Progress in Lipid Research 52 (2013) 395–408

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Progress in Lipid Research

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Review

Advancing oleaginous microorganisms to produce lipid via metabolicengineering technology

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 oxidaarachidonic acid; BE, branching enzymes; CDP-DAG, CDP-diacylglycerol; DAG, diacylglycerol; DGAT, diacylglycerol acyl-transferase; DHA, docosahexenoic aciddihydroxyacetone phosphate; DHAPAT, DHAP acyltransferase; EPA, eicosapentaenoic acid; FAS, fatty acid synthetase; FAT, acyl-ACP-thioesterase; FFA, free fatty aglyceraldehyde 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-phacyltransferase; GPD1 and GUT2, glycerol 3-phosphate dehydrogenase; KAS, b-ketoacyl-ACP synthase; LPA, lysophosphatidate; LPAT, lysophosphatidate acyl-traMAT, malonyl-CoA:ACP transacetylase; ME, malic enzyme; PA, phosphatidate; PAP, phosphatidic acid phosphatase; PDAT, phospholipid:diacylglycerol acyltransferapyruvate dehydrogenase; PEP, phosphoenolpyruvate; PEPC, phosphoenolpyruvate carboxylase; Pi, inorganic pyrophosphate; PYC, pyruvate carboxylase; SS, starch sTAG, 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: jgjiang@scut.edu.cn (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, Chinab 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 2013Received in revised form 3 May 2013Accepted 6 May 2013Available online 16 May 2013

Keywords:Oleaginous microorganismsMetabolic engineeringBiodieselMicrobial oilsLipidsMicroalgae

a b s t r a c t

With the depletion of global petroleum and its increasing price, biodiesel has been becoming one of themost 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 toscale up. Many oleaginous microorganisms can accumulate lipids, especially triacylglycerols (TAGs),which are the main materials for biodiesel production. This review is covering the related researcheson different oleaginous microorganisms, such as yeast, mold, bacteria and microalgae, which mightbecome the potential oil feedstocks for biodiesel production in the future, showing that biodiesel fromoleaginous microorganisms has a great prospect in the development of biomass energy. Microbial oilsbiosynthesis process includes fatty acid synthesis approach and TAG synthesis approach. In addition,the strategies to increase lipids accumulation via metabolic engineering technology, involving theenhancement 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 targetsfor gene transformation, and reducing PEPC activity is observed to be beneficial for lipid production.

� 2013 Elsevier Ltd. All rights reserved.

Contents

1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3962. Oleaginous microorganisms for biodiesel production . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 396

2.1. Oleaginous microalgae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3972.2. Oleaginous yeast and mold. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3972.3. Oleaginous bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 398

3. TAG biosynthesis in microorganisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 400

3.1. Fatty acid synthesis approach. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4003.2. 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) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4024.1.2. Fatty acid synthetase (FAS) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4024.1.3. Acyl-ACP-thioesterase (FAT) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 402

se; ARA,; DHAP,

cid; GAP,osphate

nsferase;se; PDH,ynthase;

9.

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). . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4024.2.2. Lysophosphatidate acyl-transferase (LPAT). . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4034.2.3. Acyl-CoA:diacylglycerol acyl-transferase (DGAT) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4034.2.4. Glycerol 3-phosphate dehydrogenase (GPDH) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 403

4.3. Regulation of related TAG biosynthesis bypass approaches . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 403

4.3.1. Acetyl-CoA synthase (ACS). . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4034.3.2. Malic enzyme (ME). . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4034.3.3. ATP:citrate lyase (ACL) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 403

4.4. Blocking competing pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 404

4.4.1. Repression of b-oxidation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4044.4.2. Repression of phospholipid biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4044.4.3. Repression of phosphoenolpyruvate carboxylase (PEPC) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4044.4.4. Repression of starch biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4044.4.5. Repression of the degradation of TAG. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 405

4.5. The multi-gene approach . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 405

5. Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 405

Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 406References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 406

1. Introduction

The earliest research on producing lipids from microorganismscould be traced back to the First World War, when Germany hadprepared with some strains of Endomyces and Fusarium sp. to pro-duce lipids to solve the cooking oil shortage problem [1]. In recentyears, high energy prices, energy and environment security, con-cerns about petroleum supplies are drawn great attention anddrive us to find a renewable biofuel. One of the most promisingrenewable 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 rapidand transitional growth. In the year 2007, there were only 20 nationsproducing biodiesel for the needs of over 200 nations; by the year2010, more than 200 nations became biodiesel producing nationsand suppliers. Global biodiesel production has massively increasedto 18.2 billion liters per year from 2000 to 2010 (Fig. 1). However,the plant oil materials require large energy and acreage for sufficientproduction of oilseed crops. For example, using the average oil yieldper hectare from various crops, the cropping area needed to meet50% 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 viewof 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 oilwas used for biodiesel production, the cost of source would accountfor 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 feedstockslimits the large-scale development of biodiesel to a large extent.Therefore, to lower the cost of oil raw materials, much attentionhas been paid to the development of microbial oils and it has beenfound that many microorganisms, such as microalgae, yeast, bacte-ria, and fungi, have the ability to accumulate oils under some specialcultivation conditions.

Compared to plant oils, microbial oils have many advantages,such as short life cycle, abundant and cheap raw materials, lesslabor required, less affections by venue, season and climate, andeasier to scale up [8,9]. Therefore, microorganisms might becomeone of the potential oil feedstocks for biodiesel production in thefuture, although there are many researches associated with micro-organism-producing oils that need to be further carried out. Thisreview is covering the related researches about different oleagi-nous microorganisms for lipids production and microbial TAG bio-synthesis process. Moreover, metabolic engineering strategies toincrease lipid production are introduced in detail.

2. Oleaginous microorganisms for biodiesel production

Oleaginous microorganisms are defined as oleaginous specieswith 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]. Whilethe 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 oleaginousmicroorganisms can accumulate oils, especially TAGs, which arethe main materials for biodiesel production. TAGs act with alcoholunder acid or alkali catalyst by transesterification to generate fattyacid 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 reducethe cost of microbial oils, exploring other carbon sources instead ofglucose 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 thecarbon sources for microbial oils accumulation. Due to the low

CH

CH2

CH2

OOC-R1

OOC-R2

OOC-R3

+ 3R'OHCatalyst

Triacylglycerol(TAG)

Alcohol

R1-COO-R'

R2-COO-R'

R3-COO-R'

Esters(Biodiesel)

+ CH

CH2

CH2 OH

OH

OH

Glycerol

Fig. 3. Biodiesel producing by alcoholysis.

0

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Oil yield (Liter/hectare)

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Oil

yie

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Lite

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Lan

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ea n

eede

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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 dryweight) 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 asthe carbon sources and sunlight as the energy for photoautotrophicculture, 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 mixotrophicculture. Scaling up for autotrophic microalgae is more complicated,since light is needed during the cultivation process. To minimizethe cost, oil production from microalgae must rely on available freesunlight, despite daily and seasonal variations in light levels [13].Heterotrophic microalgae are easily cultivated and controlled innormal fermenters. But, they require organic carbon sources foroil accumulation, which might limit the application of such micro-algae used for oil production for biodiesel production [11]. As forfungi, scaling up for fungi needs to be carried out further. To utilizecheap carbon sources for oil production from fungi opens a newway for cost reduction, which is very important for such oils usedfor biodiesel production in the future [11].

2.1. Oleaginous microalgae

Generally speaking, microalgae whose lipid contents in thecells are more than 20% of the dry weight are called oleaginousmicroalgae. The average lipid contents of algal cells varybetween 1% and 70% but can reach 90% of dry weight undercertain conditions [14,15]. Oil levels of 20–50% are quitecommon [16] (Table 1). Microalgae, such as Chlorophyta,Bacillariophyceae, have higher oil contents, and they are easierto cultivate, especially chlorella, which could be applied to indus-trial production, so they are the ideal energy microalgaeresources [17,18]. At present to breed optimal chlorella strains

suitable for large-scale breeding for biodiesel production hasaroused great attention from international energy researchinstitutions and biological energy development companies. Miaoet al. [19] acquired high lipid content of heterotrophic chlorellaby heterotrophic transformation cell engineering technology,containing the lipid compounds as high as 55% of cell dryweight, which was 4 times of the autotrophic one (14%). NationRenewable Energy Laboratory (NREL) in the United States devel-oped a ‘‘engineering Cyclotella’’ [20], whose oil content can reachmore than 60% in the laboratory condition, and more than40% in outdoor culture, while microalgae lipid content in thenature is generally 5–20%. It is obvious that using ‘‘engineeringmicroalgae’’ for biodiesel production has important economicand ecological significances.

2.2. Oleaginous yeast and mold

Since the Second World War, many highly oleaginous microor-ganisms, like Lipomyces starkeyi, Rhodotorula glutinis, Aspergillusand Mucor and so on were sequentially found. In order to furtherform productivity providing technical basis, the identification of avariety of oleaginous strains has achieved breakthrough [9]. Manyyeast species, such as Cryptococcus albidus, Cryptococcus albidun,Lipomyces lipofer, Lipomyces starkeyi, Rhodosporidium toruloides,Rhodotorula glutinis and Trichosporon pullulans, were found to beable to accumulate lipids under some cultivation conditions(Table 2), suggesting that different yeast species led to differentlipid accumulation. Many mold species, such as Asoergullusterreus, Claviceps purpurea, Tolyposporium, Mortierella alpina,Mortierella isabellina, can also accumulate lipids. Although sometypes of fungi have the ability to produce lipids (Table 2), mostfungi are explored mainly for the production of special lipids,such as DHA, GLA, EPA and ARA, and there are few reports onthe utilization of fungi oils for biodiesel production [9]. It wasreported that when Mortierella isabellina was cultivated in nitro-gen-limited and high utilization rate of glucose medium, or evenin high initial sugar concentration medium, significant fatty acidquantities were accumulated after nitrogen depletion, resultingin a notable microbial oils production of 18.0 g/L in culturemedium, containing a maximum concentration of 0.801 g GLAper liter of culture medium [36].

Table 1Oil content of some microalgae.

Microalgal species Culture conditions Lipid content(% dry weight)

Biomass productivity(g/L/day)

Lipid productivity(mg/L/day)

Refs.

ChlorophytaChlorella 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]

BacillariophyceaeChaetoceros 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]

OthersCrypthecodinium 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 2Different carbon sources used for yeast and mold lipid production.

Species Carbon sources Lipid content (% dry weight) Refs.

YeastCryptococcus 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]

MoldCunninghamella 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 oilunder some special environment. But usually, the lipid composi-tion produced by bacteria is quite different from other microbialoils. Most bacteria just produce complex lipoid [9], and only a fewbacteria can produce oils that can be used as the feedstock forbiodiesel production [12]. In bacteria, the most abundant classof neutral lipids are polyhydroxyalkanoic acids serving as intra-cellular carbon and energy storage compounds [48], but also

few examples of substantial TAG accumulation have beenreported for species mostly belonging to the actinomycetesgenera Mycobacterium [49], Nocardia, Rhodococcus [50] andStreptomyces [51] (Table 3). Recently, it was found that Gordoniasp. and Rhodococcus opacus could accumulate oils under somespecial conditions with maximum oil content of 80%, but thebiomass is only 1.88 g/L [52].

Compared to other microorganisms, many gene regulationmechanisms in fatty acid synthesis in bacteria are already under-stood [53,54]. Therefore, it is relatively easy to use biological

Table 3TAGs 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 forlipids 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. InAcinetobacter 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:diacylglycerolacyltransferase; PDAT: phospholipid:diacylglycerol acyltransferase; DHAPAT: DHAP acyltransferase; GPD1 and GUT2: encoding glycerol 3-phosphate dehydrogenase; TGL3and TGL4: encoding triacylglycerol lipases; POX1–6: encoding the six acyl-CoA oxidases; MFE1: encoding multi-functional enzyme.

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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. Itwas reported that a metabolically engineered Escherichia coli couldproduce biodiesel directly, and the fatty acid esters concentrationsof 1.28 g/L was achieved by fed-batch fermentation using renew-able carbon sources [55]. Although the yield was low, it provideda new idea for the biodiesel production.

3. TAG biosynthesis in microorganisms

TAGs, as neutral lipids, are the main materials in the productionof biodiesel. The synthesis routes of TAG in microorganisms mayconsist 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 carbonsources (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 lipidsproduction. The metabolism flux route on the utilization of carbondioxide and glucose for the formation of acetyl-CoA in microalgaewas described by Yang and Shimizu [64].

Microalgae can perform oxygenic photosynthesis and fixcarbon dioxide through Calvin cycle like plant cells. That is,microalgal cells can trap light energy as the energy source andassimilate CO2 as the carbon source. CO2 enters in chloroplastvia Calvin cycle to generate glyceraldehyde 3-phosphate (GAP)(Fig. 4). GAP is withdrawn from Calvin cycle and exported tocytoplasm for consumption [64], and then to form pyruvate viaglycolytic pathway. Finally, pyruvate is converted to acetyl-CoAby pyruvate dehydrogenase (PDH). When glucose is the carbonsource of microorganisms, it can be converted to pyruvate inthe cytoplasm after several steps by glycolytic pathway. Afterentering 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 transfermitochondrion as citrate; in the cytoplasm they are delivered as acetyl-CoA for fattyATP:citrate lyase; ME: Malic enzyme.

high, citrate enters into cytoplasm, where it is cleaved to formacetyl-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 carboxylicenzyme (ACC) and fatty acid synthase (FAS) in most microorgan-isms. From Fig. 4, ACC uses acetyl-CoA for malonyl-CoA formationand acyl chain elongation. Once malonyl-CoA is synthesized, it istransferred by malonyl-CoA:ACP transacetylase (MAT), one of thefatty acid synthase (FAS) multi-enzymatic complex subunits, toform malonyl-acyl-carrier protein (malonyl-ACP). The FAS trans-fers the malonyl moiety to acyl-carrier protein (ACP) to use it asa carbon source for the synthesis of long chain fatty acids, mainlyC16 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 catalyzedby 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 fattyacid.

3.2. TAG synthesis approach

For eukaryotes, TAG formation takes place in specializedorganelles, like the mitochondria and plastid located in theendoplasmic reticulum. In contrast, the TAG synthesis takes placein the cytoplasm of prokaryotic cells. The most important route toTAG biosynthesis is the G3P or Kennedy pathway (Fig. 4), firstdescribed by Professor Eugene Kennedy and his colleagues inthe 1950s, by means of which more than 90% of liver TAGs areproduced.

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-phosphateacyl-transferase (GPAT). This enzyme exhibits the lowest specificactivity in the TAG synthesis pathway, and is suggested to bepotentially the rate limiting step [65,66]. The LPA is then further

of acetyl group from mitochondrion to the cytoplasm. Acetyl groups pass out of theacid 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 acidphosphatase (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 acritical metabolic branch point in the Kennedy pathway for glycer-ide and glycerophosphatide synthesis [68]. Overexpression ofDGAT would commit more DAG to TAG formation rather thanphospholipid formation.

It is reported that de novo formation of PA in yeast can occureither through the glycerol-3-phosphate (G3P) or dihydroxyace-tone phosphate (DHAP) pathways. Whereas the former route ofPA 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 reducedby 1-acyl-DHAP reductase to yield LPA, which is further acylatedto PA by LPAT.

Table 4Researches about lipid synthesis by overexpressing genes or knockout genes.

Genes (enzymes) Source-species Rec

accA-D (ACC), tesA (thioesterase I) E. coli (bacteria) E. cACC Acinetobacter calcoaceticus (plant) E. cACC1(ACC) Mucor rouxii (mold) HaACC Aspergillus oryzan (mold) AspACC1(ACC) Yarrowia lipolytica (yeast) YarFAT, fabD (MAT) E. coli, Streptomyces avermitilis

MA-4680, Streptomyces coelicolor A3(2)(bacteria)

E. c

KAS III E. coli (bacteria) BraKAS III Spinacia oleracea (plant) Nic

FAT Umbellularia californica (plant) E. cFAT Diploknema Butyracea (plant) E. cFAT Ricinus communis (plant) E. cFAT Jatropha curcas (plant) E. cGPAT Safflower (plant), E. coli (bacteria) AraSLC1-1 (LPAT) Yeast RapD LRO1 (PDAT) Y. lipolytica (yeast) Y. lDGAT Arabidopsis (plant) Yea

GPD1 (GPDH) Yeast Bra

DGUT2 (GPDH) Y. lipolytica (yeast) Y. lGPD1 (GPDH) Y. lipolytica (yeast) Y. lDGUT2 (GPDH) Y. lipolytica (yeast) Y. lGPD1, DGUT2 (GPDH) Y. lipolytica (yeast) Y. lACS E. coli (bacteria) E. cDME Aspergillus nidulans (mold) AspME E. coli K-1 (bacteria) E. cmalA (ME) Mortierella alpine, Mucor circinelloides

(mold)Mu

ACL Aspergillus oryzan (mold) Asp

Antisense PEPC Agrobacterium tumefaciens (bacteria) BraAntisense PEPC Anabaena sp. (microalgae) E. cDAGPase Chlamydomonas (microalgae) ChlDAGPase Chlamydomonas reinhardtii BAFJ5

(microalgae)Chl(m

DTGL3, DTGL4 (TAG lipases) Y. lipolytica (yeast) Y. l

Multi-gene approach to enhance lipid biosynthesisACC, thioesterase, D fadD (acyl-CoA

synthetase)E. coli (bacteria), plant E. c

ACP, KAS, FAT Haematococcus pluvialis (microalgae) HaACC1, DGAT1 Y. lipolytica (yeast) Y. lPOX1-6 (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 havean acyl-CoA-independent mechanism for TAG synthesis, whichuses 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 thatAcinetobacter calcoaceticus ADP1 accumulates both wax ester(WE) and TAG, and a bifunctional wax ester synthase/acyl-CoA:diacylglycerol acyltransferase (WS/DGAT) was identified toexhibit both wax ester synthase and DGAT activities [73].

4. Metabolic engineering for TAG production

Numerous studies have been carried out using the metabolicengineering strategy to enhance the lipid accumulation in differentspecies. Some of these studies have been summarized in Table 4and will be discussed briefly in this section. They can be broadlyclassified into five different approaches: (1) overexpressing en-zymes of the fatty acid biosynthesis pathway; (2) overexpressingenzymes that enhance the TAG biosynthesis pathway; (3) regula-tion of related TAG biosynthesis bypass approaches; (4) partiallyblocking competing pathways; and (5) the multi-gene transgenicapproach.

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% lipidcontent

[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 BAFJ5icroalgae)

+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) andthioesterase 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 fattyacid, 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 anobvious increase in fatty acid production, which indicated thatsteps later in the pathway limit the flux through the pathway. Thiscould be because the overexpression of native E. coli ACC sufferedfrom feedback inhibition by acyl-ACP [100]. tesA may reduce thisinhibition by forming free fatty acid, resulting in the increased pro-duction of fatty acid. Another way to reduce this inhibition couldbe to express a non-native ACC gene that is not recognized bythe acyl-ACP of E. coli. For example, the heterologous expressionof ACC from Acinetobacter calcoaceticus in E. coli caused a 3-foldenhancement in lipid levels and thus provided support for theassumption [75].

In eukaryotic microorganisms, overexpression of ACC has beenmet with only limited improvement of lipid production. TheACC1 enzyme is responsible for providing the malonyl-CoA usedin cytoplasmic fatty acid synthesis. Heterologous expression ofACC1 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 TAGsrelative to the parental strain [77]. It was suspected that improve-ments in total lipid accumulation have been limited in eukaryotesby the strong metabolic and regulatory control maintained overthis 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 contentthrough 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 throughthe enzyme malonyl CoA:ACP transacylase (MAT). The threeE. coli strains carrying acyl-ACP thioesterase (FAT) and a fabD geneencoding MAT from E. coli, Streptomyces avermitilis MA-4680, orStreptomyces coelicolor A3(2) improved the free fatty acid produc-tion, about 11% more than the control strains, suggesting that fabDoverexpression could be used to improve free fatty acid productionby increasing the malonyl-ACP availability [79].

b-ketoacyl-ACP synthase III (KAS III) condenses acetyl-CoA withmalonyl-ACP to initiate fatty acid biosynthesis. It was defined asthe kinetic mechanism that underlay the negative regulation ofKAS III activity long-chain acyl-ACP [101]. An E. coli KAS III wasoverexpressed 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 andincreased the amounts of C18:2 and C18:3 as compared to controlplants. Such changes in fatty acid composition reflected changes inthe regulation and control of fatty acid biosynthesis, proposing thatfatty 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, aspinach (Spinacia oleracea) KAS III was expressed in tobacco (Nico-tiana tabacum) and resulted in a 300-fold increase in activity abovethe 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 inACP levels in tobacco leaves, although other FAS activities wereunaffected. Decreases in fatty acid content as a result of KAS IIIoverexpression were attributed to the decreased rates of de novofatty acid synthesis most likely by the reduced malonyl-CoA poolsfor subsequent KAS condensation reactions.

In microalgae, main key genes like ACP, KAS and FAT genesoverexpression 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 formetabolic engineering for fatty acid metabolism enhancement,probably due to the fact that FAS is a multi-enzymatic complexcontaining subunits whose activities depend on one another. Thedifficulties 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 acidscan be produced by introducing an FAT gene. The presence of theFAT will break the fatty acid elongation cycle and release free fattyacids [98,102].

It was reported that the expression of the plant (Umbellulariacalifornica) enzyme medium-chain FAT in E. coli deficient in fattyacid 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 actas feedback inhibitors for fatty acid synthsis [82]. It was examinedthe effect of different FAT on the quantities and compositions offree fatty acid produced by an E. coli strain ML103 carrying FATgene from four different plants. The strain carrying the FAT genesfrom 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, morethan 2.0 g/L at 48 h. These two strains accumulated three majorstraight chain free fatty acids, C14, C16:1 and C16 at levels about40%, 35% and 20%, respectively [83]. It was shown that the amountof 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 acylationof DHAP by acyl-CoA:DHAP acyltransferase (DHAPAT) followed byreduction of the newly formed 1-acyl-DHAP to LPA by DHAP reduc-tase (Fig. 4). S. cerevisiae GPAT mutants are equally defective inDHAPAT activity, thereby establishing the genetic identity of thetwo activities in yeast [103]. In S. cerevisiae, GAT1 and GAT2(SCT1) genes encode the major GPAT activities. In vitro substratespecificities were determined and the GAT1 gene product coulduse both G3P and DHAP with similar efficiencies and had a broadfatty acyl-CoA specificity profile, whereas the GAT2 gene productpreferred G3P over DHAP and had a 2.5- to 5-fold preference forC16 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 TAGbiosynthesis [105].

It was reported that the oil content of Arabidopsis seeds was in-creased from 8% to 29% in selected transgenic lines by expressing aplastidial safflower GPAT and an E. coli GPAT gene, demonstratingthat expression of both a bacterial and a plant GPAT gene increasedplant 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 variantLPAT, which can be used to change total fatty acid content andcomposition as well as to alter the stereospecific acyl distributionof fatty acids in seed TAGs [106], was used to transform rapeseedand Arabidopsis [85], leading to increases from 8% to 48% seed oilcontent on the seed dry weight basis, suggesting that increasingthe expression of glycerolipid acyltransferases in seeds leads to agreater 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 DGAT1or DGAT2 families present in yeast, plants, and animals, and thephospholipid:diacylglycerol acyltransferase (PDAT) catalyzes TAGformation in yeast and plants. Yeast LRO1 is incapable of synthesiz-ing sterols and encodes the PDAT. Deletion of LRO1 encoding PDATin Y. lipolytica resulted in a 40% loss of TAG [86]. Another experi-ment with a knockout mutant demonstrated the key role of thebifunctional WS/DGAT for biosynthesis of both storage lipids (TAGsand WEs) in A. calcoaceticus ADP1 [107]. The WS/DGAT enzymes inprokaryotes are not related to any known acyltransferase involvedin the formation of TAGs and WEs in eukaryotes, including theDGAT1 [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-foldincrease of TAGs accumulation [87].

The success with DGAT could be explained by the fact that thesubstrate of DGAT, DAG, could be allocated to either phospholipidbiosynthesis or TAG formation. Overexpression of DGAT wouldcommit 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 cellsof 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 byDGAT 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 intransgenic oil-seed rape (B. napus L.) under the control of theseed-specific promoter. It was found that a 2-fold increase in GPDHactivity led to a 3–4-fold increase in the level of G3P in developingseeds, resulting in a 40% increase in the final lipid content of theseed [88]. The GUT2 gene coding for another GPDH isomer, whichcatalyzes the dihydroxyacetone phosphate (DHAP) formation fromG3P 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 togetherresult in 1.5-, 2.9- and 5.6-fold respective increases in the levelof 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 lipidaccumulation 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 biosynthesisor oxidation via the TCA cycle [110]. The overexpression of ACS inE. coli showed significant reduction in acetate during glucosemetabolism. It also greatly enhanced the assimilation of acetatewhen used as the sole carbon source. Increased ACS levels presum-ably enhanced the activation of acetate to acetyl-CoA, which mayincrease the rate of fatty acid synthesis. For instance, it was ob-served that, by overexpressing the ACS gene in E. coli, the ACSactivity was increased by 9-fold, leading to a significant increaseof 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 isneeded for fatty acid biosynthesis. It has been suggested that thefunction of ME in lipid biosynthesis is to supply NADPH for fattyacid synthase and desaturases [111,112]. A mutant (scuK248) lack-ing ME activity accumulated only half the lipid (l2% of cell dryweight) accumulated by strains of Aspergillus nidulans possessingME [92].

The effect of ME was studied in filamentous fungi in correlationwith lipid accumulation [113]. It was observed that the enhancedenergy (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 enhancedactivity of ME led to the increase of the cytosolic NADPH pool,making available more reducing power for lipogenic enzymes suchas ACC, FAS and ATP:citrate lyase (ACL). It was reported that pro-viding a high level of NADPH by overexpressing ME in E. coli andadding malate to the culture medium resulted in a 4-fold increasein intracellular lipids [75].

The contribution of malA encoded for the isoforms III and IV ofME to lipid accumulation has been further supported because over-expression of the gene coding for them led to a 2.5-fold increase oflipid 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 lowME 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 forthe generation of the acetyl-CoA, which could be incorporated intofatty 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 mammalsand oleaginous yeasts and fungi. The specific activity of ACL en-zyme correlates with the specific rate of lipid synthesis, and it

OO

OH

CH2P

O

HOOH

OHO

OPhosphoenolpyruvate Pyruvate

PEPCActivate

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 Aspergillusoryzae, where a 1.7-fold increase in the productivities of fatty acidsand 1.9-fold increase of TAG relative to the parental strain were ob-served [77].

SCoA

O

O

OHO

O

HO

O FFA

Oxaloacetate

Acetyl-CoA

ACC

4.4. Blocking competing pathways

From the metabolic engineering point of view, blocking offcompeting pathways may also enhance the metabolic flux beingchanneled 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-oxidationA complementary strategy to increase lipid accumulation is to

decrease lipid catabolism. Genes involved in the activation of bothTAGs and free fatty acids synthesis, as well as genes directly in-volved in b-oxidation of fatty acids to be inactivated, sometimesresult in the increase of cellular lipid content.

Several reports have shown that knocking out genes involved inb-oxidation in S. cerevisiae not only can lead to increased amountsof intracellular free fatty acids but also results in extracellular fattyacid secretion in some instances [103,116]. Dysfunction of AOX inS. cerevisiae leading to the inactivation of the b-oxidation processwill result in the accumulation of cellular fatty acids. S. cerevisiae(nonoleaginous yeast) produces a relatively low level of lipids. Itwas explained that nonoleaginous yeasts may not be able to accu-mulate the substrate due to a feedback regulation of the oxidationprocess which would affect hydrophobic substrates (e.g., alkanes,fatty acids, and oils) transporters and down-regulate the diffusionprocess. In addition, lower levels of ATP deriving from b-oxidationwere produced in these strains, and may be used in pathways otherthan lipid storage [86].

While Y. lipolytica (oleaginous yeast) is able to accumulate lipidsto levels exceeding 50% of cell dry weight [117]. It contains sixAOXs, 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 andnumber of lipid bodies in which these fatty acids accumulate[118], suggesting the existence of a regulatory mechanism instrains 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 lipiddegradation and therefore leads indirectly to an increase in lipidaccumulation [89].

As for many microalgae, during diel light–dark cycles, initiateTAG storage during the day and deplete those stores at night tosupport cellular ATP demands and cell division. Consequently,inhibition of b-oxidation would prevent the loss of TAG duringthe 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 lipidproduction in microalgae grown in photobioreactors with exoge-nous carbon sources and continuous light.

4.4.2. Repression of phospholipid biosynthesisPhospholipid biosynthesis is another competitive pathway to

TAG formation because it competes against TAG biosynthesis fora common substrate, PA. If PA is converted into CDP-diacylglycerolinstead of DAG (Fig. 4), it enters the phospholipids synthetic path-way [66]. It was shown that inhibition of phospholipid synthesiscaused 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 topyruvate, 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, thebiosyntheses of either protein or lipid take PEP as the commonsubstrate. 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 ofPEP increases to high level to help to the formation of more pyru-vate. Meanwhile, along with consumption of acetyl-CoA for theformation of malonyl-CoA, the low level of acetyl-CoA is able toactivate the biochemical process of converting PEP to pyruvate(Fig. 6).

Many evident roles of PEPC in lipids biosynthetic pathways inoilseed crops have been reported. By expressing antisense PEPCin 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 obtainedwith transgenic soybean lines harbouring anti-PEPC gene[121,122]. Therefore, the antisense expression of PEPC gene forthe modulation of fatty acids biosynthesis in oil crops is an effec-tive method.

In microalgae, preliminary results also indicated that PEPC playsa role in the regulation of fatty acid accumulation and reducedPEPC activity by antisense expression was correlated with a46.9% increase of the lipid content in Anabaena sp., a cyanobacte-rium [95], showing a great progress in microalgae engineeringfor biodiesel production.

4.4.4. Repression of starch biosynthesisIn many microalgal cells, starch is another major carbon and en-

ergy storage compound, particularly under stress conditions. Thefirst 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) andbranching enzymes (BE) to elongate the glucan chains of the starchgranule [123].

OH

OH

H

OHH

OHH

OH

CH2

HO C O

O

OH

OP

O

HO OH

OH

OH

H

OH

OHH

OH

CH2

H

HO

P

N

NN

N

NH2

O

HOH

HH

HH

OPO

O-

O

P

O-

OOH

OH

H

OH

OHH

OH

CH2

H

HO

OP

O

HO OHO

HO OH

Starch

BESS

AGPaseATP

Pi

GAP

G-6-P G-1-P

ADP-Glucose

CO2

Fig. 7. Starch biosynthesis pathway in chloroplast. GAP: glyceraldehyde-3-phosphate; G-6-P: glucose 6-phosphate; G-1-P: glucose 1-phosphate; Pi: inorganicpyrophosphate; 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 biodieselproduction. Blocking starch synthesis has been suggested as away to boost oil accumulation. Recently, it was reported that inac-tivation of AGPase in a Chlamydomonas starchless mutant led to a10-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 lipidsynthesis 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% ofdry weight or 8- and 3.5-fold higher, respectively, than the wild-type [97]. These results confirmed the feasibility of increasing lipidproduction through redirecting photosynthetically assimilated car-bon away from starch synthesis to neutral lipid synthesis. How-ever, some growth impairment was observed in the low starchand starchless mutants. Starch may play an important role inmaintaining high photosynthetic efficiency. Shifting carbon fluxfrom starch to the lipid synthesis pathway may have a negative ef-fect on photosynthesis in algae, resulting in impairment in growthin starchless and other low starch mutants.

4.4.5. Repression of the degradation of TAGIt 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 onekey enzyme in the TAG pathway to enhance lipid biosynthesis,was suggested by a few researchers [80,124]. Until now, thereare 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, anendogenous thioesterase, and a plant thioesterase, as well asknocked 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 relationshipswith synthesis of fatty acids in H. pluvialis. These genes could bepotential candidates for better quality and higher production offatty acids for biofuel using metabolic engineering techniques [99].

Co-overexpression of two important genes in the lipid synthesispathway, 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 to62% 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 balancebetween the fatty acid and TAG synthesis pathways. Acyl-CoAintermediates function as both product and feedback inhibitorsin the fatty acid (upstream) pathway and primary precursors inthe TAG (downstream) pathway. Upregulation of the upstreampathway increases the throughput of fatty acid synthesis. Upregu-lation of the downstream pathway creates a driving force bydepleting acyl-CoA intermediates and increasing the rate of storageof TAG in lipid bodies. Indeed, coupling precursor overproductionand driving forces with a metabolic sink to enable a push and pulldynamic has become a very powerful strategy in recent efforts ofmetabolic 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, involvedin the second step of the b-oxidation pathway. It was reported thatthe inactivation of POX1-6 or MFE1 strains in which GPD1 was over-expressed or GUT2 was inactivated, showed that G3P accumulationaffects lipid accumulation mostly in strains that are defective for b-oxidation [90]. The finding indicated that TAG synthesis is limitedby the availability of G3P and fatty acids, and that the expression ofgenes involved in TAG homeostasis is regulated by the G3P shuttleand the b-oxidation pathway.

From the above mentioned, multi-gene approach is shown agreat prospect in lipid accumulation for biodiesel production.

5. Conclusion

Taken together, increasing the quantity of any of the Kennedypathway 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 sinksize. 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 byincreasing 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 accumulationachieved by overexpression of Kennedy pathway acyltransferasesmay 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 inrapeseed [128], which suggested that a greater level of controlwas exerted at the level of TAG assembly than at fatty acid synthe-sis [129].

As there are quite a few success stories in lipid overproductionusing transformed microbial strains, the knowledge obtained instudies on lipid pathways and genetic transformed organisms forenhanced lipid synthesis among various species suggests thatDGAT and ME are the most promising targets for gene transforma-tion. Of particular interest, reducing PEPC activity by expressingantisense gene was observed to be beneficial for lipid productionin microalgae. As for multi-gene approach, more trials of this strat-egy should be done due to its great effectiveness. Since oil crisis inthe mid 1970s, finding new energy resources to replace petroleumhas been a hot topic worldwide. Because of the many advantagesover the conventional energy resources, the production of biodieselhas attracted much attention in recent years. We believe that in thenear future, research into lipid metabolism of microorganisms willadvance rapidly and lipid content will increase rapidly by usingmetabolic engineering technology.

Acknowledgements

This project was supported by National Natural Foundation ofChina (Grant 31171631).

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