U/mL was secreted. Kamzolova et al. (2008) reported that a high citric acid concentration of 150 g/L is produced from mutant strain Y. lipolytica N15 grown on sunflower oil as a carbon source. Darvishi et al. (2009) described citric acid production in Y. lipolytica DSM3286 cultivated on olive oil as a carbon source. In this study, a citric acid concentration of 3.6 g/L was produced, and a lipase activity of 34.6 U/mL was secreted simultaneously. To improve citric acid production, in 1991, Kautola et al. (1991) reported the citric acid production from immobilized Y. lipolytica A-101. With the repeated-batch fermentation model, the alginate bead-immobilized cells were deemed the most suitable carrier for citric acid production. Meanwhile, the highest citric acid concentration of 16.4 g/L was produced with cells immobilized in κ-carrageenan beads in an air-lift bioreactor. Karasu-Yalcin et al. (2010) reported the effects of various factors, including initial pH, temperature, initial ammonium chloride concentration and initial concentration of various minerals, on the growth and citric acid production of two strains (Y. lipolytica NBRC 1658 and 57, respectively). From the comparative kinetic model of cell growth and citric acid production analysis, the maximum concentration of citric acid with 41.63 g/L was produced by Y. lipolytica 57 under optimized

conditions. In addition, chemical surfactants, such as Triton X-100, could be used to increase citric acid yield through changing the permeability of yeast cells (Mirbagheri et al., 2011). With the development of the global economy and the fossil fuel energy crisis, utilizing raw materials for citric acid production is becoming increasingly realistic. Glycerol, a byproduct of biodiesel production, has widely been used as a carbon source for citric acid production. Although raw glycerol, with its complex compositions, cannot be used as a carbon source for any products that will be used as a food or feed, many academic studies at the lab scale have been reported (André et al., 2009; Morgunov et al., 2013; Papanikolaou and Aggelis, 2003c). Among those research groups, the Papanikolaou group and Aggelis group have made some progress in citric acid production from Y. lipolytica using glycerol as a carbon source. For example, Papanikolaou et al. (2008a) reported that a total of 62.5 g/L of citric acid is produced by Y. lipolytica ACA-DC 50109 grown on a raw glycerol-based medium. Papanikolaou and Aggelis (2009) showed in a review that glycerol can be used as a potential renewable substrate for the production of different added-value chemicals by Y. lipolytica ACA-DC 50109. Moreover, the Rymowicz group has recently made considerable progress in organic acid production from Y. lipolytica using glycerol as a carbon source. Using raw glycerol as a carbon source, the highest citric acid yield of 124.5 g/L was produced from an acetate mutant of Y. lipolytica 1.31 under optimized conditions (Rymowicz et al., 2006). Rywińska et al. (2009) used raw and pure glycerol as carbon substrates for citric acid production by Y. lipolytica Wratislavia AWG7 and Wratislavia K1, respectively. In this study, the highest yield of citric acid (139 g/L) was produced by the Wratislavia AWG7 strain grown on raw glycerol as a carbon source. In contrast, 89.0 g/L of citric acid was produced by Wratislavia K1 grown on pure glycerol. Rymowicz et al. (2010) demonstrated that 124.2 g/L of citric acid is produced by Y. lipolytica A-1011.22 grown on glycerol under an optimized repeated batch fermentation culture. Rywińska and Rymowicz (2010) developed long-term repeated-batch cultures for citric acid production. In this research, the highest amount of citric acid, 197 g/L, was produced by Y. lipolytica Wratislavia AWG7 with crude glycerol as a carbon source. Rywińska et al. (2010a, 2010b) reported that citric acid can be produced by wild-type Y. lipolytica A-101 and its mutant strains (Wratislavia 1.31, Wratislavia AWG7 and Wratislavia K1) with glycerol and glucose as carbon sources in batch fermentation. It was demonstrated that Wratislavia 1.31 and Wratislavia AWG7 were the best citric acid producers with the potential of producing 82.0 g/L and 82.9 g/L, respectively, using pure glycerol as a carbon source. In further research, a higher amount of citric acid (155.2 g/L and 157.5 g/L in Y. lipolytica Wratislavia 1.31 and Wratislavia AWG7, respectively) was produced under an optimized fed-batch culture with 300 g/L glycerol as the carbon source. Rywińska et al. (2012) reported the effects of agitation and aeration on citric acid production. In this study, the highest citric acid concentration (92.8 g/L) and yield (0.63 g/g) were obtained at 0.24 vvm by acetate-negative mutants of the Y. lipolytica Wratislavia 1.31 strain grown on glycerol. In addition to glycerol, other raw materials can be fermented for citric acid production by Y. lipolytica. Imandi et al. (2008) demonstrated that pineapple waste can be used and fermented for citric acid production. Papanikolaou et al. (2008b) reported that a total of 28.9 g/L of citric acid is produced by Y. lipolytica ACA-DC 50109 cultivated on an olive mill wastewater-based medium enriched with glucose. Using the extract of Jerusalem artichoke tubers as a carbon source, a yield of citric acid up to 68.3 g/L was produced by Y. lipolytica SWJ-1b transformant 30 (L.F. Wang et al., 2013). X. Liu et al. (2014) showed that the hydrolysate of pretreated straw cellulose also can be used as a carbon source for citric acid production by Y. lipolytica SWJ1b. Under a batch fermentation mode, 26.7 g/L of citric acid was produced. Meanwhile, 42.4 g/L of citric acid was produced in fed-batch cultivation. Saygün et al. (2014) revealed that different various oils and oil industry residues (such as linseed oil, borage oil, canola oil, sesame oil, Echium oil, trout oil, olive pomace oil, hazelnut oil press cake, and



sunflower seed oil cake) also can be used as carbon sources for lipids and citric acid production from Y. lipolytica YB 423-12. With the development of genetic tools, engineered strains have been constructed by metabolic pathway modification to increase citric acid accumulation. In 1989, Nicaud et al. (1989) constructed the GM Y. lipolytica via expressing the chimeric gene for producing invertase to metabolize sucrose. Förster et al. (2007a) reported that sucrose can be utilized by cloning and expressing the SUC2 gene encoding invertase from S. cerevisiae in Y. lipolytica. Using sucrose as a carbon source, 127–140 g/L of citric acid was yielded by the recombinant strain Y. lipolytica H222-S4 (p67ICL1)-T5 under optimized conditions. Lazar et al. (2011) reported citric acid production from Y. lipolytica grown on different carbon sources, such as sucrose, glycerol, glucose or glucose with fructose (1:1). In this study, the SUC2 gene encoding invertase was expressed in Y. lipolytica, and the transformant Y. lipolytica 101-B56-5 was selected for further fermentation. Under nitrogen limitation in bioreactors, the highest value of citric acid that was produced in a batch culture ranged from 57.15 g/L (glycerol as a substrate) to 45.02 g/L (sucrose as a substrate). Using sucrose as a carbon source, the highest amount of biomass of 12.4 g/L was reached during the fermentation process. Lazar et al. (2013) successfully cloned the SUC2 gene encoding invertase and expressed it in Y. lipolytica under the pTEF promoter. One of the recombinant strains, Y. lipolytica JMY2593, reached the highest level of extracellular activity (4519 U/L) using sucrose as a carbon source. Meanwhile, these strains were capable of simultaneously producing citric acid from sucrose-based media. Liu et al. (2013b) generated a modified strain of Y. lipolytica SWJ-1b 87 that could display inulinase on its surface via the disruption of the ACL1 gene encoding ATP-citrate lyase (EC 4.1.3.8) and the expression of the ICL1 gene encoding isocitrate lyase (EC 4.1.3.1). Using inulin as a carbon source, 84.0 g/L of citric acid was produced. At the same time, iso-citric acid (1.8 g/L) was synthesized during fermentation. Papanikolaou et al. (2013) modified the Y. lipolytica strain by inactivating the PHD1 gene encoding 2-methyl-citrate dehydratase (EC 4.2.1.79). Under nitrogenlimited conditions, using raw glycerol as a carbon substrate, the mutant strain Y. lipolytica JMY1203 produced a total of 57.7 g/L of citric acid with a concomitant glycerol-to-citric-acid yield of 0.91 g/g. To modify the glycerol metabolic pathway, an effective strategy of cloning and expressing three genes that encoded glycerol dehydratase, its reactivator and a wide-spectrum alcohol oxidoreductase was developed in Y. lipolytica. Under optimized conditions, 58.8 g/L of citric acid was produced by recombinant strain Y. lipolytica NCYC3825 grown on glycerol as a carbon substrate. Simultaneously, a high biomass of 42 g/L and a peak lipid accumulation of 38% of total lipids in the dry cellular weight (DCW) were accumulated (Celińska and Grajek, 2013). Isocitric acid, a useful chiral building block, exists as four isomers. Dsthreo-Isocitric acid is one natural compound presented in living cells that has potential functions for food and medical applications. Generally, it can be synthesized in a form of a mixture that contains four isomers by chemical methods. However, the chemical synthesis method has some disadvantages. Y. lipolytica, a fermentation starter, also can produce isocitric acid during citric acid accumulation process. The ratio of isocitric acid to citric acid always reflects the yield potential and can be shifted using different strategies such as changing culture conditions and generating mutant or GM strains for a high yield of isocitric acid production. Förster et al. (2007b) observed that the overexpression of isocitrate lyase, which is encoded by the ICL1 gene, results in a significant shift in the citric acid/isocitric acid ratio. Holz et al. (2009) determined that the citric acid/isocitric acid ratio is affected by the overexpression of the ACO1 gene encoding aconitase (EC 4.2.1.3). Kamzolova et al. (2013) selected one wild-type strain, Y. lipolytica VKM Y-2373, from 60 yeast strains within different genera that could produce isocitric acid using rapeseed oil as a carbon source. Under optimized conditions, 70.6 g/L of isocitric acid was produced by the wildtype strain along with 22.4 g/L of citric acid, and the ratio of isocitric acid to citric acid was 1:0.32. At the same time, its mutant strain,

11

Y. lipolytica 704-UV4-A/NG50, was selected for isocitric acid production. Under the same conditions, 86 g/L of isocitric acid and 20 g/L of citric acid were produced with a ratio of isocitric acid to citric acid of 1:0.23. α-Ketoglutaric acid, an important metabolic intermediate with potential industrial applications, can be synthesized by isocitrate dehydrogenase in the TCA cycle. Usually, it can be synthesized by different chemical methods. However, these chemical pathways have some disadvantages, such as low yield, low purity and the production of toxic waste. Therefore, due to its inherent advantages, a biotechnological process is viewed as an alternative way to produce α-ketoglutaric acid. Otto et al. (2011) reviewed α-ketoglutaric acid production from different microorganisms. Generally, α-ketoglutaric acid can be produced in large amounts under carbon excess and thiamine limitation. Using ethanol as a carbon source, Chernyavskaya et al. (2000) reported that 49 g/L of α-ketoglutaric acid was produced by a thiamine-auxotrophic mutant Y. lipolytica N1 under optimized conditions. Recently, Zhou et al. (2010) constructed a thiamine-auxotrophic Y. lipolytica WSH-Z06 strain for α-ketoglutaric acid production. Using glycerol as a carbon source, 39.2 g/L of α-ketoglutaric acid was obtained from Y. lipolytica WSHZ06 under optimized conditions. Simultaneously, a high amount of pyruvic acid was also produced in this research. Yu et al. (2012) demonstrated that 66.2 g/L of α-ketoglutaric acid accumulated in Y. lipolytica WSH-Z06 under optimized conditions with glycerol as the sole carbon source. Moreover, the highest amount of α-ketoglutaric acid of 172 g/L was produced, and a mass yield coefficient of 0.70 g/g was obtained by Y. lipolytica VKM Y-2412 grown on ethanol as a carbon substrate under optimized conditions (Kamzolova et al., 2012a). Kamzolova and Morgunov (2013) isolated an ideal strain, Y. lipolytica VKM Y-2412, from 26 α-ketoglutaric acid-producing strains with rapeseed oil as a carbon source. Under optimized conditions, a yield of αketoglutaric acid of up to 102.5 g/L with a mass yield coefficient of 0.95 g/g and a volumetric α-ketoglutaric acid productivity of 0.8 g/L/h was ultimately produced by Y. lipolytica VKM Y-2412. Pyruvic acid is produced as a main byproduct during the αketoglutaric acid-accumulation process. Li et al. (2001) showed in a review that pyruvic acid can be produced by various eukaryotic and prokaryotic microorganisms (including Y. lipolytica). Morgunov et al. (2004) reported that a high pyruvic acid concentration with 61.3 g/L is produced by the thiamine-auxotrophic strain Y. lipolytica 374/4 grown on glycerol via a changing thiamine concentration. However, it was reported that high concentrations of pyruvic acid could result in limiting the effects on α-ketoglutaric acid application. To enhance the α-ketoglutaric acid yield and reduce the pyruvic acid yield, a genetic approach of modifying the target product metabolic flux was developed. Holz et al. (2011) reported the influence of the overexpression of multiple complex genes (KGD1, KGD2 and LPD1, which encode αketoglutarate dehydrogenase (EC 1.2.4.2)) on α-ketoglutaric acid production in wild-type Y. lipolytica H222. In this research, the amounts of citric acid and isocitric acid did not change significantly, whereas the production of α-ketoglutaric acid was reduced, and the production of pyruvic acid was elevated in recombinant Y. lipolytica H222-MHI under optimized conditions. To improve the α-ketoglutaric acid yield, Zhou et al. (2012) regulated the carbon flux through expressing the acetyl-CoA synthetase (EC 6.2.1.1) coding gene, ACS1, from S. cerevisiae and the ATP-citrate lyase coding gene, ACL, from Mus musculus in Y. lipolytica WSH-Z06. In this research, a high yield of αketoglutaric acid, up to 56.5 g/L, was obtained in Y. lipolytica grown on glycerol via a two-stage control strategy. However, an obvious decrease in pyruvic acid accumulation (from 35.1 g/L to 20.2 g/L) was obtained. To increase α-ketoglutaric acid production, two genes, ScPYC1 from S. cerevisiae and RoPYC2 from Rhizopus oryzae encoding pyruvate carboxylase (EC 6.4.1.1), were cloned and expressed in Y. lipolytica WSHZ06. Under optimized conditions with glycerol as the carbon substrate, the recombinant strains Y. lipolytica WSH-Z06-ScPYC1 and WSH-Z06RoPYC2 produced α-ketoglutaric acid levels of up to 53.6 g/L and 62.5 g/L, respectively (Yin et al., 2012). To decrease pyruvic acid


12


limitation, Otto et al. (2012) reported that the overexpression of the FUM1 gene, encoding fumarase (EC 4.2.1.21), and the PYC1 gene, encoding pyruvate carboxylase, has positive effects on reducing the concentrations of pyruvic acid and fumarate in Y. lipolytica. In this research, the recombinant strains, Y. lipolytica H355 (PYC1) T3, Y. lipolytica H355 (FUM1) T1 and Y. lipolytica H355 (FUM1-PYC1) T4, were constructed, and the highest amount of α-ketoglutaric acid of 138 g/L was produced in Y. lipolytica H355 (FUM1-PYC1) T4 grown on raw glycerol as the carbon source. Yovkova et al. (2014) developed a biotechnological process with overexpression of the gene IDP1, encoding isocitrate dehydrogenase (EC 1.1.1.42), and PYC1, encoding pyruvate carboxylase, in Y. lipolytica to increase α-ketoglutaric acid production. In this research, under optimized conditions, the recombinant strain Y. lipolytica H355 (PYC1-IDP1) T5 produced the highest level of α-ketoglutaric acid of 186 g/L when raw glycerol was used as the carbon source in a bioreactor. In addition, Guo et al. (2014) demonstrated the effect of the overexpression of the pyruvate dehydrogenase (EC 1.2.4.1) complex, including the α and β subunits of the E1 component, E2 and E3 components, on α-ketoglutaric acid production from Y. lipolytica. Under optimized conditions, it was revealed that a high yield of α-ketoglutaric acid (up to 43.3 g/L) was obtained via overexpressing the PDA1 gene and encoding the α unit of the E1 component in Y. lipolytica WSH-Z06 T1 grown on glycerol. Succinic acid is a building-block chemical that is widely used in various fields. Compared with the chemical-synthesis route, the biosynthesis method with many potential advantages has received more attention for producing succinic acid. Commonly, succinic acid can be produced by bacteria. Recently, Kamzolova et al. (2009) developed an efficient two-step process that includes the synthesis of α-ketoglutaric acid by Y. lipolytica and a chemical decarboxylation treatment for succinic acid production. Using ethanol as a carbon source, a yield of succinic acid of up to 63.4 g/L was produced by Y. lipolytica VKM Y2412. With the two-step process, a succinic acid yield of 69.0 g/L was produced by Y. lipolytica grown on rapeseed oil via the decarboxylation of α-ketoglutaric acid by hydrogen peroxide (Kamzolova et al., 2014a). Kamzolova et al. (2014b) obtained a high yield of succinic acid of up to 71.7 g/L with ethanol as a promising substrate, and reported that succinic acid has the potential for antibacterial and nematicidic activities to control pathogenic microorganisms in this study. Through a temperature-sensitive mutation strategy and the deletion of genes encoding succinate dehydrogenase (EC 1.3.5.1), the strain Y. lipolytica Y3314 was constructed and a succinate level of 45.5 g/L was produced under optimized conditions (Yuzbashev et al., 2010). Moreover, acetic acid, an important metabolic product with potential applications, could be produced as byproduct in Y. lipolytica. Papanikolaou et al. (2009) revealed that acetic acid can be produced by some natural Y. lipolytica strains under nitrogen-limited conditions with initial glucose (60 g/L) as a carbon source. Moreover, Chatzifragkou et al. (2011a) reported that acetic acid can be produced by Y. lipolytica LFMB 19 under different initial raw glycerol concentrations. In particular, at the initial raw glycerol concentration of 90 g/L, the highest production level of acetic acid (up to 29.2 g/L) was accumulated in this research. Chatzifragkou et al. (2011b) showed the effects of adding various essential oils of Origanum vulgare L. upon the biochemical ability of Y. lipolytica LFMB 20. It was revealed that the production of both organic acids such as citric acid and acetic acid, and mannitol were reduced significantly in Y. lipolytica LFMB 20 grown on higher essential oil concentrations in this research. Itaconic acid is one of the top twelve building-block chemicals and has wide applications. It can be naturally synthesized by chemical methods. However, a biotechnology process has attracted more interest for itaconic acid production. Aspergillus terreus is a well-known producer of itaconic acid. Compared to A. terreus, Y. lipolytica, due to its special potential, is viewed as an alternative host for itaconic acid production. The cis-aconitic acid decarboxylase (EC 4.1.1.6) can effectively catalyze the transformation of cis-aconitate into itaconate. Wang et al. (2012a)

discovered that a high level of itaconic acid can be produced from GM Y. lipolytica via expressing cis-aconitic acid decarboxylase when this strain is grown on glycerol as a carbon source. Long-chain dicarboxylic acids, as building blocks, are important raw materials in the fine chemical industry and can be used for the synthesis of various materials, such as perfumes, polymers, adhesives and macrolide antibiotics. Due to some drawbacks of the chemical synthesis pathway, an alternative bioconversion strategy has been developed for long-chain dicarboxylic acid synthesis with hydrophobic materials such as alkanes as substrates. Generally, dicarboxylic acid synthesis, corresponding to alkane degradation through the alkane monooxygenase system in the ER, occurs via the ω-oxidation pathway. However, dicarboxylic acid can be degraded through the β-oxidation pathway. Therefore, a high yield of long-chain dicarboxylic acids can be obtained through the modification of its synthesis and degradation pathways. Smit et al. (2005) reported long-chain dicarboxylic acid production from wild-type Y. lipolytica W29 and its POX-deleted strains (double, triple and quadruple deletion mutants). In this research, it was revealed that dodecanedioic acid could be synthesized in all strains. Among these strains, the quadruple deletion mutant strain was the only strain that was able to produce dicarboxylic acid from C16 alkanols and monocarboxylic acid. Moreover, Thevenieau (2006) reported that overexpression of the cytochrome P450 reductase coding gene also results in a twofold increase in long chain dicarboxylic acid production. 3.3. Single cell oils Single cell oils (SCOs), also called microbial lipids, are produced from oleaginous microorganisms. Due to their similar compositions to plant oils and fats, SCOs are viewed as alternative edible oils with potential applications. Generally, those microorganisms that can accumulate lipid to more than approximately 20% of their biomass are termed the oleaginous species (Ratledge and Wynn, 2002). Papanikolaou and Aggelis (2011a) summarized in detail the mechanism of microbial lipid accumulation (de novo pathway from hydrophilic substrates and ex novo pathway from hydrophobic substrates) and the degradation of oleaginous yeasts. Meanwhile, Papanikolaou and Aggelis (2011b) reported various kinetic models of microbial lipid accumulation in oleaginous yeasts grown on hydrophilic and hydrophobic substrates. In particular, the review presented different strategies for the production of cocoa butter-like lipids from oleaginous yeasts. Naturally, these oleaginous species include yeasts, molds and algae. To date, fewer than 30 of the 600 species are known to be oleaginous (Ratledge, 1994). Ageitos et al. (2011) reviewed the synthesis pathways of microbial oils and their metabolic characteristics in oily yeasts. These oleaginous yeasts primarily include Lipomyces starkeyi, Cryptococcus curvatus, Cryptococcus albidus, Rhodotorula glutinis, Rhodosporidium toruloides, Trichosporon pullulans and Y. lipolytica (Li et al., 2008). Naturally, only some restricted Y. lipolytica strains have the potential to produce lipids to levels exceeding 20% of cell dry weight. Ratledge and Wynn (2002) reported that the microbial lipid content of Y. lipolytica can reach 36% of dry cell biomass. Tsigie et al. (2011) showed that Y. lipolytica Po1g is able to accumulate high lipid content (58.5%) exceeding the value of 20% of cell dry weight when detoxified sugarcane bagasse hydrolysate is used as a carbon substrate. Nambou et al. (2014) reported that a lipid content up to 35% of the microbial dry biomass can be produced by Y. lipolytica DSM3286 grown on the basal minimal culture medium (S2). Therefore, Y. lipolytica, a non-conventional yeast, has received increasing attention. Recently, Abghari and Chen (2014) critically presented a review of the potential applications of Y. lipolytica used as an oleaginous cell factory platform. The metabolism of microbial oils includes fatty acid synthesis and triacylglycerol production. In general, SCOs play an important role in the supply of major PUFAs, which have potential roles in nutrition and human health. Naturally, fatty acids (C14 and/or C16) can be synthesized via a de novo pathway from hydrophilic materials or an ex novo pathway



directly from a medium containing hydrophobic materials. However, long-chain PUFAs are synthesized by elongation and/or a desaturation reaction in the ER. Generally, PUFAs, including γ-linolenic acid (GLA, C18:3, ω-6), arachidonic acid (ARA, C20:4, ω-6), EPA and DHA, are classified as either ω-3 fatty acids or ω-6 fatty acids. Janssen and Kiliaan (2014) showed in a review that ω-3 and ω-6 fatty acids have important effects on neural development, aging and neurodegeneration. Generally, the major sources of ω-6 fatty acids and ω-3 fatty acids can be obtained from various foods. Mammals, including humans, cannot synthesize linoleic acid (LA, C18:2, ω-6) or α-linolenic acid (ALA, C18:3, ω-3). Therefore, these functional PUFAs should be obtained from the diet directly or synthesized indirectly in human body via fatty acid synthesis reaction with appropriate ω-3 and ω-6 precursors as substrates. However, with the growing population and the increasing consumption of edible oils, the production of SCOs is becoming more important. Our group has made successful progress in the production of functional SCOs, including ARA-rich oil and DHA-rich oil, by fungal Mortierella alpina and Schizochytrium sp., respectively (Ji et al., 2014a, 2014b; Jin et al., 2008; Nie et al., 2014; Qu et al., 2013; Ren et al., 2009, 2014; Sun et al., 2014). A commercial scale of DHA production and a pilot scale of ARA production have been developed by Jiangsu Tiankai Biotechnology Co., Ltd. Recently, based on genetic technology and metabolic pathway modification, various functional PUFAs have been synthesized in different organisms such as P. pastoris (Wan et al., 2009), Nicotiana benthamiana (Petrie et al., 2010), Camelina sativa (Ruiz-Lopez et al., 2014), Phaeodactylum tricornutum (Hamilton et al., 2014), Arabidopis thaliana (Petrie et al., 2012a, 2012b; Qi et al., 2004) and Brassica napus (Petrie et al., 2012a). Due to its high content of LA (C18:2 of 51%), Y. lipolytica, a non-conventional yeast, is used as a suitable host for ω-3/ω-6 fatty acid production via the construction of the target product metabolic pathway. To date, there are many studies on PUFA synthesis by genetically engineered Y. lipolytica. Chuang et al. (2010) showed that the approach of the simultaneous coexpression of Δ6-desaturase and Δ12desaturase from M. alpina is applied in Y. lipolytica Po1g to increase the yield of GLA. Conjugated linoleic acid (CLA), an important compound with wide applications, can be synthesized via expressing LA isomerase with endogenous LA as a substrate. B. Zhang et al. (2012) reported the production of trans-10, cis-12-CLA via expressing the gene encoding LA isomerase from Propionibacterium acnes in Y. lipolytica Polh grown on glucose as a carbon source. Moreover, Dupont de Nemours developed a series of engineered Y. lipolytica strains for producing various ω-3 and ω-6 fatty acids, such as ARA, EPA and DHA (Damude et al., 2009a, 2009b, 2011). With the power of metabolic engineering, Xue et al. (2013) successfully showed that EPA-rich oils can be produced by engineered Y. lipolytica. To produce EPA from LA used as an inherent substrate, the complete EPA biosynthetic pathway, including 22 heterologous genes encoding different functional enzymes (Δ9elongase, Δ8-desaturase, Δ5-desaturase and Δ17-desaturase, respectively) from various long-chain PUFA-producing microorganisms, was integrated into Y. lipolytica ATCC 20362. Meanwhile, the combinational strategy including the overexpression and deletion of functional genes involved in lipid synthesis was applied in modulating the product carbon flux. Notably, these genes such as PEX10, LIP1 and LEU2 were disrupted by integration events with a non-homologous end-joining recombination. Therefore, based on the expertise in metabolic engineering and serendipitous identification, it was determined that EPA at 56.6% of the total fatty acids and lipids at up to 30% of the DCW were accumulated by Y. lipolytica Y4305 grown on glucose as a carbon source. Lipid bodies, which mainly consist of neutral lipids in the form of triacylglycerols, diacylglycerols or steryl esters, are covered by a monolayer of amphipathic phospholipids, glycolipids and/or sterols (Murphy, 2001). As the main energy storage product, triacylglycerols are always synthesized via the Kennedy pathway, which uses glycerol-3-phosphate (G-3-P) and acyl-CoA as substrates (Kennedy, 1961). The synthesis pathways and analysis methods of microbial oils from various

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oleaginous species have been reviewed (Beopoulos et al., 2009a, 2009b, 2011; Guschina and Harwood, 2006; Ratledge and Wynn, 2002; Schmidt et al., 2013). Naturally, two different pathways for lipid accumulation exist in oily microorganisms. One pathway is a de novo lipid synthesis pathway, and the other is an ex novo pathway. Papanikolaou and Aggelis (2011a) presented a significant review on lipid accumulation and degradation of oleaginous yeasts. Generally, when using water-soluble hydrophilic materials, such as glucose or glycerol, as carbon sources, microbial lipids are synthesized via the de novo lipid synthesis pathway after exhaustion of certain nutrients (such as nitrogen). In particular, under nitrogen-limited condition, the activation of AMP deaminase (EC 3.5.4.6) results in a rapid decrease in the AMP concentration, which ultimately leads to isocitrate dehydrogenase inactivation. Then, citric acid is transported into the cytosol from the mitochondria and is transformed into acetyl-CoA and oxaloacetate in the cytosol by ATP-citrate lyase, distinguishing this process from citric acid metabolism in non-oleaginous yeast. Next, acetyl-CoA is converted into malonyl-CoA by acyl-CoA carboxylase (EC 6.4.1.2) in the cytosol. In Y. lipolytica, ATP-citrate lyase consists of two subunits, which are encoded by ACL1 and ACL2. The pool of acetyl-CoA and malonyl-CoA plays an important role in the fatty acid synthesis process in oleaginous yeast. Naturally, when using a hydrophilic substrate-based nitrogenlimited medium as a carbon source, the growth of Y. lipolytica can be divided into distinct phases for microbial lipid production (including a biomass formation phase, a lipid production phase and a citric acid production phase). There is a strong interplay of microbial lipids and citric acid production in Y. lipolytica cultivated on hydrophilic substrates (such as glucose or glycerol) under nitrogen-limited conditions. However, when using hydrophobic materials as carbon substrates (such as fats and oils), microbial lipids are accumulated via an ex novo synthesis pathway and are not affected by nutrient exhaustion in Y. lipolytica. In 1984, Bati et al. (1984) showed that citric acid and microbial lipids could be produced simultaneously by C. lipolytica grown on corn oil as a carbon source. Moreover, their research revealed that the low level of dissolved oxygen played an important role in lipid production. Montet et al. (1985) reported that hydrophobic materials, including rapeseed oil, palm oil and soapstock, can also be used as carbon substrates by C. lipolytica YB 423-12 to increase lipid production. Koritala et al. (1987) showed that soybean oil can also be consumed by C. lipolytica Y-1095 for microbial lipid production. Papanikolaou et al. (2001) reported the effects of using mixtures of industrial fats containing stearic, oleic, linoleic and palmitic acid on the lipid accumulation and cellular lipid composition of Y. lipolytica LGAM S(7)1. Interestingly, a cocoa-butter substitute could be produced with a 50/50 mixture of hydrolyzed oleic rapeseed oil/stearin. Generally, the characteristics of various Y. lipolytica strains isolated from different environments are straindependent. Montet et al. (1985) showed that only 2.0% of the total lipid content was produced by C. lipolytica YB 423-12 grown on glucose. Papanikolaou et al. (2009) showed the effects of different initial glucose concentrations (30 g/L and 60 g/L) on the production of organic acids and microbial lipids, respectively, by various Y. lipolytica strains. It was revealed that citric acid, with a maximum level of 38.2–49.0 g/L, was mainly produced under nitrogen depletion conditions, whereas intracellular lipids were not produced in high amounts, i.e., less than 14%. Makri et al. (2010) reported that microbial lipid production is accompanied by citric acid production in repeated batch culture by Y. lipolytica ACA-DC 50109 grown on pure glycerol as a carbon source. This research showed that the amount of lipids exceeded 20% during lipogenic production. Moreover, it was observed that neutral lipid degradation occurred in Y. lipolytica when the rate of uptake of the carbon substrate was low. Naturally, the accumulation of microbial lipids is accompanied by lipase synthesis in Y. lipolytica grown on TAGs-rich materials. Papanikolaou et al. (2007) presented a numerical model for researching microbial lipids, biomass and extra-cellular lipase production by Y. lipolytica ACA-DC 50109 with stearin as the sole carbon source. It was revealed that microbial lipids and lipase could be produced


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simultaneously. Najjar et al. (2011) also revealed that olive oil can be utilized as a carbon substrate by Y. lipolytica CBS 7504 to trigger lipase production. In general, lipid accumulation is affected by different factors. Tehlivets et al. (2007) concluded that building blocks such as ATP, NADPH and CoA are co-factors that play important roles during the process of lipid synthesis. Ratledge (2014) reported that the NADPH generation mechanism still remains unclear. In that review, it was suggested that NADPH might be produced from different pathways, such as through the action of malic enzyme (EC 1.1.1.40), the pentose phosphate pathway and cytosolic isocitrate dehydrogenase activity. Based on genome sequence information, there is only one malic enzymecoding gene located in the mitochondrion of Y. lipolytica. However, its homologous overexpression did not significantly improve lipid accumulation (Beopoulos et al., 2011). Moreover, Zhang et al. (2013) demonstrated that heterologous overexpression of the gene encoding malic enzyme from M. alpina does not significantly result in lipid biosynthesis in Y. lipolytica Po1h. It was proposed that NADPH might be primarily provided via the pentose phosphate pathway in Y. lipolytica. Hong et al. (2011b) successfully increased lipid accumulation and PUFA production in Y. lipolytica via overexpression of the genes encoding glucose 6-phosphate dehydrogenase (EC 1.1.1.49) and 6phosphogluconolactonase (EC 3.1.1.31), which are involved in the pentose phosphate pathway. Recently, Wasylenko et al. (2015) suggested that the oxidative pentose phosphate pathway is the primary source of NADPH for lipogenesis in Y. lipolytica. Generally, the final step of triacylglycerol synthesis is catalyzed by diacylglycerol acyltransferase which contains distinct enzymes, including the phospholipid diacylglycerol acyltransferase (PDAT, EC 2.3.1.158), acyl-CoA: diacylglycerol acyltransferase (DGAT, EC 2.3.1.20). Through gene disruption of the pdat, gat1 and gat2 genes encoding diacylglycerol acyltransferase, H. Zhang et al. (2012) determined that DGAT1p significantly contributes to lipid biosynthesis in Y. lipolytica ATCC 90812. Moreover, Beopoulos et al. (2012) determined that four acyltransferase-encoding genes (LRO1, ARE1, DGA1 and DGA2) play important roles in triacylglycerol synthesis. Furthermore, many other environmental factors also play important roles in lipid accumulation. Papanikolaou and Aggelis (2011b) summarized the effects of various factors, such as carbon source, nitrogen source, C/N ratio, pH, incubation temperature and various yeast strains and fermentation modes, on the accumulation of lipid from oleaginous microorganisms (including Y. lipolytica). The Aggelis and Papanikolaou group has presented many results on the lipid production of Y. lipolytica LGAM S(7)1 grown on various carbon sources. For example, to increase the production of cocoa-butter substitute, Papanikolaou et al. (2001) used industrial fats containing mixtures of stearic, oleic, linoleic and palmitic acids for lipid production. In that research, it was revealed that the high lipid accumulation with a lipid composition similar to that of cocoa-butter occurred in Y. lipolytica LGAM S(7)1 grown on a 50/50 mixture of hydrolyzed oleic rapeseed oil and stearin. Papanikolaou et al. (2003) reported the effects of using the mixtures of stearin, raw glycerol, and glucose as carbon sources on the production of a cocoa-butter-like lipid in Y. lipolytica ACA-DC 50109. It was revealed that the maximum content of storage lipids similar to cocoa-butter (up to 3.4 g/L) was produced by Y. lipolytica ACA-DC 50109 grown on mixtures of stearin and glycerol. Naturally, the size and composition of the lipid body are affected by various carbon sources. Athenstaedt et al. (2006) reported that the number of polypeptides of the lipid body increased when Y. lipolytica W29 cells were shifted from glucose to oleic acid. Sestric et al. (2014) showed that Y. lipolytica ATCC 20460 serves as an effective producer for TAG synthesis when using different carbon sources such as pure glycerol, raw glycerol, dextrose and canola oil. Moreover, it was revealed that Y. lipolytica displayed a preference for the consumption of glycerol over dextrose. Interestingly, a large amount of citric acid was produced by Y. lipolytica grown on glucose as the sole carbon source (Papanikolaou et al., 2006). To increase lipid production without excess citric acid, Ochoa-Estopier and Guillouet (2014)

successfully developed a cultivation system by controlling the C/N ratio for modification of the metabolic flux in Y. lipolytica W29 grown on glucose. However, the degradation mechanisms of microbial lipids produced by Y. lipolytica are still entirely unclear. Beopoulos et al. (2011) presented a review on lipid catabolism in yeasts. Generally, the carbon pool used for cell growth and metabolism could not be provided solely by the extra-cellular carbon source and needed to be supplemented by biodegradation of the storage lipid. Another hypothesis of lipid catabolism was that when significant amounts of lipid were accumulated inside the yeast cells, the yeast could initiate consumption of these reserve lipids instead of transporting the remaining extra-cellular aliphatic chains inside the yeast cell. Moreover, the lipid degradation could be related to decreased membrane biological activity during the later phase of fermentation. Normally, the intra-cellular lipids, such as neutral lipids and polar lipids, are degraded into various chemicals used for meeting energy and carbon source requirements. Generally, triacylglycerols are hydrolyzed into fatty acids and glycerol by lipase. A numerical model of Y. lipolytica grown on vegetable oils was developed for the prediction of lipid accumulation and degradation in oleaginous microorganisms (Aggelis and Sourdis, 1997). In this research, it was reported that the degradation of the intra-cellular carbon pool was a very slow process. Papanikolaou et al. (2001) presented the assumptions for explaining the intra-cellular lipid degradation of Y. lipolytica LGAM S(7)1. Papanikolaou and Aggelis (2003a) developed a modeling approach for researching the kinetic behavior of Y. lipolytica ACA-DC 50109 capable of producing lipids using industrial fats as carbon sources. It was revealed that the degradation of intra-cellular storage lipids was significantly lower compared with that of the extra-cellular carbon substrates. Papanikolaou and Aggelis (2003b) reported that yeast preferentially consumes fatty acids such as C16:0 and C18:1 during the degradation phase of lipid storage. With the development of genetic technology, the traditional strategy of modulating the metabolic carbon flux has been developed for increasing lipid accumulation. Courchesne et al. (2009) reviewed different strategies (including biochemical engineering, genetic engineering and transcription factor engineering) for improving lipid production in oilrich microalgae. Liang and Jiang (2013) presented various metabolic engineering approaches for enhancing microbial oil production in different oleaginous microorganisms. Mlícková et al. (2004a) reported the effects of POX gene deletions on lipid accumulation and showed that the accumulation of lipids is strongly affected by the Aox proteins. In particular, Aox2p had an important role in increasing the amount of neural lipids. The pool of G-3-P, which plays an important role in lipid accumulation, can be modulated through GPD1 overexpression or GUT2 inactivation. Deactivation of the GUT2 gene that resulted in the enhancement of the G-3-P level, in combination with deletion of the POX1 to POX6 genes, led to a significant increase in lipid yield (Beopoulos et al., 2008). To increase lipid accumulation, the metabolic strategy of improving the G-3-P level through GPD1 gene overexpression and GUT2 gene inactivation and blocking the β-oxidation pathway by deletion of the POX1 to POX6 genes and the MFE1 gene was developed. It was revealed that modification of the metabolic pathway could affect the expression of genes involved in triacylglycerol homeostasis (Dulermo and Nicaud, 2011). Z.P. Wang et al. (2013) showed the effect of disruption of the MIG1 gene, an important repressor element in the glucose repression process, on microbial lipid metabolism. Compared with wild-type yeast, a high lipid content was produced by the mutant strain Y. lipolytica M25 in this research. Seip et al. (2013) demonstrated that all of the subunits of the SNF1/AMPK complex play important roles in lipid accumulation in Y. lipolytica. Among these components, deletion of SNF1p significantly increased lipid production. Tai and Stephanopoulos (2013) demonstrated that overexpression of ACC1 and DGA1, which are functional genes involved in lipid synthesis, has a significant effect on lipid accumulation in Y. lipolytica. Overexpression of ACC1 and DGA1 induced a significant flux diversion towards lipid



synthesis. Moreover, Dulermo et al. (2013) revealed that the inactivation of the TGL3 and TGL4 encoding intra-cellular lipases involved in triacylglycerol degradation can increase lipid accumulation in Y. lipolytica. Blazeck et al. (2014) described an efficient strategy for increasing lipid production via overexpression of five functional lipogenic genes (MAE, AMPD, ACL, DGA1 and DGA2) in combination with the deletion of MFE1 and PEX10. Under optimized conditions, lipids exceeding 25 g/L accumulated in the engineered Y. lipolytica. Recently, Qiao et al. (2015) constructed an engineered Y. lipolytica capable of producing high lipid titers of 55 g/L, which was the highest reported lipid titer for Y. lipolytica, via simultaneous overexpression of delta-9 stearoyl-CoA desaturase, acetylCoA carboxylase and diacylglyceride acyl-transferase in Y. lipolytica Po1g. Moreover, it was suggested that autophagy was a necessary but not sufficient element of high triacylglycerol accumulation. Additionally, Liu et al. (2015) presented a rapid evolutionary metabolic approach for improving lipogenesis in a pre-engineered Y. lipolytica via random mutagenesis and reported that high lipid production titers of 39.1 g/L were achieved in the ethyl methanesulfonate-evolved strain, Y. lipolytica E26E1. More interestingly, it was revealed through whole genome sequencing that there was a link between gamma-aminobutyric acid (GABA) assimilation and lipogenesis in Y. lipolytica. The use of raw materials as carbon sources for SCO production has attracted a lot of attention for the purpose of reducing high production costs and serious environment pollution. Huang et al. (2013) reviewed the utilization of raw materials for SCO production by wild-type or modified Y. lipolytica strains. Aggelis and Komaitis (1999) reported that using aqueous extracts of Teucrium polium L. can enhance SCO production by Y. lipolytica LGAM (7)1. Karanika et al. (2001) concluded that aqueous extracts of some plants of the Lamicaeae family could also increase the biomass and ratio of unsaturated to saturated fatty acids from Y. lipolytica LGAM (7)1 grown on medium with a low C/N ratio. However, Papanikolaou et al. (2008c) reported that the addition of essential oil of the Greek citrus hybrid Citrus aurantium significantly inhibits the growth of Y. lipolytica ACA-DC 50109. Moreover, Chatzifragkou et al. (2011b) revealed that higher O. vulgare L. essential oil concentrations have the same impact on the growth of Y. lipolytica LFMB 20. Moreover, the amounts of cellular saturated fatty acids, especially palmitic acid, were increased with increasing essential oil concentrations. Using industrial glycerol as a carbon substrate, reserve lipids of 3.5 g/L were produced by Y. lipolytica LGAM (7)1, while citric acid was produced under a nitrogen-limited continuous culture (Papanikolaou and Aggelis, 2002). Using stearin as the carbon source, a dry biomass of 9–12 g/L and lipids were produced under optimized conditions by Y. lipolytica ACA-DC 50109 (Papanikolaou et al., 2002a). Papanikolaou et al. (2003) demonstrated that microbial lipids can be produced in a medium containing mixtures of stearin, glycerol, and glucose as co-substrates. Papanikolaou and Aggelis (2003a) reported the biomass and lipid synthesis of Y. lipolytica ACA-DC 50109 cultivated on a mixture of hydrolyzed oleic rapeseed oil and stearin. In that research, lipids were accumulated under high stearic acid concentrations. Papanikolaou and Aggelis (2010) reported that Y. lipolytica has a particular potential capability for lipid production when using various fatty agro-industrial residues, such as stearin and oleic acid-rich materials, as carbon sources. Chi et al. (2011) determined that Y. lipolytica can assimilate and ferment various wastes such as food waste hydrolyzed broth and municipal wastewater with high concentrations of residue carbon sources and chemical oxygen demand (COD) for lipid production. Yu et al. (2011) demonstrated that hydrolysate from the dilute sulfuric acid pretreatment of wheat straw can be used for microbial oil production by Y. lipolytica. Interestingly, it was demonstrated that hemicellulosic sugars obtained from the pretreatment of lignocellulosic materials have potential applications for lipid production. Using detoxified sugarcane bagasse hydrolysate as a carbon source, 6.68 g/L of lipids and 11.42 g/L of biomass were produced by Y. lipolytica Po1g under optimized conditions (Tsigie et al., 2011). Katre et al. (2012) selected five strains with potential for lipid and biomass production by Y. lipolytica grown on medium containing glucose and wastes such as

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cooking oil as carbon sources. In that research, compared with other strains, Y. lipolytica NCIM3589 exhibited a maximal lipid/biomass coefficient (0.29 g/g grown on glucose and 0.43 g/g grown on cooking oil). Moreover, a dry cell mass of 10.75 g/L and a lipid content of 48.02% were produced from Y. lipolytica Po1g under optimized conditions with detoxified defatted rice bran hydrolysate (DRBH) as a carbon source (Tsigie et al., 2012b). Tsigie et al. (2012a) adopted a sub-critical water treatment for increasing the neutral lipid accumulation. Under optimized conditions, a maximum of 42.69% (w/w) of neutral lipids was obtained from Y. lipolytica Po1g grown on DRBH. Wadekar et al. (2012) reported that non-traditional oils, such as jatropha and karanja oils, which are rich in oleic and linoleic acids, can also be used as potential substrates for sophorolipid production. Harder et al. (2013) concluded that the culture medium containing bamboo, vinasse, sewage and yeast extract can be used for higher lipid production by Y. lipolytica. By using crude coconut fat as a carbon source, medium-chain saturated fatty acids, especially lauric acid (62% of total fatty acids), were produced by Y. lipolytica RO13 in a solid-state cultivation system (Parfene et al., 2013). Juszczyk et al. (2013) determined that the Y. lipolytica S6 strain, one of 21 strains isolated from different environments, can produce a high biomass when using glycerol as a carbon source under optimized conditions. In that research, high contents of lysine, threonine and phenylalanine/tyrosine were also produced. Using crude glycerin as a carbon source, 2.7 g/L cellular lipids were produced by Y. lipolytica IMUFRJ50678 under optimized conditions (de Lima et al., 2013). Cheirsilp and Louhasakul (2013) reported that many industrial wastes, such as crude glycerol, palm oil mill effluent, serum latex and molasses, can be used as potential fermentation sources for cell growth and microbial lipid production by Y. lipolytica. Poli et al. (2014) demonstrated that combined wastes containing crude glycerol and fresh yeast extract from brewery waste as the carbon and nitrogen sources, respectively, have promising effects on the fatty acid composition of microbial oil, with a high PUFA content produced by Y. lipolytica QU21. Moreover, Tsigie et al. (2013) reported that defatted biomass from Y. lipolytica Po1g can be used as a substrate for ethanol production. 3.4. Bioremediation With economic development and energy consumption, environments have been polluted by various materials such as heavy metals, toxic carbon compounds, oil residues and hydrocarbon materials. At present, environmental pollution is becoming increasingly serious. Bioremediation, a co-friendly environmental restoration strategy, can convert toxic materials into non-toxic materials or small molecule chemicals used for assimilation. As a potential degrader, Y. lipolytica has been reviewed for its many applications in environmental bioremediation (Bankar et al., 2009a; Fickers et al., 2005a; Zinjarde et al., 2014). For example, Strouhal et al. (2003) revealed that the strain Y. lipolytica strain CCM 4510 has considerable resistance to different heavy metals, such as cadmium (0–75 μM), nickel (0–2400 μM), cobalt (0–2400 μM) and zinc (0–9600 μM). In that research, the data showed that these heavy metals are able to incorporate into individual cell compartments of Y. lipolytica. Moreover, it was shown that metallothionein, a protective protein with a potential role in binding heavy metals, could be produced by Y. lipolytica. Ito et al. (2007) showed that Y. lipolytica also can be grown on high concentrations (2–4 mM) of copper sulfate. Bankar et al. (2009b) reported that the hexavalent chromium ion [Cr (VI)], a stable and toxic heavy metal, can be reduced from aqueous solutions by two marine Y. lipolytica strains. In that research, under optimal conditions (pH of 1.0, temperature of 35 °C, agitation of 130 rpm and contact time of 2 h), Y. lipolytica NCIM 3589 and 3590 displayed specific uptake of Cr (VI) ions of 64 mg/g at a concentration of 950 ppm and 46 mg/g at a concentration of 955 ppm, respectively. Recently, it was confirmed that Ni (II), a toxic heavy metal, could also be removed from wastewaters by marine Y. lipolytica strains (Shinde et al., 2012). Under optimal conditions with an initial Ni (II) ion concentration of 1000 mg/L, Y. lipolytica NCIM 3589 and 3590 displayed a maximum


16


uptake of 95.33 mg/g and 85.44 mg/g, respectively. Bromoalkanes, as hydrophobic organic compounds, can usually be degraded by bacteria under aerobic and anaerobic conditions. However, Vatsal et al. (2011) presented that Y. lipolytica NCIM 3589 is also able to utilize a broad range of bromoalkanes with different carbon chain lengths that vary in degree and position of bromide substitution for the first time and 1bromodecane was the best utilized substrate among the bromoalkanes tested. Petroleum oil-polluted soil or water is a serious environmental pollution problem. Moreover, lipid-rich wastes, such as fat or oil residues, are also serious pollutants. Generally, biosurfactants, which have particular properties of high emulsification activity and stability, can accelerate oil-polluted chemical bioavailability in an eco-friendly manner. Usually, these compounds can be produced by Y. lipolytica grown on hydrophobic materials such as alkanes or triacylglycerols in polluted environments. For example, one bioemulsifier, called liposan (which is mainly composed of carbohydrate and protein), was produced by Y. lipolytica grown on water-immiscible carbon substrates (Cirigliano and Carman, 1984, 1985). Zinjarde and Pant (2002) showed that the Y. lipolytica NCIM 3589, which is isolated from marine waters, can produce a cell-associated emulsifier in the presence of alkanes (C10–C18) or crude oil. Moreover, it was reported that the optional NaCl concentration (from 2 to 3%) could increase emulsifier production. Amaral et al. (2006) discovered a novel bioemulsifier (Yansan) produced by Y. lipolytica (IMUFRJ50682) grown on glucose-based culture medium. It was demonstrated that this bioemulsifier had potential applications due to its high emulsification activities. Another biosurfactant, Rufisan, which is an antimicrobial and anti-adhesive agent, was produced by Y. lipolytica UCP0988 with a high yield of approximately 4.5 g/L when this strain was grown on medium supplemented with soybean oil refinery residue (Rufino et al., 2007, 2011). The yield of bioemulsifier is affected by many factors such as the carbon source, oxygen, and pH. To date, many studies have shown the effects of various experimental factors on biosurfactant production (Albuquerque et al., 2006; Fontes et al., 2010; Sarubbo et al., 2001). Due to its biochemical characteristics, Y. lipolytica can synthesize lipases directly for lipid-rich waste treatments. Zinjarde et al. (2008) reported that the strain Y. lipolytica NCIM 3589 can produce extracellular lipases when using various crude oils as carbon sources. Therefore, many research groups and companies are paying more attention to this field. For example, Groenewald et al. (2014) mentioned that the Belgian company Artechno has developed a Y. lipolyticaderived lipase for the bioremediation of lipid-rich wastewaters. In particular, the typical wastes of olive mill wastewater and palm oil mill, due to the high COD and biological oxygen demand (BOD) values, can be utilized effectively by Y. lipolytica. Oswal et al. (2002) presented that Y. lipolytica NCIM 3589 can effectively decrease COD and BOD values of palm oil mill effluent. Papanikolaou et al. (2008b) reported that olive mill wastewater can be used as a carbon source for citric acid production by Y. lipolytica. Song et al. (2011) constructed an effective lipase-displaying recombinant Y. lipolytica (pINA1296-F-lipRS) for oily wastewater treatment. In that research, the COD of oily wastewater was effectively reduced. Sarris et al. (2011) reported the possibility of citric acid production from three different Y. lipolytica strains (ACA-YC 5028, W29 and ACA-YC 5033) using glucose-enriched olive mill wastewaters as substrates. In that research, the highest citric acid yield of 18.9 g/L was produced by Y. lipolytica ACA-YC 5033 under nitrogen limited conditions. Moreover, that research showed that the tested Y. lipolytica strains (Y. lipolytica W29 and ACA-YC 5033) could be used for olive mill wastewater bioremediation and production of addedvalue compounds. Louhasakul and Cheirsilp (2013) demonstrated that the decanted effluent from palm oil mill can be used as a potential carbon source for lipid synthesis by Y. lipolytica. Moreover, other wastes, such as pineapple waste, vegetable oil refinery residues and industrial fats, could be used as raw materials for important chemical production (Imandi et al., 2008; Papanikolaou et al., 2002a; Rufino et al., 2007). Naturally, when grown on polluted substrates, Y. lipolytica can form

biofilms capable of resisting a variety of antimicrobial agents. Dusane et al. (2012) demonstrated that rhamnolipid, a biological surfactant, has the potential to control biofilm formation by effectively disrupting biofilms. In addition, toxic materials and heavy metals, as serious environmental pollutants, can be detoxified by the Y. lipolytica strain. For example, based on the complex metabolic reactions and multiple enzymes produced by Y. lipolytica, toxic materials such as phenol and biphenyl can be detoxified through hydroxylation. The effects and detoxification mechanisms of toxic materials and heavy metals have been reviewed (Bankar et al., 2009a; Zinjarde et al., 2014). Based on these advances, it is concluded that Y. lipolytica, due to its inherent capabilities, can play an important role in bioremediation. Moreover, it is concluded that combining Y. lipolytica with other microorganisms also has positive effects on environmental bioremediation (Kaczorek et al., 2008). X. Liu et al. (2014b) reported that Y. lipolytica SWJ-1b co-cultured with the immobilized mycelia of T. reesei can utilize raw biomass, such as pretreated straw, for organic acid production. 3.5. Aromatic compounds Aromatic compounds, such as lactones and esters, can be synthesized by bioprocessing. Schrader et al. (2004) reviewed the synthesis pathways and future potential applications of flavor compounds. Lactones are important aromatic compounds and can be produced by different microorganisms such as yeasts and fungi (Romero-Guido et al., 2011). Waché et al. (2003) reviewed the biotechnological production of lactones by Y. lipolytica. Among various types of lactones, γdecalactone is an important peach-like aromatic compound that can be synthesized through β-oxidation with methyl-ricinoleate as a substrate. The accumulation of γ-decalactone is mainly related to the biochemical reaction of β-oxidation, which contains four steps. Naturally, the first and most important step of β-oxidation is catalyzed by acylCoA oxidase (Aox1–6 encoded by the POX1–6 genes) (Nicaud et al., 1998; Wang et al., 1998). These enzymes have particular roles in βdegradation. In the Aox family, Aox2 performs long-chain oxidation, whereas Aox3 mediates short-chain degradation (Mlícková et al., 2004a). In addition, Aox4p was found to be involved in lactone degradation (Groguenin et al., 2004). Moreover, other by-products such as 3hydroxy-γ-decalactone, dec-2-en-4-olide and dec-3-ec-4-olide, can be synthesized during γ-decalactone accumulation. The effective strategy of blocking β-oxidation via genetic modification was developed for modifying the carbon flux and improving target chemical accumulation to increase γ-decalactone yield (Waché et al., 2001, 2002). Through expression of the POX2 gene and disruption of the POX3 gene, γ-decalactone yield was increased in recombinant Y. lipolytica under optimized conditions with methyl ricinoleate as a carbon source (Guo et al., 2012). In addition, many environmental factors, such as substrate concentration, pH, aeration and dissolved oxygen content, also played important roles in altering the β-oxidation carbon flux (García et al., 2009). Of these factors, the strategy of improving oxygen availability can significantly improve aromatic compound production. Aguedo et al. (2005) observed the influence of different oxygenation conditions on aromatic compound accumulation. In that research, 263 mg/L of 3-hydroxy-γ-decalactone was effectively produced with a kLa of 120 h−1 from Y. lipolytica W29 grown on glucose as the carbon source. Gomes et al. (2007) reported that the kLa plays an important role in aromatic compound production. Under optimized conditions with a kLa of 70 h−1, agitation rates of 400 rpm and aeration rates of 0.6 vvm, a maximal amount of γ-decalactone of up to 141 mg/L was produced by Y. lipolytica W29 grown on methyl ricinoleate as the sole carbon source. Moreover, using methyl ricinoleate as the carbon source, Gomes et al. (2010) determined the effect of the oxygen transfer rate on γ-decalactone production from Y. lipolytica W29. Under optimized conditions, γ-decalactone productivity of up to 87 mg/L/h was observed via a new method that avoided the laborious step of washing cells, while 3hydroxy-γ-decalactone, another important compound, also accumulated.



Moradi et al. (2013) illustrated γ-decalactone accumulation from Y. lipolytica with castor oil as a carbon source. Under optimized conditions, a high concentration of γ-decalactone of up to 220 mg/L was produced by Y. lipolytica (DSM3286). Braga and Belo (2014) demonstrated the influence of the oxygen transfer rate on γ-decalactone production. In this research, a maximum γ-decalactone concentration of 5.4 g/L and a cell density of 60 g/L were obtained in Y. lipolytica W29 grown on castor oil as a carbon source. Moreover, to increase γ-decalactone production, an efficient method of cell immobilization was developed. In this research, a maximum aromatic concentration of up to 1597 mg/L was produced by Y. lipolytica W29 cells immobilized with DupUM® using castor oil as a carbon source (Braga and Belo, 2013). 2-Phenylethanol, a rose-like aroma compound, has many potential applications. It can be naturally synthesized by different microorganisms. Recently, it was demonstrated that Y. lipolytica is deemed a novel 2-phenylethanol producer. Celińska and Grajek (2013) demonstrated that 2-phenylethanol is produced as a by-product together with citric acid production with glycerol as a carbon source. A 2phenylethanol yield of up to 1 g/L was produced by the recombinant strain NCYC3825 in a shake-flask culture. Celińska et al. (2013) obtained 2 g/L of 2-phenylethanol from Y. lipolytica NCYC3825 grown on glucose under non-optimized culture conditions. Moreover, the green-note aromatic compounds, such as C6 aldehydes, can also be synthesized in Y. lipolytica. In the case of the C6 aldehydes, Bourel et al. (2004) demonstrated that hexanal and trans-2-hexenal are produced via expressing the hydroperoxide lyase-encoding gene in Y. lipolytica grown on olive oil as a fermentable substrate.

3.6. Others Polyalcohols, due to their particular characteristics, have potential applications. Recently, using Y. lipolytica, polyalcohols, such as erythritol, mannitol and arabitol, could also be produced during the

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fermentation process. The recent applications of polyalcohol production from Y. lipolytica are summarized in Table 2. Erythritol is an FDA-approved four-carbon sugar alcohol with a wide range of potential applications. Generally, erythritol can be synthesized by chemical and biotechnological pathways. Moon et al. (2010) reviewed the biotechnological pathways of erythritol and its applications. Using Y. lipolytica, erythritol can be naturally synthesized via the pentose phosphate pathway when this yeast grows on glucose as a carbon source (Ghezelbash et al., 2014; Rywińska et al., 2010b). Recently, although pure glycerol with high cost or raw glycerol with various impurities is not a suitable carbon substrate for polyalcohol production, using pure glycerol or raw glycerol for erythritol production has received more interest for academic research. Among the research groups involved in these projects, the Rymowicz group has made a lot of progress in polyalcohol production from Y. lipolytica using glycerol as a carbon source (Rymowicz et al., 2008, 2009, 2010; Rywińska et al., 2009, 2010a, 2010b, 2015). Generally, erythritol was produced with citric acid production from wild-type and mutant Y. lipolytica strains using glycerol as a carbon source. Rymowicz et al. (2009) optimized the fermentation process, and 170 g/L of erythritol was produced by mutant Y. lipolytica Wratislavia K1 in a fed-batch culture with raw glycerol as the sole carbon source, while no citric acid was obtained under the same conditions. Naturally, erythritol productivity is affected by various factors, such as the culture mode, agitation rate and osmotic pressure (Rywińska et al., 2013b, 2015; Tomaszewska et al., 2012, 2014). Mirończuk et al. (2014) developed the fed-batch culture mode for erythritol production. Under optimized conditions, up to 220 g/L erythritol was produced by mutant Y. lipolytica Wratislavia K1. Yang et al. (2014) demonstrated that high osmotic pressure can enhance cell growth and improve erythritol production. Based on the twostage osmotic pressure control fed-batch fermentation mode, 194.3 g/L of erythritol was produced by Y. lipolytica grown on glycerol as a carbon source. In addition to erythritol, other polyalcohols, such as mannitol and arabitol, were produced together with erythritol

Table 2 Fermentative production of polyols produced by Yarrowia lipolytica. Strains

Carbon sources

Polyols

Culture mode

Production (g/L)

References

DSM70562 mutant 49 A-101 Wratislavia K1 CICC 1675 CICC 1675 Wratislavia K1 Wratislavia K1 Wratislavia K1 Wratislavia AWG7 A-101-1.22 Wratislavia K1 Wratislavia K1 Wratislavia K1 LFMB 20 LFMB 19 LFMB 20 8661 UV1 Wratislavia AWG7 A-101-1.22 Wratislavia K1 Wratislavia K1 Wratislavia K1 Wratislavia K1 Wratislavia K1 Wratislavia K1 Wratislavia K1 Wratislavia AWG7 Wratislavia K1 A UV'1 Wratislavia K1 A UV'1 Wratislavia K1

Glucose Glucose Glycerol Glycerol Glycerol Raw glycerol Raw glycerol Raw glycerol Raw glycerol Raw glycerol Raw glycerol Raw glycerol Raw glycerol Raw glycerol Raw glycerol Raw glycerol Raw glycerol Raw glycerol Raw glycerol Raw glycerol Raw glycerol Pure glycerol Pure glycerol Pure glycerol Pure glycerol Pure glycerol Pure glycerol Pure glycerol Pure glycerol Pure glycerol Pure glycerol Pure glycerol

Erythritol Mannitol Erythritol Erythritol Mannitol Erythritol Erythritol Erythritol Erythritol Erythritol Erythritol Erythritol Erythritol Mannitol Mannitol Mannitol Mannitol Mannitol Mannitol Mannitol Arabitol Erythritol Erythritol Erythritol Erythritol Erythritol Mannitol Mannitol Mannitol Mannitol Arabitol Arabitol

Batch Batch Batch Fed-batch Fed-batch Fed-batch Fed-batch Fed-batch Fed-batch Reaped batch Batch Batch Reaped batch Batch Batch Batch Fed-batch Fed-batch Reaped batch Reaped batch Reaped batch Batch Batch Reaped batch Batch Reaped batch Fed-batch Batch Batch Batch Batch Batch

39.7 7.3 47.1 194.3 41.2 81 170 46.9 44.1 45.2 80.0 58.2 180 6.0 19.4 9.6 40.5 11.2 4.75 9.2 1.2 32.9 84.1 220 46.9 132 9.0 13.7 27.6 15.1 9.2 7.9

Ghezelbash et al. (2014) Rywińska et al. (2010b) Tomaszewska et al. (2014) Yang et al. (2014) Yang et al. (2014) Rymowicz et al. (2008) Rymowicz et al. (2009) Rywińska et al. (2009) Rywińska et al. (2010a) Rymowicz et al. (2010) Tomaszewska et al. (2012) Rywińska et al. (2013b) Mirończuk et al. (2014) André et al. (2009) Chatzifragkou et al. (2011a) Chatzifragkou et al. (2011b) Rymowicz et al. (2009) Rywińska et al. (2010a) Rymowicz et al. (2010) Mirończuk et al. (2014) Mirończuk et al. (2014) Rywińska et al. (2010b) Tomaszewska et al. (2012) Mirończuk et al. (2014) Rywińska et al. (2015) Rywińska et al. (2015) Rywińska et al. (2009) Rywińska et al. (2010a) Tomaszewska et al. (2012) Rywińska et al. (2013b) Tomaszewska et al. (2012) Rywińska et al. (2010a)


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accumulation (Rymowicz et al., 2009, 2010; Rywińska et al., 2009, 2010a, 2010b; Tomaszewska et al., 2012). Chatzifragkou et al. (2011a) reported that 19.4 g/L mannitol is produced by Y. lipolytica LFMB 19 with raw glycerol as the sole carbon source. Under optimal initial osmotic pressure, Yang et al. (2014) revealed that a high mannitol concentration of 41.2 g/L is produced by Y. lipolytica CICC 1675 in fed-batch fermentation. Recently, Rywińska et al. (2013b) reported that 7.9 g/L arabitol is produced by Y. lipolytica Wratislavia K1 grown on pure glycerol in batch fermentation mode. Polyhydroxyalkanoates are important biodegradable chemical biopolymers, which mainly include short-chain length, medium-chain length or long-chain length polyhydroxyalkanoates and can be synthesized by many microorganisms. Among these microorganisms, Y. lipolytica, based on its inherent characteristics, has been selected as an ideal host for polyhydroxyalkanoate production. Naturally, polyhydroxyalkanoate can be synthesized from 3-hydroxyacyl-CoA, which is an intermediate in the β-oxidation cycle. Therefore, the strategy of modification to the β-oxidation cycle has been developed for balancing the carbon flux for polyhydroxyalkanoate production. Haddouche et al. (2010) showed the effect of the multiple acyl-CoA oxidase isoenzymes encoded by POX genes (POX1 to POX6) on the flux of β-oxidation intermediates towards polyhydroxyalkanoate biosynthesis in Y. lipolytica that expressed polyhydroxyalkanoate synthase. Their research revealed that the POX3 genotype remarkably affected the polyhydroxyalkanoate production levels from different chain length fatty acids such as tridecanoic acid (C13:0) or nonanoic acid (C9:0), respectively. However, their research showed that changes in POX genes had no effects on the monomer composition of polyhydroxyalkanoate. Recently, Haddouche et al. (2011) reported that the expression of βoxidation multifunctional enzyme (MFE2) variants have potential to obtain polyhydroxyalkanoate with various monomer compositions in Y. lipolytica. Carotenoids, including more than 600 diverse compounds, are important chemicals with potential applications that can be synthesized by different organisms. However, these natural, synthetic carotenoids cannot meet the huge market demand. Therefore, it is a necessary to develop new strategies for improving carotenoid production. Based on the development of metabolic engineering and synthetic biology, a high yield of carotenoids has been produced via constructing and tuning the synthetic pathway in patented recombinant Y. lipolytica strains (Bailey et al., 2010). Lycopene is an important central carotenoid, and it can be synthesized via metabolic pathway modification. Matthäus et al. (2014) demonstrated lycopene production in a Y. lipolytica strain with raw glycerol as a carbon source. Upon deletion of the POX1 to POX6 and GUT2 genes and overexpression of the GGS1 and HMG1 genes, a yield of 16 mg/g (DCW) lycopene was ultimately achieved in fed-batch fermentation. Petroleum fuels, such as gasoline, diesel, and jet fuel, contain a complex mixture of hundreds of hydrocarbons. In particular, gasoline is a complex, variable mixture of hydrocarbons and other additives that contain a mixture of C4 to C12 short-chain hydrocarbons. However, there are only a few reports on short-chain hydrocarbon production via microbial fermentation at present. For example, Choi and Lee (2013) reported that short-chain hydrocarbons, including octane and dodecone, can be produced by an engineered E. coli strain. Interestingly, with Y. lipolytica as a platform, the highest pentane yield of 4.98 mg/L was produced via a combinational strategy with overexpression of the soybean lipoxygenase I gene (Gmlox1) and the hydroperoxide lyase gene (Gmhpl1) and deletion of the MFE1 and PEX10 genes under optimized conditions with LA as a substrate (Blazeck et al., 2013a). 4. Conclusions and prospects As a GRAS-approved non-conventional yeast, Y. lipolytica has potential applications in metabolite synthesis, functional protein production and environmental bioremediation due to its particular characteristics.

To date, many research groups have achieved important progress in this field of Y. lipolytica. For example, the Papanikolaou group at the Agricultural University of Athens and the Aggelis group at the University of Patras of Greece have presented many reports indicating that Y. lipolytica can assimilate and utilize various carbon sources (such as industrial fats, glycerol and glucose). Significantly, many potential metabolites, including organic acids and microbial lipids, can be produced when raw carbon materials are used as carbon sources. Moreover, the Rymowicz group at the Wroclaw University of Environmental and Life Sciences of Poland has also made a lot of progress in organic acid and polyalcohol production from Y. lipolytica using glycerol as a carbon source. With the development of biotechnology, the Madzak group at the INRA has developed a series of genetic and molecular tools for Y. lipolytica. Recently, Madzak (2015) presented a very synthetic review on the subject of Y. lipolytica engineering. Specifically, the established genetic tools for heterologous protein expression, secretion, and surface display, including strains, vectors, promoters, selection markers, multiple integrations and signals for secretion, targeting and display, were reviewed in detail in this mini-review. The Nicaud group at the INRA has also presented important genetic tools for tuning gene expression. Moreover, the Nicaud group has identified many functional genes involved in the metabolism and degradation of hydrophobic material. Furthermore, new surface display vectors have been developed by the Chi group at the Ocean University of China, which will enable a wide range of applications of Y. lipolytica. In addition, “omics” research on Y. lipolytica has been reported under the development of system biology technology. A genome-scale metabolic network model of Y. lipolytica has been presented (Loira et al., 2012; Pan and Hua, 2012). Morin et al. (2011) presented the transcriptome analysis of Y. lipolytica and identified functional genes involved in lipid metabolism. Zhao et al. (2014) reported a pretreatment method for analyzing intracellular metabolites from Y. lipolytica. In addition, Y. lipolytica has been used as a model for studies on mitochondrial complexes, RNA splicing and genome evolution. Moreover, Y. lipolytica is deemed as a whole-cell biocatalyst and has been applied in miscellaneous applications such as biosensor, fine chemistry and pharmaceutical applications. Therefore, Y. lipolytica is a promising cell factory that has garnered great interest. Recently, the “LipoYeasts” project was developed via constructing and optimizing metabolic pathways to synthesize important lipid-derived industrial products from engineered Y. lipolytica (Sabirova et al., 2011). Although many industrial applications of Y. lipolytica have been reported, some problems that limit the potential development of this field remain. One of the limitations is that an ideal host for further applications needs to be selected. Because the characteristics of Y. lipolytica are intrinsically strain-dependent, many products cannot be produced on commercial scale. Therefore, host selection is deemed a crucial factor for producing specific key products. In addition, many key enzymes involved in metabolic reactions are still unclear. These problems will hinder us from doing further research. With the development of biotechnology, especially the sequencing of the Y. lipolytica genome, a series of genetic and molecular tools have been constructed and used as a platform for Y. lipolytica. Moreover, with advances in yeast system biology and synthetic biology-based metabolic engineering, many different approaches, such as biochemical engineering, genetic engineering, transcription factor engineering, modular pathway engineering and compartmentalization metabolic engineering and various synthetic methods, such as promoter engineering, ribosome binding site engineering, terminator engineering, DNA assembly, RNA devices and scaffolds, will be used for tuning gene expression and balancing target product metabolic flux (Agapakis, 2014; Boyle and Silver, 2012; Keasling, 2012; Kim et al., 2014). Recently, Gao et al. (2014) reported that an entire β-carotene biosynthesis pathway with multiple fragments has been assembled via in vivo homologous recombination in Y. lipolytica ATCC 201249. Altogether, it has been concluded that these technologies will provide new ideas and opportunities for more natural products, especially added-value chemical production from Y. lipolytica in the future.



Acknowledgements This work was financially supported by the National Science Foundation for Distinguished Young Scholars of China (No. 21225626), the National Basic Research Program of China (No. 2011CBA00800), the National Natural Science Foundation of China (Nos. 21376002 and 21476111), Jiangsu Province Natural Science Foundation of China (No. BK20131405), and the National High-Tech R&D Program of China (Nos. 2014AA021701 and 2014AA021703). We also acknowledge the support of the project funded by the Priority Academic Program Development of Jiangsu Higher Education Institutions. References Abghari, A., Chen, S., 2014. Yarrowia lipolytica as an oleaginous cell factory platform for the production of fatty acid-based biofuel and bioproducts. Front. Energy Res. 2, 1–21. Agapakis, C.M., 2014. Designing synthetic biology. ACS Synth. Biol. 3, 121–128. Ageitos, J.M., Vallejo, J.A., Veiga-Crespo, P., Villa, T.G., 2011. Oily yeasts as oleaginous cell factories. Appl. Microbiol. 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