of 14/14
Review Advancing oleaginous microorganisms to produce lipid via metabolic engineering technology Ming-Hua Liang a , Jian-Guo Jiang a,b,a School of Biological Science & Engineering, South China University of Technology, Guangzhou 510006, China b College of Food Science and Engineering, South China University of Technology, Guangzhou 510640, China article info Article history: Received 1 February 2013 Received in revised form 3 May 2013 Accepted 6 May 2013 Available online 16 May 2013 Keywords: Oleaginous microorganisms Metabolic engineering Biodiesel Microbial oils Lipids Microalgae abstract With the depletion of global petroleum and its increasing price, biodiesel has been becoming one of the most promising biofuels for global fuels market. Researchers exploit oleaginous microorganisms for bio- diesel production due to their short life cycle, less labor required, less affection by venue, and easier to scale up. Many oleaginous microorganisms can accumulate lipids, especially triacylglycerols (TAGs), which are the main materials for biodiesel production. This review is covering the related researches on different oleaginous microorganisms, such as yeast, mold, bacteria and microalgae, which might become the potential oil feedstocks for biodiesel production in the future, showing that biodiesel from oleaginous microorganisms has a great prospect in the development of biomass energy. Microbial oils biosynthesis process includes fatty acid synthesis approach and TAG synthesis approach. In addition, the strategies to increase lipids accumulation via metabolic engineering technology, involving the enhancement of fatty acid synthesis approach, the enhancement of TAG synthesis approach, the regula- tion of related TAG biosynthesis bypass approaches, the blocking of competing pathways and the multi- gene approach, are discussed in detail. It is suggested that DGAT and ME are the most promising targets for gene transformation, and reducing PEPC activity is observed to be beneficial for lipid production. Ó 2013 Elsevier Ltd. All rights reserved. Contents 1. Introduction ......................................................................................................... 396 2. Oleaginous microorganisms for biodiesel production ........................................................................ 396 2.1. Oleaginous microalgae ........................................................................................... 397 2.2. Oleaginous yeast and mold........................................................................................ 397 2.3. Oleaginous bacteria .............................................................................................. 398 3. TAG biosynthesis in microorganisms ..................................................................................... 400 3.1. Fatty acid synthesis approach...................................................................................... 400 3.2. TAG synthesis approach .......................................................................................... 400 4. Metabolic engineering for TAG production................................................................................. 401 4.1. Enhancement of fatty acid synthesis approach ........................................................................ 402 4.1.1. Acetyl-CoA carboxylase (ACC) .............................................................................. 402 4.1.2. Fatty acid synthetase (FAS) ................................................................................ 402 4.1.3. Acyl-ACP-thioesterase (FAT) ............................................................................... 402 0163-7827/$ - see front matter Ó 2013 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.plipres.2013.05.002 Abbreviations: ACC, acetyl-CoA carboxylase; ACL, ATP:citrate lyase; ACP, acyl-carrier protein; AGPase, ADP-glucose pyrophosphate; AOX, acyl-CoA oxidase; ARA, arachidonic acid; BE, branching enzymes; CDP-DAG, CDP-diacylglycerol; DAG, diacylglycerol; DGAT, diacylglycerol acyl-transferase; DHA, docosahexenoic acid; DHAP, dihydroxyacetone phosphate; DHAPAT, DHAP acyltransferase; EPA, eicosapentaenoic acid; FAS, fatty acid synthetase; FAT, acyl-ACP-thioesterase; FFA, free fatty acid; GAP, glyceraldehyde 3-phosphate; GLA, gamma-linolenic acid; G-1-P, glucose 1-phosphate; G-6-P, glucose 6-phosphate; G3P, glycerol-3-phosphate; GPAT, glycerol-3-phosphate acyltransferase; GPD1 and GUT2, glycerol 3-phosphate dehydrogenase; KAS, b-ketoacyl-ACP synthase; LPA, lysophosphatidate; LPAT, lysophosphatidate acyl-transferase; MAT, malonyl-CoA:ACP transacetylase; ME, malic enzyme; PA, phosphatidate; PAP, phosphatidic acid phosphatase; PDAT, phospholipid:diacylglycerol acyltransferase; PDH, pyruvate dehydrogenase; PEP, phosphoenolpyruvate; PEPC, phosphoenolpyruvate carboxylase; Pi, inorganic pyrophosphate; PYC, pyruvate carboxylase; SS, starch synthase; TAG, triacylglycerol; WS/DGAT, wax ester synthase/acyl-CoA:diacylglycerol acyltransferase. Corresponding author at: College of Food Science and Engineering, South China University of Technology, Guangzhou 510640, China. Tel./fax: +86 20 87113849. E-mail address: [email protected] (J.-G. Jiang). Progress in Lipid Research 52 (2013) 395–408 Contents lists available at SciVerse ScienceDirect Progress in Lipid Research journal homepage: www.elsevier.com/locate/plipres

Progress in Lipid Research - miyazaki-u

  • View
    0

  • Download
    0

Embed Size (px)

Text of Progress in Lipid Research - miyazaki-u

Advancing oleaginous microorganisms to produce lipid via metabolic engineering technologyContents lists available at SciVerse ScienceDirect
Progress in Lipid Research
Review
0163-7827/$ - see front matter 2013 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.plipres.2013.05.002
Abbreviations: ACC, acetyl-CoA carboxylase; ACL, ATP:citrate lyase; ACP, acyl-carrier protein; AGPase, ADP-glucose pyrophosphate; AOX, acyl-CoA oxida arachidonic acid; BE, branching enzymes; CDP-DAG, CDP-diacylglycerol; DAG, diacylglycerol; DGAT, diacylglycerol acyl-transferase; DHA, docosahexenoic acid dihydroxyacetone phosphate; DHAPAT, DHAP acyltransferase; EPA, eicosapentaenoic acid; FAS, fatty acid synthetase; FAT, acyl-ACP-thioesterase; FFA, free fatty a glyceraldehyde 3-phosphate; GLA, gamma-linolenic acid; G-1-P, glucose 1-phosphate; G-6-P, glucose 6-phosphate; G3P, glycerol-3-phosphate; GPAT, glycerol-3-ph acyltransferase; GPD1 and GUT2, glycerol 3-phosphate dehydrogenase; KAS, b-ketoacyl-ACP synthase; LPA, lysophosphatidate; LPAT, lysophosphatidate acyl-tra MAT, malonyl-CoA:ACP transacetylase; ME, malic enzyme; PA, phosphatidate; PAP, phosphatidic acid phosphatase; PDAT, phospholipid:diacylglycerol acyltransfera pyruvate dehydrogenase; PEP, phosphoenolpyruvate; PEPC, phosphoenolpyruvate carboxylase; Pi, inorganic pyrophosphate; PYC, pyruvate carboxylase; SS, starch s TAG, triacylglycerol; WS/DGAT, wax ester synthase/acyl-CoA:diacylglycerol acyltransferase. ⇑ Corresponding author at: College of Food Science and Engineering, South China University of Technology, Guangzhou 510640, China. Tel./fax: +86 20 8711384
E-mail address: [email protected] (J.-G. Jiang).
Ming-Hua Liang a, Jian-Guo Jiang a,b,⇑ a School of Biological Science & Engineering, South China University of Technology, Guangzhou 510006, China b College of Food Science and Engineering, South China University of Technology, Guangzhou 510640, China
a r t i c l e i n f o
Article history: Received 1 February 2013 Received in revised form 3 May 2013 Accepted 6 May 2013 Available online 16 May 2013
Keywords: Oleaginous microorganisms Metabolic engineering Biodiesel Microbial oils Lipids Microalgae
a b s t r a c t
With the depletion of global petroleum and its increasing price, biodiesel has been becoming one of the most promising biofuels for global fuels market. Researchers exploit oleaginous microorganisms for bio- diesel production due to their short life cycle, less labor required, less affection by venue, and easier to scale up. Many oleaginous microorganisms can accumulate lipids, especially triacylglycerols (TAGs), which are the main materials for biodiesel production. This review is covering the related researches on different oleaginous microorganisms, such as yeast, mold, bacteria and microalgae, which might become the potential oil feedstocks for biodiesel production in the future, showing that biodiesel from oleaginous microorganisms has a great prospect in the development of biomass energy. Microbial oils biosynthesis process includes fatty acid synthesis approach and TAG synthesis approach. In addition, the strategies to increase lipids accumulation via metabolic engineering technology, involving the enhancement of fatty acid synthesis approach, the enhancement of TAG synthesis approach, the regula- tion of related TAG biosynthesis bypass approaches, the blocking of competing pathways and the multi- gene approach, are discussed in detail. It is suggested that DGAT and ME are the most promising targets for gene transformation, and reducing PEPC activity is observed to be beneficial for lipid production.
2013 Elsevier Ltd. All rights reserved.
Contents
1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 396 2. Oleaginous microorganisms for biodiesel production . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 396
2.1. Oleaginous microalgae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 397 2.2. Oleaginous yeast and mold. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 397 2.3. Oleaginous bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 398
3. TAG biosynthesis in microorganisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 400
3.1. Fatty acid synthesis approach. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 400 3.2. TAG synthesis approach . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 400
4. Metabolic engineering for TAG production. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 401
4.1. Enhancement of fatty acid synthesis approach . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 402
4.1.1. Acetyl-CoA carboxylase (ACC) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 402 4.1.2. Fatty acid synthetase (FAS) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 402 4.1.3. Acyl-ACP-thioesterase (FAT) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 402
se; ARA, ; DHAP,
cid; GAP, osphate
http://dx.doi.org/10.1016/j.plipres.2013.05.002
mailto:[email protected]
http://dx.doi.org/10.1016/j.plipres.2013.05.002
http://www.sciencedirect.com/science/journal/01637827
http://www.elsevier.com/locate/plipres
0
2
4
6
8
10
12
14
16
18
20
19
B
396 M.-H. Liang, J.-G. Jiang / Progress in Lipid Research 52 (2013) 395–408
4.2. Enhancement of TAG synthesis approach . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 402
4.2.1. Acyl-CoA:glycerol-sn-3-phosphate acyl-transferase (GPAT). . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 402 4.2.2. Lysophosphatidate acyl-transferase (LPAT). . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 403 4.2.3. Acyl-CoA:diacylglycerol acyl-transferase (DGAT) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 403 4.2.4. Glycerol 3-phosphate dehydrogenase (GPDH) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 403
4.3. Regulation of related TAG biosynthesis bypass approaches . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 403
4.3.1. Acetyl-CoA synthase (ACS). . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 403 4.3.2. Malic enzyme (ME). . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 403 4.3.3. ATP:citrate lyase (ACL) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 403
4.4. Blocking competing pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 404
4.4.1. Repression of b-oxidation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 404 4.4.2. Repression of phospholipid biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 404 4.4.3. Repression of phosphoenolpyruvate carboxylase (PEPC) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 404 4.4.4. Repression of starch biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 404 4.4.5. Repression of the degradation of TAG. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 405
4.5. The multi-gene approach . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 405
5. Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 405
1. Introduction
The earliest research on producing lipids from microorganisms could be traced back to the First World War, when Germany had prepared with some strains of Endomyces and Fusarium sp. to pro- duce lipids to solve the cooking oil shortage problem [1]. In recent years, high energy prices, energy and environment security, con- cerns about petroleum supplies are drawn great attention and drive us to find a renewable biofuel. One of the most promising renewable biofuels is biodiesel, a mixture of fatty acid methyl es- ters, and generally speaking, it is produced from vegetable oils, ani- mal fats or wasting oils [2] by transesterification of triacylglycerols (TAGs) with short chain alcohols.
The global markets for biodiesel are entering a period of rapid and transitional growth. In the year 2007, there were only 20 nations producing biodiesel for the needs of over 200 nations; by the year 2010, more than 200 nations became biodiesel producing nations and suppliers. Global biodiesel production has massively increased to 18.2 billion liters per year from 2000 to 2010 (Fig. 1). However, the plant oil materials require large energy and acreage for sufficient production of oilseed crops. For example, using the average oil yield per hectare from various crops, the cropping area needed to meet 50% of the US transport fuel needs is shown in Fig. 2. This area is ex- pressed as a percentage of the total cropping area of the US. In view of Fig. 2, microalgae appear to be the only source of biodiesel that
99 2000 2001 2002 2003 2004 2005 2006 2007 2008 2009 2010 2011
illion liters
Fig. 1. Global biodiesel production in 2000–2010 [4].
has the potential to completely displace fossil diesel. If plant oil was used for biodiesel production, the cost of source would account for 70–85% of the whole production cost [3]. On the other hand, ani- mal fat oils need to feed these animals. So the cost of oil feedstocks limits the large-scale development of biodiesel to a large extent. Therefore, to lower the cost of oil raw materials, much attention has been paid to the development of microbial oils and it has been found that many microorganisms, such as microalgae, yeast, bacte- ria, and fungi, have the ability to accumulate oils under some special cultivation conditions.
Compared to plant oils, microbial oils have many advantages, such as short life cycle, abundant and cheap raw materials, less labor required, less affections by venue, season and climate, and easier to scale up [8,9]. Therefore, microorganisms might become one of the potential oil feedstocks for biodiesel production in the future, although there are many researches associated with micro- organism-producing oils that need to be further carried out. This review is covering the related researches about different oleagi- nous microorganisms for lipids production and microbial TAG bio- synthesis process. Moreover, metabolic engineering strategies to increase lipid production are introduced in detail.
2. Oleaginous microorganisms for biodiesel production
Oleaginous microorganisms are defined as oleaginous species with oil contents excess of 20% biomass weight [10]. Microbial oils, also called single cell oils, are produced by some oleaginous micro- organisms, such as yeast, fungi, bacteria, and microalgae [8]. While the eukaryotic yeast, mold and microalgae can synthesize TAGs, which are similar with the composition of vegetable oils, and pro- karyotic bacteria can synthesize specific lipids. Many oleaginous microorganisms can accumulate oils, especially TAGs, which are the main materials for biodiesel production. TAGs act with alcohol under acid or alkali catalyst by transesterification to generate fatty acid methyl ester (biodiesel) and the by-product glycerol [11] (Fig. 3).
Generally speaking, microbial oils might become one of the po- tential feedstocks for biodiesel production in the future. To reduce the cost of microbial oils, exploring other carbon sources instead of glucose is very important especially for such oils applied to biodie- sel production. It was reported that xylose, glycerol, corn straw, and other agricultural and industrial wastes could be used as the carbon sources for microbial oils accumulation. Due to the low
CH
CH2
CH2
OOC-R1
OOC-R2
OOC-R3
0
200
400
600
800
1000
1200
1400
1600
1800
0
20000
40000
60000
80000
100000
120000
140000
160000
Percent of existing U.S. cropping area
O il
y ie
a
Fig. 2. Comparison of some sources of biodiesel [5–7]. aFor meeting 50% of all transport fuel needs of the United States. b30% oil (by dry weight) in biomass. c50% oil (by dry weight) in biomass. d70% oil (by dry weight) in biomass. Use 50 g/m2/d as the productivity of microalgae by photobioreactor for calculation the oil yields of microalgae.
M.-H. Liang, J.-G. Jiang / Progress in Lipid Research 52 (2013) 395–408 397
oil yield in most bacteria [12], researchers mainly focus on micro- algae and fungi at present. Microalgae can use carbon dioxide as the carbon sources and sunlight as the energy for photoautotrophic culture, or use organic carbon as the carbon sources instead of sun- light for heterotrophic culture. And they can also use light with or- ganic carbon as supplementary carbon sources for mixotrophic culture. Scaling up for autotrophic microalgae is more complicated, since light is needed during the cultivation process. To minimize the cost, oil production from microalgae must rely on available free sunlight, despite daily and seasonal variations in light levels [13]. Heterotrophic microalgae are easily cultivated and controlled in normal fermenters. But, they require organic carbon sources for oil accumulation, which might limit the application of such micro- algae used for oil production for biodiesel production [11]. As for fungi, scaling up for fungi needs to be carried out further. To utilize cheap carbon sources for oil production from fungi opens a new way for cost reduction, which is very important for such oils used for biodiesel production in the future [11].
2.1. Oleaginous microalgae
Generally speaking, microalgae whose lipid contents in the cells are more than 20% of the dry weight are called oleaginous microalgae. The average lipid contents of algal cells vary between 1% and 70% but can reach 90% of dry weight under certain conditions [14,15]. Oil levels of 20–50% are quite common [16] (Table 1). Microalgae, such as Chlorophyta, Bacillariophyceae, have higher oil contents, and they are easier to cultivate, especially chlorella, which could be applied to indus- trial production, so they are the ideal energy microalgae resources [17,18]. At present to breed optimal chlorella strains
suitable for large-scale breeding for biodiesel production has aroused great attention from international energy research institutions and biological energy development companies. Miao et al. [19] acquired high lipid content of heterotrophic chlorella by heterotrophic transformation cell engineering technology, containing the lipid compounds as high as 55% of cell dry weight, which was 4 times of the autotrophic one (14%). Nation Renewable Energy Laboratory (NREL) in the United States devel- oped a ‘‘engineering Cyclotella’’ [20], whose oil content can reach more than 60% in the laboratory condition, and more than 40% in outdoor culture, while microalgae lipid content in the nature is generally 5–20%. It is obvious that using ‘‘engineering microalgae’’ for biodiesel production has important economic and ecological significances.
2.2. Oleaginous yeast and mold
Since the Second World War, many highly oleaginous microor- ganisms, like Lipomyces starkeyi, Rhodotorula glutinis, Aspergillus and Mucor and so on were sequentially found. In order to further form productivity providing technical basis, the identification of a variety of oleaginous strains has achieved breakthrough [9]. Many yeast species, such as Cryptococcus albidus, Cryptococcus albidun, Lipomyces lipofer, Lipomyces starkeyi, Rhodosporidium toruloides, Rhodotorula glutinis and Trichosporon pullulans, were found to be able to accumulate lipids under some cultivation conditions (Table 2), suggesting that different yeast species led to different lipid accumulation. Many mold species, such as Asoergullus terreus, Claviceps purpurea, Tolyposporium, Mortierella alpina, Mortierella isabellina, can also accumulate lipids. Although some types of fungi have the ability to produce lipids (Table 2), most fungi are explored mainly for the production of special lipids, such as DHA, GLA, EPA and ARA, and there are few reports on the utilization of fungi oils for biodiesel production [9]. It was reported that when Mortierella isabellina was cultivated in nitro- gen-limited and high utilization rate of glucose medium, or even in high initial sugar concentration medium, significant fatty acid quantities were accumulated after nitrogen depletion, resulting in a notable microbial oils production of 18.0 g/L in culture medium, containing a maximum concentration of 0.801 g GLA per liter of culture medium [36].
Table 1 Oil content of some microalgae.
Microalgal species Culture conditions Lipid content (% dry weight)
Biomass productivity (g/L/day)
Lipid productivity (mg/L/day)
Refs.
Chlorophyta Chlorella ellipsoidea YSR03 Phototrophic 32 ± 5.9 0.07 22.38 [21] Chlorella protothecoides Heterotrophic 49 1.2 586.8 [22] Chlorella sp. Phototrophic 32.6–66.1 0.077–0.338 51–124 [23] Chlorella vulgaris Phototrophic 20–42 0.21–0.346 44–147 [24] Chlorella vulgaris Phototrophic, mixotrophic, heterotrophic 21–38 0.01–0.254 4–54 [25] Dunaliella sp. Phototrophic 12.0–30.12 1.3–3.0 360–390 [26] Haematococcus pluvialis Phototrophic 15.61–34.85 – – [27] Neochloris oleoabundans Phototrophic 7–40 0.31–0.63 38–133 [28] Neochloris oleabundans UTEX #1185 Phototrophic 19–56 0.03–0.15 10.67–38.78 [29] Pseudochlorococcum sp. Phototrophic 24.6–52.1 0.234–0.76 53–350 [30] Scenedesmus obliquus Phototrophic 21–58 0.070–0.094 19.0–43.3 [21] Tetraselmis chui Phototrophic 17.25–23.5 1.0–2.6 240–440 [26] Tetraselmis sp. Phototrophic 8.2–33.0 0.158–0.214 18.6–22.7 [31] Tetraselmis tetrathele Phototrophic 29.18–30.25 3.1–4.4 920–1340 [26]
Bacillariophyceae Chaetoceros calcitrans CS 178 Phototrophic 39.8 0.04 17.6 [32] Chaetoceros gracilis Phototrophic 15.5–60.28 3.4–3.7 530–2210 [26] Chaetoceros muelleri Phototrophic 11.67–25.25 1.2–2.7 140–670 [26] Nitzschia cf. pusilla YSR02 Phototrophic 48 ± 3.1 0065 31.4 [21] Phaeodactylum tricornutum F&M-M40 Phototrophic 18.7 0.24 44.8 [32] Skeletonema sp. CS 252 Phototrophic 31.8 0.09 27.3 [32] Thalassiosira pseudonana CS 173 Phototrophic 20.6 0.08 17.4 [32]
Others Crypthecodinium cohnii Heterotrophic 19.9 2.236 444.9 [33] Isochrysis sp. Phototrophic 6.5–21.25 0.7–2.7 150–180 [26] Isochrysis sp. Phototrophic 22.0–34.1 0.029–0.090 6.44–21.1 [31] Isochrysis zhangjiangensis Phototrophic 29.8–40.9 0.667–3.1 66.2–140.9 [34] Nannochloropsis oculata Phototrophic 22.75–23.0 2.4–3.4 550–790 [26] Nannochloropsis oculata NCTU-3 Phototrophic 22.7–41.2 0.296–0.497 84–151 [35] Nannochloropsis sp. Phototrophic 21.3–37.8 0.021–0.064 4.59–20.0 [31] Pavlova salina CS 49 Phototrophic 30.9 0.16 49.4 [32] Rhodomonas sp. Phototrophic 9.5–20.5 0.018–0.064 2.06–6.04 [31] Thalassiosira weissflogii Phototrophic 6.25–13.21 0.5–1.5 20 [26]
Table 2 Different carbon sources used for yeast and mold lipid production.
Species Carbon sources Lipid content (% dry weight) Refs.
Yeast Cryptococcus curvatus Glycerol 25 [37] Lipomyces starkeyi Glucose and xylose 61 [38] Rhodosporidium toruloides Y4 Glucose (Batch culture) 48 [39] Rhodosporidium toruloides Y4 Glucose (Fed-batch culture) 67.5 [39] Rhodotorula glutinis Monosodium glutamate wastewater 20 [40] Trichosporon fermentans Glucose 62.4 [41] Yarrowia lipolytica Industrial glycerol 43 [42]
Mold Cunninghamella echinulata Potato starch wastewater 19.03 [43] Cunninghamella echinulata Xylose 57.5 [44] Mortierella isabellina Xylose 65.5 [44] Mortierella isabellina High glucose 50–55 [45] Mucorales fungi Sunflower oil 42.7–65.8 [46] Trichoderma harzianum Q2–37 Corn straw 20.5 ± 0.36 [47]
398 M.-H. Liang, J.-G. Jiang / Progress in Lipid Research 52 (2013) 395–408
2.3. Oleaginous bacteria
Just like fungi, some sort of bacteria also can accumulate oil under some special environment. But usually, the lipid composi- tion produced by bacteria is quite different from other microbial oils. Most bacteria just produce complex lipoid [9], and only a few bacteria can produce oils that can be used as the feedstock for biodiesel production [12]. In bacteria, the most abundant class of neutral lipids are polyhydroxyalkanoic acids serving as intra- cellular carbon and energy storage compounds [48], but also
few examples of substantial TAG accumulation have been reported for species mostly belonging to the actinomycetes genera Mycobacterium [49], Nocardia, Rhodococcus [50] and Streptomyces [51] (Table 3). Recently, it was found that Gordonia sp. and Rhodococcus opacus could accumulate oils under some special conditions with maximum oil content of 80%, but the biomass is only 1.88 g/L [52].
Compared to other microorganisms, many gene regulation mechanisms in fatty acid synthesis in bacteria are already under- stood [53,54]. Therefore, it is relatively easy to use biological
Table 3 TAGs production in bacteria.
Bacteria Carbon sources Lipid content Refs.
Acinetobacter baylyi ADP1(MT) Sodium gluconate and glycerol 12.4% cell dry weight [56] Alcanivorax borkumensis SK2 Pyruvate >23% cell dry weight [57] Gordonia sp. DG Agro-industrial wastes 57.8 mg/L culture medium [52] Mycobacterium tuberculosis H37Rv Limiting nutrient(IL1) 11.9% cell dry weight [58] Nocardia globerula 432 Pristine and acetate >49.7% cell dry weight [59] Rhodococcus opacus PD630 Agro-industrial wastes 88.9 mg/L culture medium [52] Rhodococcus opacus PD630 High glucose 38% cell dry weight [60] Rhodococcus opacus PD630 Sugar beet molasses and sucrose 38.4% cell dry weight [61] Rhodococcus opacus PD630 Glucose >50% cell dry weight [62] Streptomyces coelicolor TR0958 Glucose 83.0 ± 0.5% cell dry weight [63] Streptomyces coelicolor TR0123 Glucose 64 ± 2% cell dry weight [63]
Fig. 4. The fatty acid and TAG biosynthesis pathway in microorganisms. For microalgae, both inorganic carbon (CO2) and organic carbon sources (glucose) can be utilized for lipids production. For yeasts, de novo formation of LPA can occur either through the G3P or DHAP pathways. In yeasts, the DGAT and PDAT catalyze TAG formation. In Acinetobacter calcoaceticus ADP1 (bacteria), WS/DGAT exhibits the DGAT activity. GAP: glyceraldehyde 3-phosphate; DHAP: dihydroxyacetone phosphate; PEP: phosphoenolpyruvate; ACP: acyl-carrier protein; FFA: free fatty acid; G3P: glycerol-3-phosphate; LPA: lysophosphatidate; PA: phosphatidate; DAG: diacylglycerol; CDP- DAG: CDP-diacylglycerol; TAG: triacylglycerol; PDH: pyruvate dehydrogenase; PEPC: phosphoenolpyruvate carboxylase; ME: Malic enzyme; ACL: ATP:citrate lyase; ACC: acetyl-CoA carboxylase; MAT: malonyl-CoA:ACP transacetylase; FAS: fatty acid synthetase; FAT: acyl-ACP-thioesterase; GPAT: glycerol-3-phosphate acyltransferase; LPAT: lysophosphatidate acyl-transferase; PAP: phosphatidic acid phosphatase; DGAT: diacylglycerol acyl-transferase; WS/DGAT: wax ester synthase/acyl-CoA:diacylglycerol acyltransferase; PDAT: phospholipid:diacylglycerol acyltransferase; DHAPAT: DHAP acyltransferase; GPD1 and GUT2: encoding glycerol 3-phosphate dehydrogenase; TGL3 and TGL4: encoding triacylglycerol lipases; POX1–6: encoding the six acyl-CoA oxidases; MFE1: encoding multi-functional enzyme.
M.-H. Liang, J.-G. Jiang / Progress in Lipid Research 52 (2013) 395–408 399
400 M.-H. Liang, J.-G. Jiang / Progress in Lipid Research 52 (2013) 395–408
engineering technology, genetic engineering, and metabolic engi- neering to modify bacteria to improve its oil accumulation. It was reported that a metabolically engineered Escherichia coli could produce biodiesel directly, and the fatty acid esters concentrations of 1.28 g/L was achieved by fed-batch fermentation using renew- able carbon sources [55]. Although the yield was low, it provided a new idea for the biodiesel production.
3. TAG biosynthesis in microorganisms
TAGs, as neutral lipids, are the main materials in the production of biodiesel. The synthesis routes of TAG in microorganisms may consist of the following two approaches: fatty acid synthesis ap- proach and TAG synthesis approach.
3.1. Fatty acid synthesis approach
It is known that both inorganic carbon (CO2) and organic carbon sources (glucose, acetate, etc.) can be utilized by microalgae for lip- ids production. While glucose is also the common source of micro- organisms such as yeasts, molds and microalgae for lipids production. The metabolism flux route on the utilization of carbon dioxide and glucose for the formation of acetyl-CoA in microalgae was described by Yang and Shimizu [64].
Microalgae can perform oxygenic photosynthesis and fix carbon dioxide through Calvin cycle like plant cells. That is, microalgal cells can trap light energy as the energy source and assimilate CO2 as the carbon source. CO2 enters in chloroplast via Calvin cycle to generate glyceraldehyde 3-phosphate (GAP) (Fig. 4). GAP is withdrawn from Calvin cycle and exported to cytoplasm for consumption [64], and then to form pyruvate via glycolytic pathway. Finally, pyruvate is converted to acetyl-CoA by pyruvate dehydrogenase (PDH). When glucose is the carbon source of microorganisms, it can be converted to pyruvate in the cytoplasm after several steps by glycolytic pathway. After entering into the mitochondria, pyruvate is converted to acet- yl-CoA, which condenses with oxaloacetate, a TCA intermediate, to form citrate. When mitochondrial citrate levels are sufficiently
Fig. 5. Citrate-pyruvate shuttle in eukaryotic microorganisms. It is a shuttle for transfer mitochondrion as citrate; in the cytoplasm they are delivered as acetyl-CoA for fatty ATP:citrate lyase; ME: Malic enzyme.
high, citrate enters into cytoplasm, where it is cleaved to form acetyl-CoA and oxaloacetate. This is called citrate-pyruvate shut- tle (Fig. 5).
The elongation of carbon chain of fatty acids is mainly depen- dent on the reaction of two enzyme systems, acetyl-CoA carboxylic enzyme (ACC) and fatty acid synthase (FAS) in most microorgan- isms. From Fig. 4, ACC uses acetyl-CoA for malonyl-CoA formation and acyl chain elongation. Once malonyl-CoA is synthesized, it is transferred by malonyl-CoA:ACP transacetylase (MAT), one of the fatty acid synthase (FAS) multi-enzymatic complex subunits, to form malonyl-acyl-carrier protein (malonyl-ACP). The FAS trans- fers the malonyl moiety to acyl-carrier protein (ACP) to use it as a carbon source for the synthesis of long chain fatty acids, mainly C16 and C18, like palmitic acid (C16:0), stearic acid (C18:0), pal- mitoleic acid (C16:1), oleic acid (C18:1) and linoleic acid (C18:2). Some functional fatty acids like GLA, ARA, EPA, DHA are existed [8]. Each cycle of C2 addition is initiated by a reaction catalyzed by a b-ketoacyl-ACP synthase (KAS) [44] and involves the conden- sation of a malonyl-ACP with an acyl acceptor. At last, acyl-ACP- thioesterase (FAT) cleaves the acyl chain and liberates the fatty acid.
3.2. TAG synthesis approach
For eukaryotes, TAG formation takes place in specialized organelles, like the mitochondria and plastid located in the endoplasmic reticulum. In contrast, the TAG synthesis takes place in the cytoplasm of prokaryotic cells. The most important route to TAG biosynthesis is the G3P or Kennedy pathway (Fig. 4), first described by Professor Eugene Kennedy and his colleagues in the 1950s, by means of which more than 90% of liver TAGs are produced.
The first step of TAG synthesis is the acylation of glycerol-3- phosphate (G3P) with an acyl-CoA to form lysophosphatidate (LPA), which is catalyzed by acyl-CoA:glycerol-sn-3-phosphate acyl-transferase (GPAT). This enzyme exhibits the lowest specific activity in the TAG synthesis pathway, and is suggested to be potentially the rate limiting step [65,66]. The LPA is then further
of acetyl group from mitochondrion to the cytoplasm. Acetyl groups pass out of the acid synthesis. PDH: pyruvate dehydrogenase; PYC: pyruvate carboxylase; ACL:
M.-H. Liang, J.-G. Jiang / Progress in Lipid Research 52 (2013) 395–408 401
condensed, catalyzed by lysophosphatidate acyl-transferase (LPAT), with another acyl-CoA to produce phosphatidate (PA) [67]. Afterwards, PA can be dephosphorylated by phosphatidic acid phosphatase (PAP) to produce diacylglycerol (DAG). At last, synthe- sis of TAG is catalyzed by acyl-CoA:diacylglycerol acyl-transferase (DGAT), which incorporates the third acyl-CoA into DAG. This en- zyme is also known as an important regulator for this pathway [68,69]. It was observed that the deficits in TAG synthesis are asso- ciated with a striking accumulation of DAG, confirming DAG as a critical metabolic branch point in the Kennedy pathway for glycer- ide and glycerophosphatide synthesis [68]. Overexpression of DGAT would commit more DAG to TAG formation rather than phospholipid formation.
It is reported that de novo formation of PA in yeast can occur either through the glycerol-3-phosphate (G3P) or dihydroxyace- tone phosphate (DHAP) pathways. Whereas the former route of PA synthesis is present in bacteria and all types of eukaryotes, the DHAP pathway is restricted to yeast and mammalian cells [67,70,71]. DHAP is acylated at the sn-1 position by DHAP acyl- transferase (DHAPAT), and the product 1-acyl-DHAP is reduced by 1-acyl-DHAP reductase to yield LPA, which is further acylated to PA by LPAT.
Table 4 Researches about lipid synthesis by overexpressing genes or knockout genes.
Genes (enzymes) Source-species Rec
accA-D (ACC), tesA (thioesterase I) E. coli (bacteria) E. c ACC Acinetobacter calcoaceticus (plant) E. c ACC1(ACC) Mucor rouxii (mold) Ha ACC Aspergillus oryzan (mold) Asp ACC1(ACC) Yarrowia lipolytica (yeast) Yar FAT, fabD (MAT) E. coli, Streptomyces avermitilis
MA-4680, Streptomyces coelicolor A3(2) (bacteria)
E. c
KAS III E. coli (bacteria) Bra KAS III Spinacia oleracea (plant) Nic
FAT Umbellularia californica (plant) E. c FAT Diploknema Butyracea (plant) E. c FAT Ricinus communis (plant) E. c FAT Jatropha curcas (plant) E. c GPAT Safflower (plant), E. coli (bacteria) Ara SLC1-1 (LPAT) Yeast Rap D LRO1 (PDAT) Y. lipolytica (yeast) Y. l DGAT Arabidopsis (plant) Yea
GPD1 (GPDH) Yeast Bra
DGUT2 (GPDH) Y. lipolytica (yeast) Y. l GPD1 (GPDH) Y. lipolytica (yeast) Y. l DGUT2 (GPDH) Y. lipolytica (yeast) Y. l GPD1, DGUT2 (GPDH) Y. lipolytica (yeast) Y. l ACS E. coli (bacteria) E. c DME Aspergillus nidulans (mold) Asp ME E. coli K-1 (bacteria) E. c malA (ME) Mortierella alpine, Mucor circinelloides
(mold) Mu
Antisense PEPC Agrobacterium tumefaciens (bacteria) Bra Antisense PEPC Anabaena sp. (microalgae) E. c DAGPase Chlamydomonas (microalgae) Chl DAGPase Chlamydomonas reinhardtii BAFJ5
(microalgae) Chl (m
Multi-gene approach to enhance lipid biosynthesis ACC, thioesterase, D fadD (acyl-CoA
synthetase) E. coli (bacteria), plant E. c
ACP, KAS, FAT Haematococcus pluvialis (microalgae) Ha ACC1, DGAT1 Y. lipolytica (yeast) Y. l POX1-6 (AOXs), MFE1, GPD1, DGUT2 Y. lipolytica (yeast) Y. l
: multiply increased; +: increased; : decreased; D: knockout.
Dahlqvist et al. [72] found that some plants and yeast also have an acyl-CoA-independent mechanism for TAG synthesis, which uses phospholipids as acyl donors and DAG as acceptor. This reac- tion is catalyzed by an enzyme that we call phospholipid:diacyl- glycerol acyltransferase (PDAT). It was also found that Acinetobacter calcoaceticus ADP1 accumulates both wax ester (WE) and TAG, and a bifunctional wax ester synthase/acyl- CoA:diacylglycerol acyltransferase (WS/DGAT) was identified to exhibit both wax ester synthase and DGAT activities [73].
4. Metabolic engineering for TAG production
Numerous studies have been carried out using the metabolic engineering strategy to enhance the lipid accumulation in different species. Some of these studies have been summarized in Table 4 and will be discussed briefly in this section. They can be broadly classified into five different approaches: (1) overexpressing en- zymes of the fatty acid biosynthesis pathway; (2) overexpressing enzymes that enhance the TAG biosynthesis pathway; (3) regula- tion of related TAG biosynthesis bypass approaches; (4) partially blocking competing pathways; and (5) the multi-gene transgenic approach.
eiver-species Note Refs.
oli (bacteria) 6 fatty acid synthesis [74] oli (bacteria) 3 lipid content [75]
nsenula polymorpha (yeast) +40% fatty acid content [76] ergillus oryzan (mold) No significant increase [77] rowia lipolytica (yeast) 2 lipid content [78] oli (bacteria) +11% fatty acid content [79]
ssica napus (plant) +KAS III activity [80] otiana tabacum (plant) 300 KAS III activity, 20% fatty
acid content [81]
oli (bacteria) +fatty acid synthesis [82] oli ML103 (bacteria) >0.2 g/L fatty acid content [83] oli ML103 (bacteria) >2.0 g/L fatty acid content [83] oli ML103 (bacteria) >2.0 g/L fatty acid content [83] bidopsis (plant) +29% oil content [84] eseed, Arabidopsis (plant) +48% oil content [85]
ipolytica (yeast) 40% TAG content [86] st 200–600 DGAT activity,
3–9 TAG content [87]
ssica napus L. (plant) 2 GPDH activity, +40% lipid content
[88]
ipolytica (yeast) 3 lipid content [89] ipolytica (yeast) 1.5 TAG content [90] ipolytica (yeast) 2.9 TAG content [90] ipolytica (yeast) 5.6 TAG content [90] oli (bacteria) 9 ACS activity [91] ergillus nidulans (mold) 50% lipid content [92] oli BL21 (bacteria) 4 lipid content [75] cor circinelloides (mold) 2.5 lipid content [93]
ergillus oryzae (mold) 1.7 fatty acid content, 1.9 TAG content
[77]
ssica napus (plant) +6.4–18% oil content [94] oli DH5a (bacteria) +46.9% lipid content [95] amydomonas (microalgae) 10 TAG content [96] amydomonas reinhardtii BAFJ5 icroalgae)
+46.4%, 3.5 lipid content [97]
ipolytica (yeast) +lipid production [90]
oli (bacteria) 20 fatty acid content [98]
ematococcus pluvialis (microalgae) +fatty acid synthesis [99] ipolytica (yeast) 5 lipid content [78] ipolytica (yeast) +lipid accumulation [90]
402 M.-H. Liang, J.-G. Jiang / Progress in Lipid Research 52 (2013) 395–408
4.1. Enhancement of fatty acid synthesis approach
4.1.1. Acetyl-CoA carboxylase (ACC) As for microorganisms, a previous study showed that co-
expression of E. coli ACC (encoded by accA, accB, accC, accD) and thioesterase I (encoded by the tesA gene) resulted in a 6-fold in- crease in the rate of fatty acid synthesis with 6.6 nmol of free fatty acid, confirming that ACC catalyzing the committing step was in- deed the rate-limiting step for fatty acid biosynthesis in this strain [74]. However, expressing the E. coli ACC alone did not cause an obvious increase in fatty acid production, which indicated that steps later in the pathway limit the flux through the pathway. This could be because the overexpression of native E. coli ACC suffered from feedback inhibition by acyl-ACP [100]. tesA may reduce this inhibition by forming free fatty acid, resulting in the increased pro- duction of fatty acid. Another way to reduce this inhibition could be to express a non-native ACC gene that is not recognized by the acyl-ACP of E. coli. For example, the heterologous expression of ACC from Acinetobacter calcoaceticus in E. coli caused a 3-fold enhancement in lipid levels and thus provided support for the assumption [75].
In eukaryotic microorganisms, overexpression of ACC has been met with only limited improvement of lipid production. The ACC1 enzyme is responsible for providing the malonyl-CoA used in cytoplasmic fatty acid synthesis. Heterologous expression of ACC1 from the oleaginous fungus Mucor rouxii in the non-oleagi- nous yeast Hansenula polymorpha was able to achieve only a 40% increase in total fatty acid content [76]. In addition, the ACC-en- hanced expression in Aspergillus oryza also did not show a signifi- cant increase in productivities of either fatty acids or TAGs relative to the parental strain [77]. It was suspected that improve- ments in total lipid accumulation have been limited in eukaryotes by the strong metabolic and regulatory control maintained over this enzyme, possibly by free fatty acids. However, Yarrowia lipoly- tica might represent a regulatory exception in eukaryotic microor- ganisms, lending much to its oleaginous nature, as Tai et al. [78] mentioned that a 2-fold increase was achieved in lipid content through a commensurate overexpression of endogenous ACC1.
4.1.2. Fatty acid synthetase (FAS) The first committed step of fatty acid biosynthesis is the conver-
sion of acetyl-CoA to malonyl-CoA by an ATP-dependent ACC fol- lowed by the conversion of malonyl-CoA to malonyl-ACP through the enzyme malonyl CoA:ACP transacylase (MAT). The three E. coli strains carrying acyl-ACP thioesterase (FAT) and a fabD gene encoding MAT from E. coli, Streptomyces avermitilis MA-4680, or Streptomyces coelicolor A3(2) improved the free fatty acid produc- tion, about 11% more than the control strains, suggesting that fabD overexpression could be used to improve free fatty acid production by increasing the malonyl-ACP availability [79].
b-ketoacyl-ACP synthase III (KAS III) condenses acetyl-CoA with malonyl-ACP to initiate fatty acid biosynthesis. It was defined as the kinetic mechanism that underlay the negative regulation of KAS III activity long-chain acyl-ACP [101]. An E. coli KAS III was overexpressed in the Brassica napus [80], causing a significant in- crease in KAS III activity. However, the fatty acid profile of the stor- age lipids was affected, which decreased the amounts of C18:1 and increased the amounts of C18:2 and C18:3 as compared to control plants. Such changes in fatty acid composition reflected changes in the regulation and control of fatty acid biosynthesis, proposing that fatty acid biosynthesis was not controlled by one rate-limiting en- zyme, such as ACC, but rather was shared by a number of compo- nent enzymes of the fatty acid biosynthetic machinery. Recently, a spinach (Spinacia oleracea) KAS III was expressed in tobacco (Nico- tiana tabacum) and resulted in a 300-fold increase in activity above the wild type. However, rather than an increase in fatty acid con-
tent, a 20% decrease was observed [81]. An interesting and unex- pected consequence of KAS III overexpression was an increase in ACP levels in tobacco leaves, although other FAS activities were unaffected. Decreases in fatty acid content as a result of KAS III overexpression were attributed to the decreased rates of de novo fatty acid synthesis most likely by the reduced malonyl-CoA pools for subsequent KAS condensation reactions.
In microalgae, main key genes like ACP, KAS and FAT genes overexpression in Haematococcus pluvialis for fatty acid biosynthe- sis had significant correlations with monounsaturated fatty acid (MUFA) synthesis and polyunsaturated fatty acid (PUFA) synthesis, proposing that ACP, KAS, and FAT in H. Pluvialis may play an impor- tant role in fatty acid synthesis and may be rate limiting genes, which probably could be modified for the further study of meta- bolic engineering to improve the quality and production of micro- algae biofuel [99].
It seems that the subunits of FAS are challenging targets for metabolic engineering for fatty acid metabolism enhancement, probably due to the fact that FAS is a multi-enzymatic complex containing subunits whose activities depend on one another. The difficulties experienced with the heterologous expression of mul- ti-enzymatic complexes such as FAS were also likely due to the dif- ferences in multipoint controls among different species.
4.1.3. Acyl-ACP-thioesterase (FAT) FATs are a group of enzymes that catalyze the hydrolysis of
acyl-ACPs to form the free fatty acids and ACP. Free fatty acids can be produced by introducing an FAT gene. The presence of the FAT will break the fatty acid elongation cycle and release free fatty acids [98,102].
It was reported that the expression of the plant (Umbellularia californica) enzyme medium-chain FAT in E. coli deficient in fatty acid degradation could result in a very high level hydrolytic activ- ity and cause a minor accumulation of medium-chain fatty acids, which suggested that acyl-ACP intermediates might normally act as feedback inhibitors for fatty acid synthsis [82]. It was examined the effect of different FAT on the quantities and compositions of free fatty acid produced by an E. coli strain ML103 carrying FAT gene from four different plants. The strain carrying the FAT genes from Diploknema Butyracea produced the quantity of free fatty acid (>0.2 g/L) while the strains carrying FAT genes from Ricinus com- munis and Jatropha curcas produced the most free fatty acid, more than 2.0 g/L at 48 h. These two strains accumulated three major straight chain free fatty acids, C14, C16:1 and C16 at levels about 40%, 35% and 20%, respectively [83]. It was shown that the amount of free fatty acid accumulated depends on the acyl-ACP thioester- ase used.
4.2. Enhancement of TAG synthesis approach
4.2.1. Acyl-CoA:glycerol-sn-3-phosphate acyl-transferase (GPAT) GPAT catalyses the first reaction of TAG synthesis to form LPA
via the Kennedy pathway. LPA can also be formed by the acylation of DHAP by acyl-CoA:DHAP acyltransferase (DHAPAT) followed by reduction of the newly formed 1-acyl-DHAP to LPA by DHAP reduc- tase (Fig. 4). S. cerevisiae GPAT mutants are equally defective in DHAPAT activity, thereby establishing the genetic identity of the two activities in yeast [103]. In S. cerevisiae, GAT1 and GAT2 (SCT1) genes encode the major GPAT activities. In vitro substrate specificities were determined and the GAT1 gene product could use both G3P and DHAP with similar efficiencies and had a broad fatty acyl-CoA specificity profile, whereas the GAT2 gene product preferred G3P over DHAP and had a 2.5- to 5-fold preference for C16 fatty acyl chains [104]. Metabolic studies show that TAG bio- synthesis increases 50% in DGAT1 yeast and decreases by 50% in
M.-H. Liang, J.-G. Jiang / Progress in Lipid Research 52 (2013) 395–408 403
DGAT2 yeast, indicating that GAT2 initiates the major route for TAG biosynthesis [105].
It was reported that the oil content of Arabidopsis seeds was in- creased from 8% to 29% in selected transgenic lines by expressing a plastidial safflower GPAT and an E. coli GPAT gene, demonstrating that expression of both a bacterial and a plant GPAT gene increased plant seed oil content and seed weight [84].
4.2.2. Lysophosphatidate acyl-transferase (LPAT) LPA is further acylated by LPAT to form PA. The enhancement of
seed TAG content by a microsomal LPAT homologous to the endog- enous enzyme is remarkable because such an effect has been ob- served only when the yeast SLC1-1 gene, encoding a variant LPAT, which can be used to change total fatty acid content and composition as well as to alter the stereospecific acyl distribution of fatty acids in seed TAGs [106], was used to transform rapeseed and Arabidopsis [85], leading to increases from 8% to 48% seed oil content on the seed dry weight basis, suggesting that increasing the expression of glycerolipid acyltransferases in seeds leads to a greater flux of intermediates through the Kennedy pathway and re- sults in enhanced TAG accumulation.
4.2.3. Acyl-CoA:diacylglycerol acyl-transferase (DGAT) DGAT catalyzes, as discussed previously, the last step of TAG
formation to form TAG from DAG and fatty acyl CoA. The DGAT1 or DGAT2 families present in yeast, plants, and animals, and the phospholipid:diacylglycerol acyltransferase (PDAT) catalyzes TAG formation in yeast and plants. Yeast LRO1 is incapable of synthesiz- ing sterols and encodes the PDAT. Deletion of LRO1 encoding PDAT in Y. lipolytica resulted in a 40% loss of TAG [86]. Another experi- ment with a knockout mutant demonstrated the key role of the bifunctional WS/DGAT for biosynthesis of both storage lipids (TAGs and WEs) in A. calcoaceticus ADP1 [107]. The WS/DGAT enzymes in prokaryotes are not related to any known acyltransferase involved in the formation of TAGs and WEs in eukaryotes, including the DGAT1 [108] and DGAT2 [69,87] families, the WS of the jojoba plant (Simmondsia chinensis) [109] or the acyl-CoA independent PDAT (LRO1) catalyzing TAG formation in yeast and plants [72].
It was reported that transformations of yeast with the Arabidop- sis DGAT were performed. A 200–600-fold increase of DGAT activ- ity in the transformed yeast was observed, which led to a 3–9-fold increase of TAGs accumulation [87].
The success with DGAT could be explained by the fact that the substrate of DGAT, DAG, could be allocated to either phospholipid biosynthesis or TAG formation. Overexpression of DGAT would commit more DAG to TAG formation rather than phospholipid for- mation. Olukoshi and Packter [51] have reported a strong correla- tion between DGAT activity, growth phase and TAG content in cells of Streptomyces sp., and they suggested that this enzyme is proba- bly responsible for the switch from membrane phospholipid for- mation to TAG biosynthesis during the stationary growth phase. All results above seem to suggest that the reaction catalyzed by DGAT is an important rate-limiting step in lipid biosynthesis.
4.2.4. Glycerol 3-phosphate dehydrogenase (GPDH) The GPDH isoform involved in reduction of DHAP into G3P is
GPD1. G3P provides the activated glycerol backbone for TAG syn- thesis. A yeast gene coding for cytosolic GPD1 was expressed in transgenic oil-seed rape (B. napus L.) under the control of the seed-specific promoter. It was found that a 2-fold increase in GPDH activity led to a 3–4-fold increase in the level of G3P in developing seeds, resulting in a 40% increase in the final lipid content of the seed [88]. The GUT2 gene coding for another GPDH isomer, which catalyzes the dihydroxyacetone phosphate (DHAP) formation from G3P was deleted in Y. lipolytica in order to boost G3P availability, leading to a 3-fold increase in lipid accumulation compared to
the wild-type strain [89]. It was observed that GPD1 overexpres- sion, GUT2 inactivation or both mutations of Y. lipolytica together result in 1.5-, 2.9- and 5.6-fold respective increases in the level of G3P leading to an increase of TAG accumulation [90].
4.3. Regulation of related TAG biosynthesis bypass approaches
A few enzymes that are not directly involved in lipid metabo- lism have also been demonstrated to influence the rate of lipid accumulation by increasing the pool of essential metabolites for li- pid biosynthesis. The following are a few examples.
4.3.1. Acetyl-CoA synthase (ACS) ACS catalyzes the conversion of acetate into acetyl-CoA. The
metabolic utilization of acetate can contribute to lipid biosynthesis or oxidation via the TCA cycle [110]. The overexpression of ACS in E. coli showed significant reduction in acetate during glucose metabolism. It also greatly enhanced the assimilation of acetate when used as the sole carbon source. Increased ACS levels presum- ably enhanced the activation of acetate to acetyl-CoA, which may increase the rate of fatty acid synthesis. For instance, it was ob- served that, by overexpressing the ACS gene in E. coli, the ACS activity was increased by 9-fold, leading to a significant increase of the assimilation of acetate from the medium, which can contrib- ute to lipid biosynthesis [91].
4.3.2. Malic enzyme (ME) ME carries out the irreversible decarboxylation of malate to
pyruvate, accompanied by the formation of NADPH, which is needed for fatty acid biosynthesis. It has been suggested that the function of ME in lipid biosynthesis is to supply NADPH for fatty acid synthase and desaturases [111,112]. A mutant (scuK248) lack- ing ME activity accumulated only half the lipid (l2% of cell dry weight) accumulated by strains of Aspergillus nidulans possessing ME [92].
The effect of ME was studied in filamentous fungi in correlation with lipid accumulation [113]. It was observed that the enhanced energy (NADPH) supply as a result of ME overexpression was uti- lized by the enzymes involved in TAG synthesis and led to en- hanced lipid production. It was observed that the enhanced activity of ME led to the increase of the cytosolic NADPH pool, making available more reducing power for lipogenic enzymes such as ACC, FAS and ATP:citrate lyase (ACL). It was reported that pro- viding a high level of NADPH by overexpressing ME in E. coli and adding malate to the culture medium resulted in a 4-fold increase in intracellular lipids [75].
The contribution of malA encoded for the isoforms III and IV of ME to lipid accumulation has been further supported because over- expression of the gene coding for them led to a 2.5-fold increase of lipid accumulation in Mucor circinelloides in comparison with a leu- cine auxotrophic strain, from 12% of the biomass to 30% [93]. How- ever, ME activity still disappeared by the end of the lipid- accumulation phase. The possibility existed that a second bottle- neck, i.e., another limiting step in the fatty acid synthesis pathway, may have arisen after eliminating the bottleneck caused by the low ME activity. It provided proofs that a fully operating leucine bio- synthetic pathway was required for the accumulation of lipids, inferring that endogenously produced leucine was degraded for the generation of the acetyl-CoA, which could be incorporated into fatty acid biosynthesis [114].
4.3.3. ATP:citrate lyase (ACL) ACL catalyzes the conversion of citrate to acetyl-CoA and oxalo-
acetate and is a key enzyme for lipid accumulation in mammals and oleaginous yeasts and fungi. The specific activity of ACL en- zyme correlates with the specific rate of lipid synthesis, and it
O O
PEPC Activate
404 M.-H. Liang, J.-G. Jiang / Progress in Lipid Research 52 (2013) 395–408
may be inferred that the enzyme is possibly the rate-limiting reac- tion for lipid biosynthesis [115]. Recently, a great increase in pro- ductivity was seen in the ACL-enhanced expression in Aspergillus oryzae, where a 1.7-fold increase in the productivities of fatty acids and 1.9-fold increase of TAG relative to the parental strain were ob- served [77].
SCoA
O
O
OHO
O
HO
4.4. Blocking competing pathways
From the metabolic engineering point of view, blocking off competing pathways may also enhance the metabolic flux being channeled to TAG biosynthesis.
TCA cycle
SCoA
Malonyl-CoA
Fig. 6. Possible role of PEPC in the fatty acid biosynthesis pathway. PEPC: phosphoenolpyruvate carboxylase; ACC: acetyl-CoA carboxylase.
4.4.1. Repression of b-oxidation A complementary strategy to increase lipid accumulation is to
decrease lipid catabolism. Genes involved in the activation of both TAGs and free fatty acids synthesis, as well as genes directly in- volved in b-oxidation of fatty acids to be inactivated, sometimes result in the increase of cellular lipid content.
Several reports have shown that knocking out genes involved in b-oxidation in S. cerevisiae not only can lead to increased amounts of intracellular free fatty acids but also results in extracellular fatty acid secretion in some instances [103,116]. Dysfunction of AOX in S. cerevisiae leading to the inactivation of the b-oxidation process will result in the accumulation of cellular fatty acids. S. cerevisiae (nonoleaginous yeast) produces a relatively low level of lipids. It was explained that nonoleaginous yeasts may not be able to accu- mulate the substrate due to a feedback regulation of the oxidation process which would affect hydrophobic substrates (e.g., alkanes, fatty acids, and oils) transporters and down-regulate the diffusion process. In addition, lower levels of ATP deriving from b-oxidation were produced in these strains, and may be used in pathways other than lipid storage [86].
While Y. lipolytica (oleaginous yeast) is able to accumulate lipids to levels exceeding 50% of cell dry weight [117]. It contains six AOXs, encoded by the POX1 to POX6 genes, that catalyze the limit- ing step of peroxisomal b-oxidation. Aox2p expression in Y. lipoly- tica regulates the size of cellular TAG pools and the size and number of lipid bodies in which these fatty acids accumulate [118], suggesting the existence of a regulatory mechanism in strains with altered POX genotype. The mobilization of lipid sup- plies is regulated by the presence of the AOX proteins. Modifica- tions of the POX genotype is useful in preventing lipid degradation and therefore leads indirectly to an increase in lipid accumulation [89].
As for many microalgae, during diel light–dark cycles, initiate TAG storage during the day and deplete those stores at night to support cellular ATP demands and cell division. Consequently, inhibition of b-oxidation would prevent the loss of TAG during the night, but most likely at the cost of reducing growth. This strat- egy, therefore, may not be beneficial for microalgae grown in out- door open ponds, but it may be a valid strategy to increase lipid production in microalgae grown in photobioreactors with exoge- nous carbon sources and continuous light.
4.4.2. Repression of phospholipid biosynthesis Phospholipid biosynthesis is another competitive pathway to
TAG formation because it competes against TAG biosynthesis for a common substrate, PA. If PA is converted into CDP-diacylglycerol instead of DAG (Fig. 4), it enters the phospholipids synthetic path- way [66]. It was shown that inhibition of phospholipid synthesis caused the formation of abnormally long fatty acids, due to supple- mentary elongation cycles [102].
4.4.3. Repression of phosphoenolpyruvate carboxylase (PEPC) The third competitive pathway is the conversion of phospho-
enolpyruvate (PEP) to oxaloacetate, which is catalyzed by PEPC. TAG biosynthesis requires PEP (which converts successively to pyruvate, acetyl-CoA, malonyl-CoA and then fatty acids) [119]. PEPC also plays a key role in photosynthesis. Besides plants, the en- zyme is also found in photoautotrophic microalgae like cyanobac- teria, but not in animals or fungi [120]. That is to say, the biosyntheses of either protein or lipid take PEP as the common substrate. It is converted into oxaloacetate by PEPC and then di- rectly participates in the anabolism of protein. On the other hand, PEP can be converted into pyruvate by the pyruvate kinase. Pyru- vate is then transformed to acetyl-CoA by pyruvate dehydrogenase, finally participated in the anabolism of fatty acid. While the activ- ity of PEPC is reduced by antisense inhibition, the concentration of PEP increases to high level to help to the formation of more pyru- vate. Meanwhile, along with consumption of acetyl-CoA for the formation of malonyl-CoA, the low level of acetyl-CoA is able to activate the biochemical process of converting PEP to pyruvate (Fig. 6).
Many evident roles of PEPC in lipids biosynthetic pathways in oilseed crops have been reported. By expressing antisense PEPC in B. napus [94], it was achieved a 6.4–18% increase in oil content, suggesting that reducing PEPC activity enhanced the lipid accumu- lation. Significantly enhanced lipid contents were also obtained with transgenic soybean lines harbouring anti-PEPC gene [121,122]. Therefore, the antisense expression of PEPC gene for the modulation of fatty acids biosynthesis in oil crops is an effec- tive method.
In microalgae, preliminary results also indicated that PEPC plays a role in the regulation of fatty acid accumulation and reduced PEPC activity by antisense expression was correlated with a 46.9% increase of the lipid content in Anabaena sp., a cyanobacte- rium [95], showing a great progress in microalgae engineering for biodiesel production.
4.4.4. Repression of starch biosynthesis In many microalgal cells, starch is another major carbon and en-
ergy storage compound, particularly under stress conditions. The first committed step of starch synthesis (Fig. 7) in the plastid is cat- alyzed by ADP-glucose pyrophosphorylase (AGPase), which con- verts glucose 1-phosphate and ATP to ADP-glucose and Pi. Subsequently, ADP-glucose is used by starch synthases (SS) and branching enzymes (BE) to elongate the glucan chains of the starch granule [123].
OH
OH
H
Fig. 7. Starch biosynthesis pathway in chloroplast. GAP: glyceraldehyde-3-phosphate; G-6-P: glucose 6-phosphate; G-1-P: glucose 1-phosphate; Pi: inorganic pyrophosphate; AGPase: ADP-Glucose pyrophosphate; SS: starch synthase; BE: branching enzymes.
M.-H. Liang, J.-G. Jiang / Progress in Lipid Research 52 (2013) 395–408 405
The model green microalgae Chlamydomonas reinhardtii has re- cently emerged as a model to test genetic engineering or cultiva- tion strategies aiming at increasing lipid yields for biodiesel production. Blocking starch synthesis has been suggested as a way to boost oil accumulation. Recently, it was reported that inac- tivation of AGPase in a Chlamydomonas starchless mutant led to a 10-fold increase in TAG, suggesting that shunting of photosyn- thetic carbon partitioning from starch to TAG synthesis may repre- sent a more effective strategy than direct manipulation of the lipid synthesis pathway to overproduce TAG [96]. BAFJ5, one of the mu- tants defective in the small subunit of AGPase in C. reinhardtii, accumulated neutral and total lipid of up to 32.6% and 46.4% of dry weight or 8- and 3.5-fold higher, respectively, than the wild- type [97]. These results confirmed the feasibility of increasing lipid production through redirecting photosynthetically assimilated car- bon away from starch synthesis to neutral lipid synthesis. How- ever, some growth impairment was observed in the low starch and starchless mutants. Starch may play an important role in maintaining high photosynthetic efficiency. Shifting carbon flux from starch to the lipid synthesis pathway may have a negative ef- fect on photosynthesis in algae, resulting in impairment in growth in starchless and other low starch mutants.
4.4.5. Repression of the degradation of TAG It was reported that the high levels of lipids resulted from the
repression of genes (TGL3 and TGL4, encoding triacylglycerol li- pases) in Y. lipolytica involved in the degradation of TAG [90], sug- gesting that it is a feasible approach to increase lipid production.
4.5. The multi-gene approach
The multi-gene approach, i.e., overexpressing more than one key enzyme in the TAG pathway to enhance lipid biosynthesis, was suggested by a few researchers [80,124]. Until now, there are more and more literatures on the feasibility of this strategy, probably due to the effectiveness in manipulating multiple genes.
For example, a comprehensive modification of E. coli, which re- sulted in a 20-fold increase in free fatty acid production, overex- pressed the lipid biosynthesis genes encoding ACC, an endogenous thioesterase, and a plant thioesterase, as well as knocked out a gene involved in b-oxidation of fatty acids, acyl- CoA synthetase (encoded by fadD) [98]. As for microalgae, the
key rate-limiting genes of fatty acid synthesis may include ACP, KAS and FAT because their expression showed linear relationships with synthesis of fatty acids in H. pluvialis. These genes could be potential candidates for better quality and higher production of fatty acids for biofuel using metabolic engineering techniques [99].
Co-overexpression of two important genes in the lipid synthesis pathway, ACC1 and DGAT1, in oleaginous yeast Y. lipolytica, pro- vided an enhanced driving force towards the production of lipids, resulting that ACC1 + DGAT1 strain was able to accumulate up to 62% of its cell dry weight, almost 5-fold greater than the control [78]. The enhanced lipid accumulation observed in the strains co- expressing ACC1and DGAT1 is presumably due to a better balance between the fatty acid and TAG synthesis pathways. Acyl-CoA intermediates function as both product and feedback inhibitors in the fatty acid (upstream) pathway and primary precursors in the TAG (downstream) pathway. Upregulation of the upstream pathway increases the throughput of fatty acid synthesis. Upregu- lation of the downstream pathway creates a driving force by depleting acyl-CoA intermediates and increasing the rate of storage of TAG in lipid bodies. Indeed, coupling precursor overproduction and driving forces with a metabolic sink to enable a push and pull dynamic has become a very powerful strategy in recent efforts of metabolic engineering, particularly for biofuels [125–127].
Another example, the genes POX1-6 encoded the six AOXs of Y. lipolytica catalyze the first and rate-limiting step of b-oxidation. And the gene MFE1 encodes the multi-functional enzyme, involved in the second step of the b-oxidation pathway. It was reported that the inactivation of POX1-6 or MFE1 strains in which GPD1 was over- expressed or GUT2 was inactivated, showed that G3P accumulation affects lipid accumulation mostly in strains that are defective for b- oxidation [90]. The finding indicated that TAG synthesis is limited by the availability of G3P and fatty acids, and that the expression of genes involved in TAG homeostasis is regulated by the G3P shuttle and the b-oxidation pathway.
From the above mentioned, multi-gene approach is shown a great prospect in lipid accumulation for biodiesel production.
5. Conclusion
Taken together, increasing the quantity of any of the Kennedy pathway acyltransferases results in elevated TAG content consis-
406 M.-H. Liang, J.-G. Jiang / Progress in Lipid Research 52 (2013) 395–408
tent with an augmentation of the pool of LPA, PA, and DAG inter- mediates and a feed-forward enhancement of storage lipid sink size. Apparently, the supply of precursor fatty acyl groups and glyc- erol-3-phosphate responds via feedback signaling to meet this in- creased demand. The modest increases in TAG content attained by increasing the supply of fatty acids, for example, via the engineer- ing of an increased expression of ACC in the plastid [124], com- pared with the substantial augmentation of TAG accumulation achieved by overexpression of Kennedy pathway acyltransferases may indicate that fatty acid utilization represents the more impor- tant limitation on storage lipid accumulation. This inference is sup- ported by the studies on metabolic control analysis conducted in rapeseed [128], which suggested that a greater level of control was exerted at the level of TAG assembly than at fatty acid synthe- sis [129].
As there are quite a few success stories in lipid overproduction using transformed microbial strains, the knowledge obtained in studies on lipid pathways and genetic transformed organisms for enhanced lipid synthesis among various species suggests that DGAT and ME are the most promising targets for gene transforma- tion. Of particular interest, reducing PEPC activity by expressing antisense gene was observed to be beneficial for lipid production in microalgae. As for multi-gene approach, more trials of this strat- egy should be done due to its great effectiveness. Since oil crisis in the mid 1970s, finding new energy resources to replace petroleum has been a hot topic worldwide. Because of the many advantages over the conventional energy resources, the production of biodiesel has attracted much attention in recent years. We believe that in the near future, research into lipid metabolism of microorganisms will advance rapidly and lipid content will increase rapidly by using metabolic engineering technology.
Acknowledgements
This project was supported by National Natural Foundation of China (Grant 31171631).
References
[1] Liu B, Sun Y, Liu YH, Zhao ZB. Progress on microbial glyceride biosynthesis and metabolic regulation in oleaginous microorganisms. Acta Microbiol Sin 2005;45:153–6.
[2] Kulkarni MG, Dalai AK. Waste cooking oils – an economical source for biodiesel: a review. Ind Eng Chem Res 2006;45:2901–13.
[3] Gerpen JV. Business management for biodiesel producers. NREL Technical, Report 2004; NREL/SR-510-36242:175.
[4] Renewables 2011; GLOBAL STATUS REPORT. [5] Biodiesel. Vegetable oil yields. <http://journeytoforever.org/
biodiesel_yield.html>. [6] Michael B. Widescale Biodiesel production from algae. University of New
Hampshire (US) Biodiesel Group, 2004; <http://www.resilience.org/stories/ 2004-10-03/widescale-biodiesel-production-algae>.
[7] State Fact Sheets: United States. Economic Research Service, USDA, 2013; <http://www.ers.usda.gov/data-products/state-fact-sheets/state- data.aspx?StateFIPS=00#FC>.
[8] Ma YL. Microbial oils and its research advance. Chin J Bioprocess Eng 2006;4:7–11.
[9] Yi SJ, Zheng YP. Research and application of oleaginous microorganism. China Foreign Energy 2006;11:90–4.
[10] Meng X, Yang JM, Xu X, Zhang L, Ni QJ, Xian M. Biodiesel production from oleaginous microorganisms. Renew Energy 2009;34:1–5.
[11] Li Q, Du W, Liu DH. Perspectives of microbial oils for biodiesel production. Appl Microbiol Biotechnol 2008;80:749–56.
[12] Xue FY, Zhang X, Tan TW. Research advanceand prospect in microbial oils. Chin J Bioprocess Eng 2005;3:23–7.
[13] Chisti Y. Biodiesel from microalgae. Biotechnol Adv 2007;25:294–306. [14] Metting FB. Biodiversity and application of microalgae. J Ind Microbiol
Biotechnol 1996;17:477–89. [15] Spolaore P, Joannis-Cassan C, Duran E, Isambert A. Commercial applications of
microalgae. J Biosci Bioeng 2006;101:87–96. [16] Zhang YW, Liu W. Advances in the research of microalgae bioenergy. Mar Sci
2012;36:132–8.
[17] Xiong W, Li XF, Xiang JY, Wu QY. High-density fermentation of microalga Chlorella protothecoides in bioreactor for microbio-diesel production. Appl Microbiol Biotechnol 2008;78:29–36.
[18] Miao XL, Wu QY. High yield bio-oil production from fast pyrolysis by metabolic controlling of Chlorella protothecoides. J Biotechnol 2004;110:85–93.
[19] Miao XL, Wu QY. Bio-oil fuel production from microalgae after heterotrophic growth. Renew Energy 2004;4:41–4.
[20] Dunahay TG, Jarvis EE, Zeiler KG, Roessler PG, Brown LM. Genetic engineering of microalgae for fuel production. Appl Biochem Biotechnol 1992;34:331–9.
[21] Abou-Shanab RAI, Hwang JH, Cho Y, Min B, Jeon BH. Characterization of microalgal species isolated from fresh water bodies as a potential source for biodiesel production. Appl Energy 2011;88:3300–6.
[22] Gao CF, Zhai Y, Ding Y, Wu QY. Application of sweet sorghum for biodiesel production by heterotrophic microalga Chlorella protothecoides. Appl Energy 2010;87:756–61.
[23] Hsieh CH, Wu WT. Cultivation of microalgae for oil production with a cultivation strategy of urea limitation. Bioresour Technol 2009;100:3921–6.
[24] Feng YJ, Li C, Zhang DW. Lipid production of Chlorella vulgaris cultured in artificial wastewater medium. Bioresour Technol 2011;102:101–5.
[25] Liang Y, Sarkany N, Cui Y. Biomass and lipid productivities of Chlorella vulgaris under autotrophic, heterotrophic and mixotrophic growth conditions. Biotechnol Lett 2009;31:1043–9.
[26] Araujo GS, Matos LJ, Goncalves LR, Fernandes FA, Farias WR. Bioprospecting for oil producing microalgal strains: evaluation of oil and biomass production for ten microalgal strains. Bioresour Technol 2011;102:5248–50.
[27] Damiani MC, Popovich CA, Constenla D, Leonardi PI. Lipid analysis in Haematococcus pluvialis to assess its potential use as a biodiesel feedstock. Bioresour Technol 2010;101:3801–7.
[28] Li YQ, Horsman M, Wang B, Wu N, Lan CQ. Effects of nitrogen sources on cell growth and lipid accumulation of green alga Neochloris oleoabundans. Appl Microbiol Biotechnol 2008;81:629–36.
[29] Gouveia L, Marques AE, da Silva TL, Reis A. Neochloris oleabundans UTEX #1185: a suitable renewable lipid source for biofuel production. J Ind Microbiol Biotechnol 2009;36:821–6.
[30] Li YT, Han DX, Sommerfeld M, Hu QA. Photosynthetic carbon partitioning and lipid production in the oleaginous microalga Pseudochlorococcum sp. (Chlorophyceae) under nitrogen-limited conditions. Bioresour Technol 2011;102:123–9.
[31] Huerlimann R, de Nys R, Heimann K. Growth, lipid content, productivity, and fatty acid composition of tropical microalgae for scale-up production. Biotechnol Bioeng 2010;107:245–57.
[32] Rodolfi L, Zittelli GC, Bassi N, Padovani G, Biondi N, Bonini G, et al. Microalgae for oil: strain selection, induction of lipid synthesis and outdoor mass cultivation in a low-cost photobioreactor. Biotechnol Bioeng 2009;102:100–12.
[33] Couto RM, Simoes PC, Reis A, Da Silva TL, Martins VH, Sanchez-Vicente Y. Supercritical fluid extraction of lipids from the heterotrophic microalga Crypthecodinium cohnii. Eng Life Sci 2010;10:158–64.
[34] Feng DN, Chen ZA, Xue S, Zhang W. Increased lipid production of the marine oleaginous microalgae Isochrysis zhangjiangensis (Chrysophyta) by nitrogen supplement. Bioresour Technol 2011;102:6710–6.
[35] Chiu SY, Kao CY, Tsai MT, Ong SC, Chen CH, Lin CS. Lipid accumulation and CO(2) utilization of Nannochloropsis oculata in response to CO(2) aeration. Bioresour Technol 2009;100:833–8.
[36] Tang SH, Chen MK, Yang JB, Ni Q, He DP, Chen T. Research of producing oil by Mortierella isabellina. China Oil Fat 2007;32:35–7.
[37] Meesters PAEP, Huijberts GNM, Eggink G. High-cell-density cultivation of the lipid accumulating yeast Cryptococcus curvatus using glycerol as a carbon source. Appl Microbiol Biotechnol 1996;1996:575–9.
[38] Zhao X, Kong XL, Hua YY, Feng B, Zhao ZB. Medium optimization for lipid production through co-fermentation of glucose and xylose by the oleaginous yeast Lipomyces starkeyi. Eur J Lipid Sci Technol 2008;110:405–12.
[39] Li YH, Zhao ZB, Bai FW. High-density cultivation of oleaginous yeast Rhodosporidium toruloides Y4 in fed-batch culture. Enzyme Microb Technol 2007;41:312–7.
[40] Xue FY, Miao JX, Zhang X, Luo H, Tan TW. Studies on lipid production by Rhodotorula glutinis fermentation using monosodium glutamate wastewater as culture medium. Bioresour Technol 2008;99:5923–7.
[41] Zhu LY, Zong MH, Wu H. Efficient lipid production with Trichosporon fermentans and its use for biodiesel preparation. Bioresour Technol 2008;99:7881–5.
[42] Papanikolaou S, Aggelis G. Lipid production by Yarrowia lipolytica growing on industrial glycerol in a single-stage continuous culture. Bioresour Technol 2002;82:43–9.
[43] Wang HX, Deng ZS, Zhou S, Zhang XY. Using potato starch waste water to produce GLA with Cunninghamella echinulata. China Oil Fat 2007;32:49–51.
[44] Fakas S, Papanikolaou S, Batsos A, Galiotou-Panayotou M, Mallouchos A, Aggelis G. Evaluating renewable carbon sources as substrates for single cell oil production by Cunninghamella echinulata and Mortierella isabellina. Biomass Bioenergy 2009;33:573–80.
[45] Papanikolaou S, Komaitis M, Aggelis G. Single cell oil (SCO) production by Mortierella isabellina grown on high-sugar content media. Bioresour Technol 2004;95:287–91.
M.-H. Liang, J.-G. Jiang / Progress in Lipid Research 52 (2013) 395–408 407
[46] Certik M, Baltészov L, Šajbidor J. Lipid formation and c-linolenic acid production by Mucorales fungi grown on sunflower oil. Lett Appl Microbiol 1997;25:101–5.
[47] Wang X, Hu JD, Zhang XJ, Ren Y, Yang HT. Screening on Trichoderma isolates producing lipid by decomposing corn straw. Chn Brw 2012;31:154–8.
[48] Steinbüchel A. Polyhydroxyalkanoic acids. Biomaterials 1991:123–213. [49] Barksdale L, Kim KS. Mycobacterium. Bacteriol Rev 1977;41:217–372. [50] Alvarez HM, Kalscheuer R, Steinbuchel A. Accumulation of storage lipids in
species of Rhodococcus and Nocardia and effect of inhibitors and polyethylene glycol. Fett/Lipid 1997;99:239–46.
[51] Olukoshi ER, Packter NM. Importance of stored triacylglycerols in Streptomyces: possible carbon source for antibiotics. Microbiology 1994;140:931–43.
[52] Gouda MK, Omar SH, Aouad LM. Single cell oil production by Gordonia sp. DG using agro-industrial wastes. World J Microbiol Biotechnol 2008;24:1703–11.
[53] Wentzel A, Ellingsen TE, Kotlar H, Zotchev SB, Throne-Holst M. Bacterial metabolism of long chain n-alkanes. Appl Microbiol Biotechnol 2007;76:1209–21.
[54] Alvarez HM, Steinbuchel A. Triacylglycerols in prokaryotic microorganisms. Appl Microbiol Biotechnol 2002;60:367–76.
[55] Kalscheuer R, Stolting T, Steinbuchel A. Microdiesel: Escherichia coli engineered for fuel production. Microbiology 2006;152:2529–36.
[56] Santala S, Efimova E, Kivinen V, Larjo A, Aho T, Karp M, et al. Improved triacylglycerol production in Acinetobacter baylyi ADP1 by metabolic engineering. Microb Cell Fact 2011;10:36.
[57] Kalscheuer R, Stoveken T, Malkus U, Reichelt R, Golyshin PN, Sabirova JS, et al. Analysis of storage lipid accumulation in Alcanivorax borkumensis: evidence for alternative triacylglycerol biosynthesis routes in bacteria. J Bacteriol 2007;189:918–28.
[58] Bacon J, Dover LG, Hatch KA, Zhang Y, Gomes JM, Kendall S, et al. Lipid composition and transcriptional response of Mycobacterium tuberculosis grown under iron-limitation in continuous culture: identification of a novel wax ester. Microbiology 2007;153:1435–44.
[59] Alvarez HM, Souto MF, Viale A, Pucci OH. Biosynthesis of fatty acids and triacylglycerols by 2,6,10,14-tetramethyl pentadecane-grown cells of Nocardia globerula 432. FEMS Microbiol Lett 2001;200:195–200.
[60] Kurosawa K, Boccazzi P, de Almeida NM, Sinskey AJ. High-cell-density batch fermentation of Rhodococcus opacus PD630 using a high glucose concentration for triacylglycerol production. J Biotechnol 2010;147:212–8.
[61] Voss I, Steinbüchel A. High cell density cultivation of Rhodococcus opacus for lipid production at a pilot-plant scale. Appl Microbiol Biotechnol 2001;55:547–55.
[62] Miller N. Process design and modeling for the production of striacylglycerols (TAGs) in Rhodococcus opacus PD630: Massachusetts Institute of Technology (MIT); 2012.
[63] Arabolaza A, Rodriguez E, Altabe S, Alvarez H, Gramajo H. Multiple pathways for triacylglycerol biosynthesis in Streptomyces coelicolor. Appl Environ Microb 2008;74:2573–82.
[64] Yang C, Hua Q, Shimizu K. Energetics and carbon metabolism during growth of microalgal cells under photoautotrophic, mixotrophic and cyclic light-autotrophic//dark-heterotrophic conditions. Biochem Eng J 2000;6:87–102.
[65] Cao Z, Gao H, Liu M, Jiao P. Engineering the acetyl-CoA transportation system of Candida tropicalis enhances the production of dicarboxylic acid. Biotechnol J 2006;1:68–74.
[66] Coleman R, Lee DP. Enzymes of triacylglycerol synthesis and their regulation. Prog Lipid Res 2004;43:134–76.
[67] Athenstaedt K, Daum G. Phosphatidic acid, a key intermediate in lipid metabolism. Eur J Biochem 1999;266:1–16.
[68] Oelkers P, Cromley D, Padamsee M, Billheimer JT, Sturley SL. The DGA1 gene determines a second triglyceride synthetic pathway in yeast. J Biol Chem 2002;277:8877–81.
[69] Sandager L, Gustavsson MH, Stahl U, Dahlqvist A, Wiberg E, Banas A, et al. Storage lipid synthesis is non-essential in yeast. J Biol Chem 2002;277:6478–82.
[70] Racenis PV, Lai JL, Das AK, Mullick PC, Hajra AK, Greenberg ML. The acyl dihydroxyacetone phosphate pathway enzymes for glycerolipid biosynthesis are present in the yeast Saccharomyces cerevisiae. J Bacteriol 1992;174:5702–10.
[71] Minskoff SA, Racenis PV, Granger J, Larkins L, Hajra AK, Greenberg ML. Regulation of phosphatidic acid biosynthetic enzymes in Saccharomyces cerevisiae. J Lipid Res 1994;35:2254–62.
[72] Dahlqvist A, Stahl U, Lenman M, Banas A, Lee M, Sandager L, et al. Phospholipid: diacylglycerol acyltransferase: an enzyme that catalyzes the acyl-CoA-independent formation of triacylglycerol in yeast and plants. Proc Natl Acad Sci USA 2000;97:6487–92.
[73] Kalscheuer R, Steinbuchel A. A novel bifunctional wax ester synthase/acyl- CoA:diacylglycerol acyltransferase mediates wax ester and triacylglycerol biosynthesis in Acinetobacter calcoaceticus ADP1. J Biol Chem 2003;278:8075–82.
[74] Davis MS, Solbiati J, Cronan Jr JE. Overproduction of acetyl-CoA carboxylase activity increases the rate of fatty acid biosynthesis in Escherichia coli. J Biol Chem 2000;275:28593–8.
[75] Meng X, Yang J, Cao Y, Li L, Jiang X, Xu X, et al. Increasing fatty acid production in E. coli by simulating the lipid accumulation of oleaginous microorganisms. J Ind Microbiol Biotechnol 2011;38:919–25.
[76] Ruenwai R, Cheevadhanarak S, Laoteng K. Overexpression of acetyl-CoA carboxylase gene of Mucor rouxii enhanced fatty acid content in Hansenula polymorpha. Mol Biotechnol 2009;42:327–32.
[77] Tamano K, Bruno KS, Karagiosis SA, Culley DE, Deng S, Collett JR. Increased production of fatty acids and triglycerides in Aspergillus oryzae by enhancing expressions of fatty acid synthesis-related genes. Appl Microbiol Biotechnol 2013;97:269–81.
[78] Tai M, Stephanopoulos G. Engineering the push and pull of lipid biosynthesis in oleaginous yeast Yarrowia lipolytica for biofuel production. Metab Eng 2013;15:1–9.
[79] Zhang X, Agrawal A, San KY. Improving fatty acid production in Escherichia coli through the overexpression of malonyl CoA-acyl carrier protein transacylase. Biotechnol Prog 2012;28:60–5.
[80] Verwoert GS, Linden KH, Walsh MC, Nijkamp HJ, Stuitje AR. Modification of Brassica napus seed oil by expression of the Escherichia coli fabH gene, encoding 3-ketoacyl-acyl carrier protein synthase III. Plant Mol Biol 1995;27:875–86.
[81] Dehesh K, Tai H, Edwards P, Byrne J, Jaworski JG. Overexpression of 3- ketoacyl-acyl-carrier protein synthase IIIs in plants reduces the rate of lipid synthesis. Plant Physiol 2001;125:1103–14.
[82] Voelker TA, Davies HM. Alteration of the specificity and regulation of fatty acid synthesis of Escherichia coli by expression of a plant medium-chain acyl- acyl Carrier protein thioesterase. J Bacteriol 1994;176:7320–7.
[83] Zhang X, Li M, Agrawal A, San KY. Efficient free fatty acid production in Escherichia coli using plant acyl-ACP thioesterases. Metab Eng 2011;13:713–22.
[84] Jain RK, Coffey M, Lai K, Kumar A, MacKenzie SL. Enhancement of seed oil content by expression of glycerol-3-phosphate acyltransferase genes. Biochem Soc Trans 2000;28:958–61.
[85] Zou J, Katavic V, Giblin EM, Barton DL, MacKenzie SL, Keller WA, et al. Modification of seed oil content and acyl composition in the brassicaceae by expression of a yeast sn-2 acyltransferase gene. Plant cell 1997;9:909–23.
[86] Beopoulos A, Chardot T, Nicaud JM. Yarrowia lipolytica: a model and a tool to understand the mechanisms implicated in lipid accumulation. Biochimie 2009;91:692–6.
[87] Bouvier-Nave P, Benveniste P, Oelkers P, Sturley SL, Schaller H. Expression in yeast and tobacco of plant cDNAs encoding acyl CoA:diacylglycerol acyltransferase. Eur J Biochem 2000;267:85–96.
[88] Vigeolas H, Waldeck P, Zank T, Geigenberger P. Increasing seed oil content in oil-seed rape (Brassica napus L.) by over-expression of a yeast glycerol-3- phosphate dehydrogenase under the control of a seed-specific promoter. Plant Physiol 2007;5:431–41.
[89] Beopoulos A, Mrozova Z, Thevenieau F, Le Dall MT, Hapala I, Papanikolaou S, et al. Control of lipid accumulation in the yeast Yarrowia lipolytica. Appl Environ Microb 2008;74:7779–89.
[90] Dulermo T, Nicaud JM. Involvement of the G3P shuttle and beta-oxidation pathway in the control of TAG synthesis and lipid accumulation in Yarrowia lipolytica. Metab Eng 2011;13:482–91.
[91] Lin H, Castro NM, Bennett GN, San KY. Acetyl-CoA synthetase overexpression in Escherichia coli demonstrates more efficient acetate assimilation and lower acetate accumulation: a potential tool in metabolic engineering. Appl Microbiol Biotechnol 2006;71:870–4.
[92] Wynn JP, Ratledge C. Malic enzyme is a major source of NADPH for lipid accumulation by Aspergillus nidulans. Microbiology 1997;143:253–7.
[93] Zhang Y, Adams IP, Ratledge C. Malic enzyme: the controlling activity for lipid production? Overexpression of malic enzyme in Mucor circinelloides leads to a 2.5-fold increase in lipid accumulation. Microbiology 2007;153:2013–25.
[94] Chen J, Lang C, Hu Z, Liu Z, Huang R. Antisense PEP gene regulates to ratio of protein and lipid content in Brassica napus seeds. Chin J Agric Biotechnol 1999;7:316.
[95] Hou DJ, Shi DJ, Cai ZF, Song DH, Wang XK. Regulation of lipids synthesis in transgenic Escherichia coli by inserting cyanobacterial sense and antisense pepcA Gene. Chin J Biotechnol 2008;28:52–8.
[96] Li Y, Han D, Hu G, Dauvillee D, Sommerfeld M, Ball S, et al. Chlamydomonas starchless mutant defective in ADP-glucose pyrophosphorylase hyper- accumulates triacylglycerol. Metab Eng 2010;12:387–91.
[97] Li Y, Han D, Hu G, Sommerfeld M, Hu Q. Inhibition of starch synthesis results in overproduction of lipids in Chlamydomonas reinhardtii. Biotechnol Bioeng 2010;107:258–68.
[98] Lu X, Vora H, Khosla C. Overproduction of free fatty acids in E. coli: implications for biodiesel production. Metab Eng 2008;10:333–9.
[99] Lei AP, Chen H, Shen GM, Hu ZL, Chen L, Wang JX. Expression of fatty acid synthesis genes and fatty acid accumulation in Haematococcus pluvialis under different stressors. Biotechnol Biofuels 2012;5:18–28.
[100] Davis MS, Cronan Jr JE. Inhibition of Escherichia coli acetyl coenzyme A carboxylase by acyl-acyl carrier protein. J Bacteriol 2001;183:1499–503.
[101] Heath RJ, Rock CO. Inhibition of b-ketoacyl-acyl carrier protein synthase III (fabH) by acyl-acyl carrier protein in Escherichia coli. J Biol Chem 1996;271:10996–1000.
[102] Jiang P, Cronan Jr JE. Inhibition of fatty acid synthesis in Escherichia coli in the absence of phospholipid synthesis and release of inhibition by thioesterase action. J Bacteriol 1994;176:2814–21.
[103] Scharnewski M, Pongdontri P, Mora G, Hoppert M, Fulda M. Mutants of Saccharomyces cerevisiae deficient in acyl-CoA synthetases secrete fatty acids due to interrupted fatty acid recycling. FEBS J 2008;275:2765–78.
[104] Zheng ZF, Zou JT. The initial step of the glycerolipid pathway: identification of glycerol 3-phosphate/dihydroxyacetone phosphate dual substrate acyltransferases in Saccharomyces cerevisiae. J Biol Chem 2001;276:41710–6.
[105] Zaremberg V, McMaster CR. Differential partitioning of lipids metabolized by separate yeast glycerol-3-phosphate acyltransferases reveals that phospholipase D generation of phosphatidic acid mediates sensitivity to choline-containing lysolipids and drugs. J Biol Chem 2002;277:39035–44.
[106] Nagiec MM, Wells GB, Lester RL, Dickson RC. A suppressor gene that enables Saccharomyces cerevisiae to grow without making phospholipids encodes a protein that resembles an Escherichia coli fatty acyltransferase. J Biol Chem 1993;268:22156–63.
[107] Waltermann M, Stoveken T, Steinbuchel A. Key enzymes for biosynthesis of neutral lipid storage compounds in prokaryotes: properties, function and occurrence of wax ester synthases/acyl-CoA: diacylglycerol acyltransferases. Biochimie 2007;89:230–42.
[108] Cases S, Smith SJ, Zheng YW, Myers HM, Lear SR, Sande E, et al. Identification of a gene encoding an acyl CoA diacylgly-cerol acyltransferase, a key enzyme in triacylglycerol synthesis. Proc Natl Acad Sci USA 1998;95:13018–23.
[109] Lardizabal KD, Metz JG, Sakamoto T, Hutton WC, Pollard MR, Lassner MW. Purification of a jojoba embryo wax synthase, cloning of its cDNA, and production of high levels of wax in seeds of transgenic Arabidopsis. Plant Physiol 2000;122:645–56.
[110] Brown TDK, Jones-Mortimer MC, Kornberg HL. The enzymic interconversion of acetate and acetyl-coenzyme A in Escherichia coli. J Gen Microbiol 1977;102:327–36.
[111] Evans CT, Ratledge C. Possible regulatory roles of ATP:citrate lyase, malic enzyme, and AMP deaminase in lipid accumulation by Rhodosporidium toruloides CBS 14. Can J Microbiol 1985;31:1000–5.
[112] Kendrick A, Ratledge C. Desaturation of polyunsaturated fatty acids in Mucor circinelloides and the involvement of a novel membrane-bound malic enzyme. Eur J Biochem 1992;209:667–73.
[113] Wynn JP, Hamid AA, Ratledge C. The role of malic enzyme in the regulation of lipid accumulation in filamentous fungi. Microbiology 1999;145:1911–7.
[114] Rodriguez-Frometa RA, Gutierrez A, Torres-Martinez S, Garre V. Malic enzyme activity is not the only bottleneck for lipid accumulation in the oleaginous fungus Mucor circinelloides. Appl Microbiol Biotechnol 2013;97:3063–72.
[115] Boulton CA, Ratledge C. Correlation of lipid accumulation in yeasts with possession of ATP:citrate lyase. J Gen Microbiol 1981;127:169–76.
[116] Michinaka Y, Shimauchi T, Aki T, Nakajima T, Kawamoto S, Shigeta S, et al. Extracellular secretion of free fatty acids by disruption of a fatty acyl-CoA synthetase gene in Saccharomyces cerevisiae. J