Lipids of oleaginous yeasts. Part I: Biochemistry of single cell oil production

Review Article

Lipids of oleaginous yeasts. Part I: Biochemistry of singlecell oil production

Seraphim Papanikolaou1 and George Aggelis2

1 Laboratory of Food Microbiology and Biotechnology, Department of Food Science and Technology,

Agricultural University of Athens, Athens, Greece2 Unit of Microbiology, Department of Biology, Division of Genetics, Cell and Development Biology, University

of Patras, Patras, Greece

In the first part of this review, the biochemistry of lipid accumulation in the oleaginous microorganisms is

depicted. Lipid biosynthesis form sugars and related substrates is a secondary anabolic activity,

conducted after essential nutrient (usually nitrogen) depletion in the medium. Due to this

exhaustion, the carbon flow is directed towards the accumulation of intracellular citric acid that is

used as acetyl-CoA donor in the cytoplasm. Acetyl-CoA generates cellular fatty acids and subsequently

triacylglycerols. Lipid accumulation from hydrophobic substrates is a growth associated process, being

independent from nitrogen exhaustion in the medium.Medium fatty acids are incorporated with various

incorporation rates and are either dissimilated for growth needs or become ‘‘substrate’’ for intracellular

biotransformations. ‘‘New’’ fatty acid profiles (in both extra- and intracellular lipids) that did not

previously exist in the medium are likely to be produced. Oleaginous microorganisms consume their

own storage lipids when their metabolic abilities cannot be saturated by the extracellular carbon source.

Reserve lipid breakdown is independent from the type of the carbon source used for lipid accumulation.

In most cases it is accompanied by lipid-free biomass production. Lipid mobilization is a specific process,

as preferential degradation of the neutral lipid fractions is observed.

Keywords: Biodiesel / Lipid accumulation – degradation / Lipid biotechnology / Oleaginous microorganisms / Single

cell oil

Received: January 10, 2011 / Revised: March 19, 2011 / Accepted: April 12, 2011

DOI: 10.1002/ejlt.201100014

1 Introduction

The utilization of oleaginous microorganisms (these that can

accumulate lipids to more than 20% w/w, in their dry cell

mass) as sources of oils and fats (the so-called single cell oils,

SCOs) in large-scale operations presents a great industrial

interest. Although the production price of SCOs is higher

compared with the traditional utilization of common oils and

fats due to the obligatory maintenance of aseptic conditions

[1], several alternative options for large-scale production of

SCOs exist, since the price of various naturally occurring

lipids and fats of the plant and animal kingdom can tremen-

dously vary (from 0.3 to up to 100US $ per kg—see: Ratledge

and Wynn [2]). Therefore identification of microorganisms

capable of producing in increased quantities lipids with struc-

ture and composition similar to that of high-value fats, and

subsequent large-scale production can present an enormous

financial interest [2]. Furthermore, various oleaginousmicro-

organisms have the potentiality to present remarkable growth

and production of SCO on wastes and by-products of the

argo-industrial sector [3–8]; thus, valorization of these resi-

dues together with production of potentially high-added

value lipid could increase the viability of the process being

simultaneously beneficial for the environment.Moreover, the

continuously increasing demand and utilization of the so-

called ‘‘1st generation biodiesel’’ (fatty acid methyl-esters—

FAMEs deriving from trans-esterification of principally plant

oils) that has occurred the last 15 years has increased the price

Correspondence: Dr. Seraphim Papanikolaou, Assistant Professor in

Food Bioprocesses, Laboratory of Food Microbiology and Biotechnology,

Department of Food Science and Technology, Agricultural University of

Athens, 75 Iera Odos, 11855 Athens, Greece

E-mail: [email protected]

Fax: þ30-210-5294700

Abbreviations: Aox, acyl-CoA oxidases; DOT, dissolved oxygen tension

(in %, v/v); Fru, fructose (in g/L); Glc, glucose (in g/L); Glol, glycerol (in

g/L); L, total intra-cellular lipid (in g/L); PUFAs, polyunsaturated fatty acids;

S, substrate fat (in g/L); SCO, single cell oil; TAGs, triacylglycerols; X, total

biomass (in g/L); Xf, lipid-free biomass (in g/L)

Eur. J. Lipid Sci. Technol. 2011, 113, 1031–1051 1031

� 2011 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim www.ejlst.com

of various foodstuffs, and this situation has led to the neces-

sity of discovery of non-conventional sources of oils, that

could be subsequently converted into biodiesel. The oleagi-

nous microorganisms (yeasts, molds, and algae) are con-

sidered as potential candidates for the production of this

lipid that would result in the generation of the so-called

‘‘2nd generation’’ biodiesel [7, 9–16]. Finally, specifically

the oleaginous yeasts, with their unicellular form, are con-

sidered as the more appropriate organisms that have been

used as tools in the comprehension of numerous phenomena

of lipid biochemistry [2, 4, 17–20].

A very large variety of substrates has been used as carbon

sources for the oleaginous microorganisms such as analytical-

grade or industrially-derived (therefore, of low-cost) sugars

or sugar-enriched wastes or residues, polysaccharides,

N-acetylglucosamine, hydrolysates of various products or

by-products, vegetable oils, pure free fatty acids, FAMEs,

fatty byproducts or wastes, n-alkanes, ethanol, glycerol, and

organic acids [4, 7, 8, 10, 13–18, 21–30].

The microbial lipids are mainly composed of triacylgly-

cerols (TAGs) [1, 2, 27, 31, 32]. Other components pre-

sented in non-negligible quantities are free fatty acids

[25, 26, 31–34], other neutral lipids (such as monoacylgly-

cerols, diacylglycerols, and steryl-esters), sterols and polar

fractions (e.g. phospholipids, sphingolipids, glycolipids)

[5, 27, 31, 32, 35]. As a general remark it should be stressed

that when growth is carried out on various hydrophobic

substances utilized as substrates (the so-called ‘‘ex novo’’

lipid accumulation), the microbial lipid produced contains

lower quantities of TAGs compared with growth elaborated

on sugar-based substrates (the so-called ‘‘de novo’’ lipid

accumulation) [5, 25, 31, 32, 36–40]. Additionally, the oil

accumulated by the oleaginous fungi is more unsaturated

than that of the yeasts [2, 18–20, 28–30, 41]. This is the

main reason for which the oleaginous molds have been prin-

cipally used in order to produce lipids rich in polyunsaturated

fatty acids (PUFAs) of medical and dietetical interest like

g-linolenic acid (D6,9,12C18:3) [5, 6, 28–32, 42–49], dihomo-

g-linolenic acid, arachidonic acid, docosahexaenoic acid,

eicosapentanoic acid, etc (for reviews see: Ratledge [4];

Certik and Shimizu [50]; Ratledge and Wynn [2];

Sakuradani and Shimizu [51]; Sakuradani et al. [52]). In

contrast, few oleaginous yeast strains have been revealed

capable to synthesize such types of ‘‘rare’’ PUFAs

[2, 18, 20, 30]. Nevertheless, the last years, the utilization

of genetic engineering has resulted in the construction of

genetically modified yeast strains (principally belonging to

the species Yarrowia lipolytica), that are capable of (super)-

expressing various desaturases (e.g. D5-, D6-, D8-desaturate),

elongases and acyl-transferases, therefore, being, capable to

synthesize the above mentioned ‘‘rare’’ PUFAs and to trans-

fer these newly synthesized molecules into TAGs [53–56].

In the present review article, we will be interested in the

biochemical events related with the lipid accumulation in the

oleaginous yeasts when various carbon sources (hydrophilic

or hydrophobic ones) are utilized as substrates. Also the

events related with storage lipid degradation (turnover) in

the oleaginous microorganisms will be assessed and compre-

hensively discussed.

2 The biochemistry of lipid accumulation anddegradation in the oleaginousmicroorganisms

2.1 Lipid production from fermentation of sugars andrelated substrates used as carbon source

2.1.1 Substrates used

De novo accumulation of cellular lipids is an anabolic bio-

chemical process in which, by virtue of quasi-inverted b-

oxidation reaction series, acetyl-CoA issued by the intermedi-

ate cellular metabolism, generates cellular fatty acids. Fatty

acids are then esterified with glycerol generating structural

(phospholipids, sphingolipids, etc) and reserve (mainly

TAGs) lipids [2, 3, 7, 18–20, 23].

With the notable exception of cellulose and methanol a

very high number of carbon sources have been considered as

substrates for the de novo lipid biosynthesis from oleaginous

microorganisms. Concerning the conversion of cellulose into

SCO, few papers have recently appeared in the international

literature [57, 58]. Sugar-based media such as simple sugars

(e.g. glucose and fructose), lactose, sucrose, whey, glucose-

enriched wastes, molasses, etc [5, 6, 10, 16, 31, 32, 46, 47,

59–68] have been used. Xylose-based media have been

recently considered as substrates of noticeable importance

due to the abundance of xylose as substrate, deriving after

chemical hydrolysis of various lignocellulosic materials [11,

28, 69, 70]. In addition, the utilization of sugar-based sub-

strates more complicated compared with glucose, such as

polysaccharides (e.g. starch and pectin), has been studied.

Although the above substrates are similarly metabolized, in

some cases the results that have been achieved, in terms of

both lipid and fatty acid composition of the SCO produced,

presented notable differences [45, 48, 71].

The stoichiometry of glucose (and similar sugars such

as lactose, fructose, etc) metabolism indicates that about

1.1 moles of acetyl-CoA are generated from 100 g of glucose

(�0.56 moles) catabolized [18, 28, 30]. As far as xylose is

concerned, this compound can be either metabolized through

the phosphoketolase reaction, which is the most efficient

pathway yielding around 1.2 moles of acetyl-CoA per

100 g of xylose (�0.66 moles) utilized, or the pentose phos-

phate pathway, where around 1.0 mole of acetyl-CoA is

formed per 100 g of xylose utilized [18, 28]. Therefore, if

all the acetyl-CoA produced is channeled towards lipid syn-

thesis, the maximum theoretical yield of SCO produced per

glucose consumed is around 0.32 g/g [18]. This value is

somewhat higher concerning the fermentation of xylose

(around 0.34 g/g), assuming that oleaginous microorganisms

1032 S. Papanikolaou and G. Aggelis Eur. J. Lipid Sci. Technol. 2011, 113, 1031–1051


utilize exclusively the phosphoketolase pathway for xylose

assimilation. With reference to glycerol, the maximum theor-

etical yield of SCO is around 0.30 g/g [18]. However, even

under ideal conditions for SCO production (e.g. highly aer-

ated chemostat cultures) lipid yield on glucose consumed can

rarely be higher than 0.22 g/g [1, 2, 7], whereas in other

reports this threshold is considered to be in the level of

0.20 g/g or even lower [28, 46, 47]. However, in

Thamnidium elegans grown on sucrose in shake flasks, the

conversion yield of lipid produced per sugar consumed was

�0.24 g/g, while utilization of other sugars (glucose or fruc-

tose) equally resulted in exceptional conversion yields, i.e.

>0.20 g/g [65]. Maximum conversion yields of the same

magnitude compared with growth of T. elegans on sucrose

(�0.23 g/g) have been reported forCunninghamella echinulata

cultivated on xylose in shake-flask experiments [28]. The

conversion threshold of glycerol into SCO is around

0.10 � 0.02 g/g, in general lower than that obtained on glu-

cose, xylose or other sugars, regardless of the oleaginous

strain used [28, 29, 49, 72–78], presumably due to the

relatively poor regulation of the enzymes involved in the

primary metabolic steps of glycerol assimilation (e.g. glycerol

kinase, 3-P-glycerol dehydrogenase) [7, 76]. In contrast, the

respective value is slightly higher (e.g.�0.15 g/g) for the case

of algae like Schizochytrium limacinum growing heterotroph-

ically on glycerol [79–81], while only in two cases so far

conversion of glycerol into SCO has been performed with

a conversion yield higher than 0.20 g/g; it was either the case

of Aspergillus sp. strains that stored remarkable amounts of

intra-cellular lipids and also produced quantities of extra-

cellular oxalic acid cultured in shake flasks [82], or the case of

the fungus T. elegans that produced indeed high SCO

quantities (from 5.9 to 11.6 g/L corresponding to lipid in

dry weight ranging between 60 and 70% w/w), with conver-

sion yield ranging between 0.16 and 0.22 g/g, equally in

shake-flask experiments [83].

Ethanol has been considered as a potential substrate for

the de novo lipid biosynthesis of the oleaginous microorgan-

isms by a number of investigators [18, 59, 84–86], given that

it is considered as a very proper one since no residual carbon

arises from its uses in fermentation processes [18]. Taking

into consideration that ethanol is by far the more reduced

substrate by any other considered yet for the process of de

novo lipid accumulation, the final stoichiometric balance for

SCO synthesis from ethanol could result in a theoretical yield

of 0.54 g of lipid per 1 g of ethanol consumed [18].

Nevertheless such high conversion yields have never been

achieved in the literature with the conversion threshold of

ethanol into SCO being around 0.31 g/g [18, 29, 84].

Coming back to glycerol, even though this substrate

presents a slightly lower theoretical conversion yield com-

pared with glucose, this carbon source is of substantial and

increasing importance due to its appearance into the market

volume in continuously growing quantities due to application

into an industrial scale of the bio-diesel production process.

In general, the utilization of glycerol as a microbial substrate,

refers mainly to the production of 1,3-propanediol by bac-

teria (for review see: Papanikolaou [87]), whilst to the best of

our knowledge, only some investigations have been con-

ducted so far with (bio-diesel derived waste) glycerol utilized

as substrate (or co-substrate) by microorganisms in order to

produce SCO [26, 28, 29, 49, 72–83]. In the above-cited

papers trials were performed with Cryptococcus curvatus, S.

limacinum,Y. lipolytica,Mortierella isabellina,C. echinulata and

T. elegans strains. Moreover, in a recent investigation,

Chatzifragkou et al. [83] have performed a relatively exten-

sive screening of yeast and Zygomycetes strains (species

Candida boidinii, C. curvata, C. oleophila, C. pulcherrima, C.

echinulata, M. isabellina, M. ramanniana, Mucor sp., Pichia

membranifaciens, Rhodotorula sp., T. elegans, Y. lipolytica,

Zygosaccharomyces rouxii, Zygorhynchus moelleri) cultivated

on bio-diesel derived glycerol in conditions favoring the pro-

duction of SCO (nitrogen-limited conditions) and they have

found that although yeast strains in general produced higher

quantities of biomass compared with the molds, the pro-

duction of lipid was increased in the fungal strains. The more

promising SCO producer amongst all strains tested was the

Zygomycete T. elegans [83].

Citric acid [88], acetic acid [89–91], or other low-mol-

ecular weight organic acids [90, 92, 93] have been equally

considered as substrates for SCO production. Specially, as far

as acetic acid is concerned, it is considered as a remarkable

pollutant, generated either in the process water of Uranium

bleaching or as effluent issued from the Fischer–Tropsch

reaction [90, 93]; the investigations, thus, concerning its

biotransformation in SCO are very interesting in both econ-

omical and ecological terms.

2.1.2 Biochemistry of de novo lipid accumulation

All microorganisms are capable to synthesize lipids, though

only the oleaginous strains may accumulate inside their cells

significant lipid quantities (i.e. >20% w/w, on dry cell basis).

In a series of investigations, it has been demonstrated that the

oleaginous microorganisms do not possess a hyperactive sys-

tem of fatty acid biosynthesis, but they are capable of pro-

ducing in significant quantities, acetyl-CoA, the basic unit of

fatty acid biosynthesis [3, 20]. The biochemistry, thus, of de

novo lipid biosynthesis, may be divided in two distinct par-

ties: the intermediate cellular metabolism and the biosyn-

thesis of TAGs.

The net product of glycolysis is pyruvic acid, which passes

through the mitochondrial membrane to the mitochondrion

matrix. Pyruvate-dehydrogenase catalyzes the formation of

acetyl-CoA from pyruvic acid, and acetyl-CoA either enters

inside the Krebs cycle, or is transported again into the cyto-

plasm in order to enhance biosynthesis of cellular fatty acids

[2, 3, 18, 20]. Since the mitochondrial membrane is not

permeable by acetyl-CoA, the transformation of this com-

pound to acetyl-carnitine (catalyzed by carnitine-acyl-trans-

Eur. J. Lipid Sci. Technol. 2011, 113, 1031–1051 The biochemistry of oleaginous yeasts 1033


ferase) should be necessary for the transport of this unit into

the cytosol. Though, the capital role carnitine-acyl-transfer-

ase is exactly the opposite, namely the transport of the acetyl-

CoA, issued by b-oxidation, inside the mitochondrion

matrix. Minimal amounts of acetylcarnitine may pass

through the mitochondrion matrix in order to enter the

cytosol, and this is the case of de novo lipid biosynthesis of

the non-oleaginous microorganisms [2, 4, 20]. In the olea-

ginous microorganisms, acetyl-CoA that constitutes the pre-

cursor of intracellular biosynthesis of fatty acids derives from

breakdown of citric acid that under some circumstances has

been previously accumulated inside the mitochondria and

then is transported into the cytosol (for reviews see: Ratledge

[18, 20]; Ratledge and Wynn [2]; Fakas et al. [30]).

The key step for lipid accumulation in the oleaginous

microorganisms is the change of intracellular concentration

of various metabolites, conducted after exhaustion of some

nutrients into the culture medium [2, 7, 18, 20]. In most of

the performed studies, the essential nutrient the depletion of

which induced the accumulation of reserve lipid is that of

nitrogen, whereas the biochemistry of de novo lipid accumu-

lation of lipid has been completely elucidated only when

extracellular nitrogen in the limiting factor of microbial

growth [1–4, 7, 8, 30]. Nitrogen exhaustion provokes a rapid

decrease of the concentration of intracellular AMP (adeno-

sine monophosphate), since, by virtue of AMP-desaminase,

the microorganism cleaves AMP in IMP (inosine monophos-

phate) and NH4þ ions. The NH4

þ ions constitute a comp-

lementary nitrogen source, necessary for synthesis of cell

material after the extracellular nitrogen limitation [94].

The excessive decrease of intra-cellular AMP concentration

alters the Krebs cycle function; NADþ- (and in various cases

also NADPþ-) isocitrate dehydrogenase, enzyme responsible

for the transformation of isocitric to a-ketoglutaric acid,

losses its activity, since it is allosterically activated by intra-

cellular AMP [47, 95–97]. Thus, iso-citric acid is accumu-

lated inside the mitochondrion. This acid is found in

equilibrium with citrate (reaction catalyzed by isocitrate

acotinase). When the intra-mitochondrial citric acid concen-

tration reaches a critical value, citrate enters the cytoplasm in

exchange with malate [98]. Finally, citric acid is cleaved by

the ATP-citrate lyase (ATP-CL), the enzyme-key of lipid

accumulation process in the oil-bearing microorganisms, in

acetyl-CoA and oxaloacetate [94, 96, 99] (for reviews see:

Ratledge [18, 20]; Ratledge andWynn [2]). Acetyl-CoA, by a

quasi-inverted b-oxidation process, will generate the cellular

fatty acids (for reviews see: Ratledge and Wynn [2];

Papanikolaou and Aggelis [7]; Fakas et al. [30]). NADPH,

indispensable fatty acid biosynthesis, is provided by the inter-

mediate cellular metabolism, in which the importance of

malic enzyme has been considered as crucial for various

oil-bearing microorganisms, specifically in the cases in which

NADPH is produced exclusively by virtue of the reaction

catalyzed by the above mentioned enzyme under nitrogen-

limited conditions [2, 97]. Schematically, the intermediate

cellular metabolism of the oleaginous microorganisms in

which lipid accumulation is performed after nitrogen exhaus-

tion from the medium, is presented in Fig. 1 (adapted by

Davies and Holdsworth [3]).

ATP-CL is an enzymatic complex that is considered to be

the most important factor to account for the oleaginicity of

the various microorganisms, being absent in the non-oleagi-

nous microbial cells [18]. The reaction catalyzed through the

use of ATP-CL is depicted as follows [18, 20]:

Net product of the action of ATP-CL, therefore, is acetyl-

CoA, that will be further converted into intra-cellular fatty

acids. If ATP-CL enzymatic complex does not exist, nitrogen

exhaustion leads in the accumulation of citric acid inside the

cytoplasm. In this case, citric acid either will be excreted into

the culture medium (case of citric acid production by

Aspergillus niger and Candida sp. strains—see: Ratledge

[20]; Papanikolaou and Aggelis [7]) or will provoke the

inhibition of the 6-phosphoro-fructokinase, having as result

intracellular accumulation of polysaccharides based on

Nitrogen limitationNitrogen limitation

Function of AMPFunction of AMP--desaminasedesaminase

Degradation of intraDegradation of intra--cellular AMPcellular AMP

Deactivation of NADDeactivation of NAD-- (and potentially NADP(and potentially NADP--) dependent) dependentdehydrogenasedehydrogenase

Citric acid accumulation inside the mitochondrionCitric acid accumulation inside the mitochondrion

Citric acid transport outside the Citric acid transport outside the mitochondrion, in exchange withmitochondrion, in exchange with malatemalate

Citric acid cleavage, formation of acetylCitric acid cleavage, formation of acetyl--CoACoA,,reaction catalyzed by ATPreaction catalyzed by ATP--citrate citrate lyaselyase

Condensation reactions of acetylCondensation reactions of acetyl--CoACoA for for acylacyl--CoACoA biosynthesis biosynthesis ––Onset of fatty acids accumulationOnset of fatty acids accumulation

Figure 1. Consecutive steps leading to the de novo lipid biosynth-

esis in oleaginous microorganisms growing under nitrogen-limited

conditions (from Davies and Holdsworth [3], adapted).



6-phosporo-glucose (case of Aureobasidium pullulans—see:

Galiotou-Panayotou et al. [100]). Due to the significant

biochemical similarity between the intracellular de novo lipid

accumulation and the extracellular secretion and production

of citric acid, the last years, the yeasts have been divided and

classified by various authors as either lipid-accumulating or

citric acid-producing ones [49,101–103]. The pattern of inter-

mediate metabolism and de novo lipid biosynthesis is pre-

sented in Fig. 2.

The first reaction of fatty acid biosynthesis after acetyl-

CoA generation, is catalyzed by a biotin-dependent acetyl-

CoA carboxylase [18]. Its action is described as follows:

Acetyl-CoAþHCO3 þ ATP ! Malonyl-CoAþ Pi

This reaction is considered as the restricting step for fatty acid

biosynthesis. It is activated by the presence of citric acid in the

oleaginous strain Candida sp. 107, but not in the non-ole-

aginous strain Candida utilis [104]. In fact, in the absence of

citrate, the abovementioned enzyme exists asmultifunctional

inactive protein with a molecular size of 2.4 � 105 daltons,

while the presence of citrate induces enzyme monomers

aggregation into an active macro-structural protein with a

size of 7–9 � 106 daltons [18]. Following the generation of

malonyl-CoA, the biosynthesis of fatty acids is performed

with the aid of the multi-enzymatic complex of fatty acid

synthetase (FAS). The effectuated reaction series can be

summarized as follows [3, 30]:

Acetyl-CoAþ 7malonyl-CoAþ 14NADPH

! Palmitoyl-CoAþ 7CO2 þ 14NADPþ 7 CoASH

þ 6H2O

Generally, the multi-enzymatic complexes of FAS and

ATP-CL are inhibited by the presence of exogenous long

aliphatic chains (e.g. fatty acids, n-alkanes, etc) found into the

culture medium [34, 105]. Whatever the origin of the intra-

cellular aliphatic chains, after the biosynthesis of fatty-CoA

esters, an esterification with glycerol takes place in order for

the reserve lipids to be stocked in the form of TAGs [18, 20].

This synthesis is conducted mainly by the so-called pathway

of a-glycerol phosphate acylation (for reviews see: Ratledge

[18]; Davies and Holdsworth [3]; Athenstaedt and Daum

[106]; Mullner and Daum [107]; Fakas et al. [30]). In this

metabolic pathway, free-fatty acids are activated to coenzyme

A and are subsequently used for the acylation of the glycerol

backbone to synthesize TAGs. In the first step of TAGs

assembly, glycerol-3-phosphate (G-3-P) is acylated by G-

3-P acyltranferase (GAT) at the sn-1 position to yield 1-

acyl-G-3-P (lysophospatidic acid-LPA), which is then further

acylated by lysophosphatidic acid acyltransferase (also named

1-acyl-G-3-P acyltransferase-AGAT) in the sn-2 position to

yield phosphatidic acid (PA). This is followed by dephos-

phorylation of PA by phosphatidic acid phosphohydrolase

(PAP) to release diacylglycerol (DAG). In the final step DAG

is acylated either by diacylglycerol acyltransferase or phos-

pholipid diacylglycerol acyltransferase to produce TAGs (for

reviews see: Ratledge [18]; Davies and Holdsworth [3];

Athenstaedt and Daum [106]; Mullner and Daum [107];

Fakas et al. [30]). The steps of TAG assembly through the

a-glycerol phosphate acylation pathway, a pathway that is

very commonly used in the oleaginous microorganisms, are

depicted in Fig. 3. Alternatively, PA can be synthesized

through the dihydroxyacetone-phosphate (DHAP) pathway

as follows (studies performed principally on the non-

oleaginous yeast Saccharomyces cerevisiae—for reviews see:

Athenstaedt and Daum [106]; Mullner and Daum [107]):

DHAP is acylated at the sn-1 position by the enzyme DHAP

acyltransferase (DHAPAT). The product, 1-acyl-DHAP, is

reduced by 1-acyl-DHAP reductase (ADR) to yield 1-acyl-

G-3-P, which is further acylated to yield PA, reaction cata-

lyzed by AGAT. Finally, as far as the structure of the TAGs

produced is concerned, although their final composition

could theoretically be a random substitution of acyl-CoA

groups into glycerol, in the case of the oleaginous microor-

ganisms that have been examined, the glycerol sn-2 position

in most of the cases is occupied by unsaturated fatty acids

(therefore, vegetable-type TAGs are produced—see: Thorpe

and Ratledge [36]; Ratledge [18, 20]; Guo and Ota [108]).

Glycerol

3-P-Glycerol

Pyruvate abc

Pyruvate CH3COSCoA

Oxaloacetate

MalateMDm

dMalate

PD

CSCitrate

Iso-citrate Ac

a-ketoglutarate ICDH

Krebs

ICL

Citrate

Secretion

ACL

CH3COSCoA

Malonyl-SCoA

FAS

TAGs

Oxaloacetate

MDc

MENADPH-CO2

NADPH

NADP

GK

3-P-Dihydroxyacetone

ATPADP

Glycolysis

Hexose, pentose Glyconeogenesis

DHAkGlucose

Biomass

ATP ADP

NAD NADH

Biomass

ATP

ADP

NAD

NADH

CYTOSOL

MITOCHONDION

Figure 2. Intermediate metabolism in the oleaginous microorgan-

isms. (a–c) Systems of pyruvate transport from cytoplasm to mito-

chondrion and inversely for the malate. (d) System of citrate and

malate transport between cytoplasm and mitochondrion. Enzymes:

Ac, acotinase; ACC, acetyl-CoA carboxylase; ACL, ATP-citrate

lyase; FAS, fatty acid synthetase; ICDH, iso-citrate dehydrogenase;

MDc, malate dehydrogenase (cytoplasmic); MDm, malate dehydro-

genase (mitochondrial); PD, pyruvate dehydrogenase; PFK, phos-

pho-fructo-kinase; PK, pyruvate kinase (from Ratledge [18];

Papanikolaou and Aggelis [7], adapted).



In batch experiments, lipid accumulation involving de

novo biosynthesis pathway in the oleaginous microorganisms

is essentially a secondary anabolic activity, since, according to

the biochemical steps previously illustrated, it is enhanced

after the nitrogen depletion of the culture medium due to

‘‘metabolic overflow’’ at carbon excess conditions (for

reviews see: Ratledge [4]; Ratledge and Wynn [2]; Dyal

and Narine [41]; Fakas et al. [30]; Papanikolaou and

Aggelis [8]). However, it should be stressed in this point that

the limitation of nitrogen is not the sole factor of crucial

importance for accumulation of storage lipid during growth

of oleaginous microorganisms on sugars or similarly metab-

olized compounds. For instance, in recent investigations

it has been demonstrated that the oleaginous yeast

Rhodosporidium toruloides can present efficient production

of biomass and accumulation of storage lipid in nitrogen-rich

media, provided that phosphorus or sulphate is the limiting

factor of cell growth [14, 109]. Interestingly enough, the

cultivation on sulphate- or phosphate-limited nitrogen-rich

media induced remarkable changes in the total fatty acid

composition of storage lipids produced, with (noticeable)

increment of the cellular saturated fatty acid content of R.

toruloides when sulphate-limited conditions were imposed

[109]. In the later case, thus, with a simple fermentation

technique and based on this property of R. toruloides, a

cocoa-butter substitute was produced (for more details see

part II [155] of the current review-article). In any case, the

property of microorganisms to store significant quantities of

SCO in nitrogen-rich and phosphate- or sulphate-limited

media, can be exploited in industrial level for the valorization

of sugar-rich andnitrogen-rich agro-industrial residues [14, 109].

In general, growth of oleaginous microorganisms on sugar (or

similarly metabolized compounds) based nitrogen- (or sul-

phate- or phosphate-) limited media can be divided into two

distinct phases: (a) phase in which all nutrients are available

for themicrobial growth (the balanced growth phase). During

this phase, cell growth (including total biomass—X and lipid-

free biomass—Xf production) is carried out, and substrate

(sugar) is consumed with relatively high assimilation rates. In

this phase, cellular lipids (L) that include quantities of TAGs

are produced but, in general, they contain polar fractions (e.g.

sphingolipids, phospholipids, etc.) corresponding to mem-

brane lipids [5, 31]. In this phase, total lipid in dry matter

corresponds to a value of 5–10% w/w; (b) lipid accumulation

phase. In this phase, neutral lipids (mostly TAGs) are accu-

mulated inside the microbial cells or fungal mycelia, while

substrate (sugar) uptake could potentially present somehow

lower uptake rate compared with the balanced growth phase.

It should also be stressed that in several cases (e.g. exper-

iments performed with the oleaginousmoldsM. isabellina and

T. elegans or also with other oleaginous microorganisms),

during nitrogen-limiting phase besides lipid, also lipid-free

biomass is synthesized. This lipid-free biomass increment

despite nitrogen-limited conditions into the medium

indicates partial cell growth (and cell proliferation) and prin-

cipally accumulation of non-lipid storage materials like intra-

cellular polysaccharides [28, 29, 46]. A pattern of lipid

accumulation kinetics in an oleaginous microorganism

during growth on sugar-based nitrogen-limited media is pre-

sented in Fig. 4 (data from Papanikolaou et al. [65]). It is

easily understood that the onset of lipid accumulation is given

after exhaustion of assimilable nitrogen into the culture

medium. Furthermore, it can be seen that after complete

exhaustion of sugar from the fermentation medium, some

quantities of previously stored lipid are degraded (lipid turn-

over) in favor of lipid-free material formation.

In general, de novo synthesized yeast lipid is composed

of C16 and C18 fatty acids. Palmitic acid (C16:0) constitutes

the 15–25% w/w, of total lipids, whilst palmitoleic (D9C16:1)

is, in general, presented in percentages inferior than 5% w/w

[4, 7, 20, 24]. Likewise, stearic acid (C18:0) is generally a

minor component of the yeast lipid (5–8% w/w). Oleic acid

(D9C18:1) is the principal fatty acid accumulated inside the

yeast cells (amounts sometimes higher than 70% w/w),

while linoleic (D9,12C18:2) is found in the second position

RCOSCoA +3-GLYCEROL-PHOSPHATE

CoA

Glycerol-3-phosphate acyl transferase

LYSOPHOSPHATIDIC ACID

RCOSCoA

CoA

Lysophosphatidic acidacyl transferase

PHOSPHATIDIC ACID

Pi

H2O

Phosphatidic acid phosphohydrolase

DIACYLGLYCEROL

RCOSCoA

CoA

Diacylglycerol acyltransferase or Phospholipid diacylglycerol acyltransferase

TRIACYLGLYCEROL

Figure 3. Formation of intracellular triacylglycerols via the pathway

of a-glycerol phosphate acylation (from Ratledge [18, 4], adapted).



(15–25% w/w) [4, 20]. More unsaturated fatty acids (e.g. a-

linolenic acid—D9,12,15C18:3) are not frequently synthesized

into the yeast lipid reserves [22]. It may be assumed, there-

fore, the yeast lipid produced is, in general, composed of

unsaturated fatty acids. Only in rare cases (e.g. case of

Lipomyces starkeyi DSM 70295 growing on glucose-enriched

sewage sludge—see: Angerbauer et al. [10]) the quantity of

saturated fatty acids accumulated by the oleaginous yeasts

(principally C16:0 and to lesser extent C18:0) is higher than

50% w/w, of total lipids. In fact, the above event (the poten-

tiality of producing through de novo fatty acid accumulation

process globally saturated microbial lipids) is considered to

be the limiting step for the synthesis of microbial analogous of

expensive exotic fats (e.g. cocoa-butter) [3, 8, 20, 110].

Various strategies have been performed in order to alleviate

the above disadvantage. The fatty acid profiles of several

oleaginous yeasts when de novo lipid accumulation is per-

formed are depicted in Table 1.

2.2 Lipid production from fermentation ofhydrophobic substrates used as carbon source

2.2.1 Substrates used

A somehow restricted number of yeast strains have been

recorded to be capable of growing on fats and at the same

time accumulate significant lipid quantities. These yeasts

belong to the genera Torulopsis (T. versatilis, Torulopsis sp.),

Candida [C. tropicalis, C. guilliermondii, Yarrowia (C.) lipoly-

tica], Trichosporon, Geortichum and to the species Pichia meth-

anolica, Apiotrichum curvatun and Rhodosporidium toruloides

[21, 25, 33, 37, 38, 40, 117–128]. It is evident that the

number ofmicroorganisms that are capable to consume soaps

and free-fatty acids is higher, since culture on these materials

is done regardless of the lipolytic capacity of the microorgan-

ism used. In contrast, the microorganisms that are able to

proceed with TAG or fatty-esters break-down, should obli-

gatory possess an active lipase system into their enzymatic

arsenal [8, 17, 122, 129, 130]. As far as the yeast Y. lipolytica

is concerned, in various reports in the past years it was

considered as a non-oleaginous microorganism, since it

had been assumed as ineffectual of accumulating significant

lipid quantities from sugars or similarly metabolized com-

pounds during submerged growth in nitrogen-limited media

[20]. However, the capacity of at least some Y. lipolytica

strains to accumulate high lipid quantities (up to 60% w/

w, in dry weight) when various fats or oils were used as sole

carbon and energy source is out of question indicating the

oleaginicity of this microorganism [25, 33, 37, 40, 122, 124,

125, 128, 131] (for reviews see: Beopoulos et al. [27];

Papanikolaou and Aggelis [8]; Sabirova et al. [132]). It

should also be noticed that in at least one case a strain of

Y. lipolytica produced significant intra-cellular fat quantities

(23–43%w/w, lipid in dry weight) when cultivated on glucose

or glycerol (de novo fatty acid accumulation) in highly aer-

ated bioreactor experiments [64, 74, 76], whereas on the

other hand, small production of lipids occurred in shake-flask

experiments, in which the nitrogen limitation imposed led to

remarkable production of extra-cellular citric acid [49, 133].

The fatty materials utilized as substrate from the oleagi-

nous strains may be vegetable oils [33, 37, 119, 122, 129,

130, 134], fatty esters (methyl-, ethyl-, butyl-, or vinyl-esters

of fatty acids) [21], soap-stocks [120], pure free-fatty acids

[117, 118, 126, 127], industrial fats composed of free-fatty

acids of animal or vegetable origin [25, 40, 124, 125, 133]

and crude fish oils [38, 39, 108]. In the case of the growth of

0

2

4

6

8

10

12

0

5

10

15

20

25

30

0 50 100 150 200 250 300

X (g/L)

L (g/L)

Fru (g/L)

Bio

mas

s (X

), Li

pids

(L) [

g/L]

Fru

ctos

e (F

ru) [

g/L]

Time [h]

0

20

40

60

80

100

120

140

0

10

20

30

40

50

60

70

80

0 50 100 150 200 250 300

Ammonium ions (mg/L)

Lipid in dry weight (%, w/w)

Am

mon

ium

ion

s (m

g/L)

Lipi

d in

dry

wei

ght

(%, w

/w)

Time [h]

(a)

(b)

Figure 4. Kinetics of the oleaginous fungus Thamnidium elegans

growing on fructose in shake-flask nitrogen-limited experiments

(initial fructose at 30 g/L). Representation of biomass production

(X, g/L), lipid production (L, g/L), fructose consumption (Fru, g/L)

(a), ammonium ions assimilation and lipid in dry weight evolution (b)

(data from Papanikolaou et al. [65]).



various microorganisms on media in which hydrophobic

materials are found as substrate (or co-substrate), the

quantity of TAGs accumulated into the total accumulated

lipids, can be substantially lower compared with the lipids

produced through the de novo lipid accumulation pathway;

for instance, in the case of strains of the molds Amylomyces

rouxii and Cunninghamella blakesleeana and the yeast C. lip-

olytica cultured on soybean oil utilized as the sole substrate,

intra-cellular lipid produced contained significant free-fatty

acids quantities ranging between 30 and 83% w/w, of total

intra-lipid produced, while TAGs represented in various

cases a marginal compound of SCO produced [37].

Likewise, in the case of the oleaginous yeast Candida sp.

107, fractionation of the lipids produced from this microor-

ganism grown on various n-alkanes showed that, although the

TAGs still constituted the major SCO fraction, their relative

proportion was less than when glucose was utilized as the sole

growth substrate, while loss in TAGs was compensated for by

corresponding increase in the quantity of phospholipids [36].

Yeast lipid containing TAGs but also considerable quantities

of free-fatty acids has been reported during growth of C.

lipolytica 1094 on corn oil utilized as the sole substrate

[33]. Likewise, Aoki et al. [123] have studied the effect of

the addition of various hydrophobic materials (e.g. TAGs,

fatty acid ethyl-esters or free-fatty acids) used as glucose co-

substrates upon the biochemical behavior of the yeast P.

methanolica, and it was revealed that the intra-cellular lipid

compounds were almost of the same chemical structure with

the fatty material that was added into the medium. It was also

interesting to state that in the case of the addition of free-fatty

acids, fat accumulation ten-fold increased compared with the

addition of methyl-esters or TAGs (lipid accumulation

around 20% w/w, in dry matter against 2.5–2.8% w/w,

respectively), while in the case of added free-fatty acids as

co-substrate, SCO produced contained around 93% w/w,

free-fatty acids into the total microbial lipids stored [123].

Growth of Geotrichum sp. and C. guilliermondii on crude fish

oils resulted in the accumulation of TAGs in quantities 30–

60% w/w, into the total SCO produced (substantially lower

TAG quantities compared with de novo lipid accumulation

from sugars—see i.e.: Ratledge [18, 20]), while polar lipids or

non-identified fatty compounds were also found in noticeable

concentrations [38, 108]. Finally, growth of Y. lipolytica

ACA-DC 50109 on industrial fats composed of free-fatty

Table 1. Fatty acid composition of lipid produced by various yeast strains growing on sugars (or similarly-metabolized like glycerol,

molasses, etc) substrates in culture conditions favoring the accumulation of microbial lipid

Strain Lipid (% w/w) C16:0 C16:1 C18:0 C18:1 C18:2 C18:3 Reference

Candida sp. 107a) 37.1 37 1 14 36 7 T. Gill et al. [104]

Candida sp. 107 n.r. 28 n.r. 8 41 17 17 Davies [22]

Candida sp. 40.3 23 13 3 54 5 2 Aggelis et al. [24]

Rhodotorula gracilis 41.0 21 T. 13 51 11 3 Choi et al. [111]

Candida curvatab) 29.1 36 T. 14 40 7 T. Evans and Ratledge [59]

Candida curvatab) 28.0 37 T. 10 44 6 T. Evans and Ratledge [59]

Apiotrichum curvarumb) 31.0 34 T. 10 43 7 2 Hassan et al. [113]

Cryptococcus curvatusb) 38.0 24 T. 10 46 9 6 Hassan et al. [114]

Cryptococcus curvatusb) 25.0 18 T. 16 50 16 T. Meesters et al. [72]

Cryptococcus curvatusb) 50.0 31 - 22 42 1 n.r. Wu et al. [15]

Cruptococcus albidus 46.3 14 T. 9 53 18 2 Hansson and Dostalek [112]

Cruptococcus albidus n.r 20 n.r 11 59 6 6 Davies [22]

Yarrowia lipolytica 43.2 15 2 11 47 21 3 Papanikolaou and Aggelis [74]

Yarrowia lipolytica 30.7 12 11 9 57 11 T. Andre et al. [75]

Yarrowia lipolytica a) 22.3 13 17 6 55 7 n.r. Makri et al. [76]

Rhodosporidium toruloides 67.5 20 1 15 47 13 3 Li et al. [9]

Rhodosporidium toruloides 65.2 34 T. 13 48 1 T. Hu et al. [115]

Rhodosporidium toruloides 62.1 26 2 5 62 3 T. Wu et al. [14]

Rhodosporidium toruloides 55.6 43 T. 16 35 2 T. Wu et al. [109]

Lipomyces starkeyi 68.0 56 2 14 26 T. T. Angerbauer et al. [10]

Lipomyces starkeyi 61.5 37 4 6 49 1 T. Zhao et al. [11]

Rhodotorula mucilaginosa 48.6 22 2 9 55 11 T. Zhao et al. [116]

Trichosporon capitatum 37.6 12 1 2 74 9 n.r. Wu et al. [16]

Rhodotorula sp. 22.0 22 1 7 56 12 n.r. Chatzifragkou et al. [83]

Candida oleophila 15.3 13 3 7 66 11 n.r. Chatzifragkou et al. [83]

T. <0.5% w/w; n.r.: not reported.a) Representation of the neutral fraction of microbial lipids produced.b) Cryptococcus curvatus was formely Candida curvata and then Apiotrichum curvatum; thus these microorganisms in fact are the same species.



acids (a fully saturated derivative of tallow called ‘‘stearin’’, or

on mixtures of stearin with fully hydrolyzed oleic rapeseed

oil), resulted in SCO production in which TAGs were the

principal lipid fraction of the accumulated fat (45–55% w/w,

of total lipids), while free-fatty acids were produced in sig-

nificant amounts as well (about 30–40% w/w, of total lipids)

[25, 40]. Likewise, fundamental biochemical differences exist

between de novo lipid accumulation from nitrogen-limited

sugar-based substrates and ex novo lipid production from fats

or other hydrophobic compounds utilized as sole carbon and

energy source. These differences will be depicted in the

following chapter.

2.2.2 The biochemistry of ex novo lipid accumulation

The free-fatty acids (existed as initial substrate or produced

after lipase-catalyzed hydrolysis of the TAGs/fatty esters) are

incorporated, with the aid of active transport, inside the

microbial cell. The incorporated fatty acids are either dis-

similated for growth needs or become a substrate for endo-

cellular bio-transformations (synthesis of ‘‘new’’ fatty acid

profiles which did not exist previously in the substrate) [25,

38–40, 122, 125, 130]. The dissimilated free-fatty acids will

be degraded, by virtue of the process of b-oxidation, into

smaller chain acyl-CoAs and acetyl-CoA, reactions catalyzed

by the various acyl-CoA oxidases (Aox) (for reviews see:

Fickers et al. [135]; Beopoulos et al. [27]), providing, thus,

firstly the necessary energy for cell growth and maintenance

(channel of acetyl-CoA inside the Krebs cycle), and secondly

the formation of organic substances (intermediate metab-

olites) which constitute the precursors for the synthesis of

cellular materials [4].

The degradation of hydrophobic materials has been

studied in enzymatic and molecular level in significant details

by strains of the non-conventional yeast Y. lipolytica [35, 118,

126, 127, 136–140]. This yeast when cultivated on TAG-

type substrates has been reported to secrete an extra-cellular

lipase called Lip2p, encoded by the LIP2 gene [137]. This

gene encoded for the biosynthesis of a precursor pre-mature

protein with Lys-Arg cleavage site. The secreted lipase was

reported to be a 301-amino-acid glycosylated polypeptide,

which belongs to the TAG hydrolase family (EC 3.1.1.3).

The Lip2p precursor protein was processed by the KEX2-like

endoprotease encoded by the geneXPR6, whereas deletion of

the above gene resulted in the secretion of an active but fewer

stable pro-enzyme [27, 135, 137]. Simultaneously other

intra-cellular lipases (Lip7p and Lip8p) may also be secreted

into the culture medium, that present different fatty acid

specificities, with maximum activity being displayed againstD9C18:1, C6:0 (capronic acid) and C10:0 (capric acid) fatty

acids (for reviews see: Fickers et al. [135]; Beopoulos et al.

[27]). Then, the released free-fatty acids, produced after

lipase-catalyzed hydrolysis of the TAGs would be incorpor-

ated inside the yeast cells. It is interesting to state that for the

case ofY. lipolytica yeast, the various individual substrate fatty

acids are incorporated inside the microbial cell with different

rates [25, 40, 125], before being subjected to degradation

performed by the various intra-cellular Aox. In fact, it has

been revealed that the aforementioned biochemical process is

a multi-step reaction requiring different enzymatic activities

of five acyl-CoA oxidase isozymes (Aox1p through Aox5p),

encoded by the POX1 through POX5 genes [118, 126, 127,

135, 136, 139, 140]. Aox3p is specific for short chain acyl-

CoAs, Aox2p preferentially oxidizes long-chain acyl-CoAs

while Aox1p, Aox4p and Aox5p do not appear of being

sensitive in the chain length of the aliphatic acyl-CoA chain

[27, 135, 138, 139]. Moreover, the physiological function of

the above-mentioned oxidases has been investigated by gene

disruption [136]; mutations in Aox4 and Aox5 resulted in an

increase in total Aox activity. The growth of mutant Y. lip-

olytica strains was analyzed and in the presence of POX1 gene

only, strains did not grow on fatty acids, whereas POX4 alone

elicited partial growth, while the growth of the double POX2-

POX3-deleted mutant was normal on media containing pureD9C18:1 as the sole carbon source [136, 139]. b-Oxidation

contributes one mole of NADH and one mole of FADH2 for

every 1 mole of acetyl-CoA generated, before the entering of

acetyl-CoA inside the Krebs cycle [4, 20], and is depicted in

Fig. 5.

Principal biochemical differences exist between de novo

and ex novo lipid biosynthesis; in the later case, lipid accumu-

lation occurs simultaneously with cell growth, being entirely

independent from nitrogen exhaustion from the culture

medium [25, 40, 122, 129–131, 134]. When fats or other

hydrophobic materials are utilized as the sole carbon and

energy source, accumulation of reserve lipid is a growth-

coupled process [25, 40, 129, 131] in which lipid is accu-

mulated simultaneously with lipid-free material formation in

the presence of assimilable nitrogen into the culture medium.

A pattern of lipid accumulation kinetics in an oleaginous

microorganism during growth on fat-based media (is pre-

sented in Fig. 6 (data from Papanikolaou et al. [131]).

Moreover, in order to further demonstrate the independence

RCH2- CH 2 - CH 2- CO - SCoA RCH2- CH = CH- CO - SCoA

FAD FADH

RCH2- CHOH - CH2- CO - SCoA

+ H2O

NAD

NADH

RCH2- CO - CH2- CO - SCoACH3- CO - SCoA

RCH2- COSCoA

New cycle of reactions

+

+ CoA-SH

Figure 5. b-Oxidation process of a fatty acid with even number of

carbon atoms (from Ratledge [4], adapted).



of the process of ex novo lipid accumulation from the extra-

cellular nitrogen availability of the culture medium,

Papanikolaou et al. [131] have performed submerged cul-

tures of Y. lipolytica strain ACA-DC 50109 on stearin at two

initial concentrations (S0 ¼ 10 or 20 g/L) and different nitro-

gen concentrations [(NH4)2SO4 ¼ 0.0–10.0 g/L] and

indeed, it has been demonstrated that in all instances signifi-

cant biomass and SCO quantities were produced (Fig. 7—

data from Papanikolaou et al. [131]). However, although

total cellular lipid was produced in significant quantities

regardless of the nitrogen availability into the medium, the

higher quantities of lipid were produced when substantially

low initial nitrogen concentration media were employed

(Fig. 7). Moreover, it should be stressed that with the aid

of genetic engineering, there has been a construction of

‘‘obese’’ yeast strains (strains belonging to the species Y.

lipolytica), in which the various genes encoding to the Aox

have been subjected to disruption [126, 127]. Therefore

mutantY. lipolytica strains in which various Aox were deleted,

have been constructed [126, 127]. These strains were the

JMY 798 (MTLY 36-2P) and JMY 794 (MTLY 40-2P)

deriving from the wild strain W29 (ATCC 20460), and

despite the disruption of the genes encoding for the synthesis

of the various Aox, the genetically modified strains presented

significant and comparable growth with the W29 wild strain

during growth on pure fatty oleic acid, but in the former case

noticeable quantities of microbial lipids were accumulated as

high-size intra-cellular lipid droplets, the so-called lipid

bodies [126, 127]. The lipid bodies are considered as depots

of neutral lipids (TAGs and to lesser extent steryl-esters)

enclosed by polar fractions, while in a relatively recent devel-

opment, extensive proteomic analysis of the lipid-particle

proteins has been elaborated [35].

When fats are used as sole carbon and energy source,

there is not any de novo lipid biosynthesis from the acetyl-

CoA, since fatty acid synthetase (FAS) and ATP-citrate lyase

(ACL) are strongly inhibited by the presence of exogenous

aliphatic chains. Therefore, the presence of hydrophobic

0

5

10

15

20

25

0 50 100 150 200 250 300

X (g/L)

S (g/L)

Xf (g/L)

L (g/L)

Bio

mas

s, X

(g/L

); Li

pid-

free

bio

mas

s, X

f (g/

L);

Su

bstr

ate

fat,

S (g

/L);

Cel

lula

r lip

id, L

(g/L

)

Time (h)

0

20

40

60

80

100

0 50 100 150 200 250 300

DOT (%, v/v)

Lipid (%, w/w)

O2 s

atu

rati

on (%

, v/v

);

Lipi

d in

dry

wei

ght

(%, w

t/w

t)

Time (h)

(a)

(b)

Figure 6. Kinetics of the oleaginous yeast Yarrowia lipolytica on an

industrial derivative of tallow composed of free-fatty acids (stearin)

used as the sole carbon source. Representation of biomass produc-

tion (X, g/L), lipid production (L, g/L), lipid-free material production

(Xf, g/L), substrate fat consumption (S, g/L) (a), dissolved oxygen

tension (DOT, % v/v) and lipid in dry weight evolution (b) (data from

Papanikolaou et al. [131]).

0

10

20

30

40

50

60

0 0.25 0.5 1 2 4 5 6 10 1 2 3 4 5

Maximum accumulated lipid (%, w/w)

Perc

enta

ge (%

, w/w

)

Ammonium sulphate (g/L)

S0

SL/g 01=0=20 g/L

Figure 7. Kinetics of the oleaginous yeast Yarrowia lipolytica on an

industrial derivative of tallow composed of free-fatty acids (stearin)

used as the sole carbon source. Representation of maximum SCO

produced (% w/w, in dry mass) during growth at initial stearin con-

centration 10 or 20 g/L and variations in the initial concentration of

ammonium sulphate (data from Papanikolaou et al. [131]).



materials in the culture medium should be generally incom-

patible with de novo biogenesis of cellular lipids even if

glucose or other hydrophilic materials presented into the

medium and were co-metabolized with the exogenous fatty

acids [34, 105]. For instance, Meyer and Schweizer [105]

proceeded with growth of S. cerevisiae and C. lipolytica strains

on mixtures of glucose and pentadecanoic acid. It has been

reported that, although the extra-cellular concentration of

pentadecanoıc acid was indeed low (about 0.3% w/v), the

fatty acid synthetase activity of the strains was completely

repressed, having as a result, only the presence of odd fatty

acids into the cellular lipids. However, in more recent inves-

tigations it has been demonstrated that some de novo bio-

synthesis of fatty acids from glucose or glycerol could

potentially occur, in spite of the presence of significant

exogenous quantities of fatty materials; for instance, the

biochemical behavior of P. methanolica HA-32 was studied

in media composed of glucose (at 50 g/L) and fish oil (or its

derivatives ethyl-esters or free-fatty acids) (at 30 g/L). The

fish oil (or its derivatives) contained significant quantities of

docosahexanoic acid (DHA—D6,9,12,15,17,19C22:6), a fatty

acid that through the de novo lipid accumulation from glu-

cose can never be synthesized in P. methanolica [123].

Although the extra-cellular presence of TAGs or free-fatty

acids into the culture medium resulted in an intra-cellular

fatty acid profile similar to that of the substrate fat, and,

therefore, significant DHA quantities were detected into

the SCO produced presumably through its direct incorpora-

tion from the fatty substrate, the presence of fatty acid ethyl-

esters resulted in the synthesis of a cellular lipid containing

very restricted quantities of DHA, and presenting similarities

with the one that had been synthesized when glucose was

used as the sole carbon source [123]. Therefore, the yeast P.

methanolica could perform de novo fatty acid biosynthesis

from glucose, in spite of the presence of exogenous aliphatic

onto the culture medium [123]. Likewise, in investigations

performed with the yeast Y. lipolytica ACA-DC 50109 in

submerged cultures, in which growth was supported by the

simultaneous use of stearin (mixtures of free-fatty acids and

mainly of C16:0 and C18:0) and industrial glycerol or glu-

cose, extensive studies of the intra-cellular lipid profile of

microbial lipid produced, equally suggested some de novo

synthesis of intra-cellular fatty acids, in spite of the presence

of long-chain fatty acids found into the culturemedium [26, 102];

when Y. lipolytica ACA-DC 50109 had been cultured on

stearin utilized as the sole substrate, no dehydrogenation

or elongation reactions were conducted in cellular level

[40], while the microbial lipid produced was almost com-

pletely saturated and was composed mainly of C18:0 fatty

acid [40, 131]. Given that growth ofY. lipolytica on glucose or

glycerol used as the sole substrate is accompanied by syn-

thesis of a lipid that is globally unsaturated [49, 74, 102, 133],

enrichment of the reserve lipid with unsaturated fatty acids

during growth on glucose/stearin or glycerol/stearin mixtures

(principally D9C18:1 and D9,12C18:2) that occurred, indicates

de novo fatty acid biosynthesis despite the presence of long

aliphatic chains into the medium (see relevant results in

Table 2).

The oleaginous yeasts growing on various hydrophobic

materials (e.g. oils, fats, free-fatty acids, FAMEs), accumu-

late and at the same time modify the fatty acid composition of

the employed fatty substance utilized as substrate [21, 25, 39,

40, 108, 117, 119, 122, 125]. The phenomena that are

controlling the fatty acid composition of the cellular lipids

are the specific rate of incorporation of substrate aliphatic

chains inside themicrobial cells and the intra-cellular changes

of fatty acids defined by the enzymatic capabilities of the

microorganism [25, 122]. In addition, the (typo- and

stereo-) specificity of the microbial lipases versus the sub-

strate fat (in the case in which TAGs are used as carbon

source), can equally potentially lead to a biomodification of

the substrate fatty acid composition as a function of the

bioconversion time [33, 122, 125]. For all of the above cited

reasons, the fatty acid profile of the accumulated reserve lipid

can potentially be significantly distinguishable comparedwith

that of the initial fat substrate employed. Various oleaginous

yeasts, possessing active systems of intra-cellular desatura-

tions (desaturases D9 and D12), can potentially be implicated

in the accumulation of a more unsaturated fat compared with

the substrate [21, 117, 120, 141]. In other cases (growth on

various fish oils) yeast strains may consume more rapidly

from the substrate the saturated (C14:0 and C16:0) and

oligounsaturated (D9C18:1) fatty acids, storing into their cells

the polyunsaturated fatty acids (e.g. DHA) fatty acids, result-

ing equally in the synthesis of a SCO more unsaturated

compared with the substrate fat [39]. On the other hand,

various yeasts present the tendency to dissimilate for growth

and maintenance the unsaturated and lower aliphatic chain

fatty acids, and accumulate the saturated ones, resulting,

thus, in the synthesis of cellular lipids more saturated com-

pared with the substrate [25, 40, 122, 125]. Likewise, in other

cases, the accumulated lipid composition is almost the same

with that of the substrate [33, 37, 119], whereas, finally, it is

interesting to indicate that in some cases the fatty acid com-

position of the intra-cellular free-fatty acids can present

remarkable differences in comparison with the stored

TAGs [128]. The fatty acid profiles of the initial substrate

fat used and the accumulated lipid for fermentations per-

formed by oleaginous microorganisms is presented in

Table 3. Likewise, in various cases growth of oleaginous

microorganisms on fatty mixtures may result in selective

uptake of the various individual fatty acids from the culture

medium, and during fermentation the extra-cellular fat also

may present significant compositional differences compared

with the initial substrate fat [37, 125, 142]. The fatty acid

profiles of the initial lipid substrate and the remaining lipid

fraction after fat fermentations performed by eukaryotic

microorganisms are presented in Table 4.



2.3 Degradation of reserve lipid in the oleaginousmicroorganisms

After exhaustion or decrease in the uptake rate of the carbon

source in the culture medium the oleaginous microorgan-

isms, as a general rule, consume their own lipid reserves.

Several types of studies have been performed for this purpose

using oleaginous yeasts, molds and bacteria and under var-

ious environmental conditions, breakdown (turnover) of the

previously produced lipophilic materials was conducted, with

this process being performed simultaneously with biosyn-

thesis of new lipid-free biomass [5, 6, 25, 37, 40, 47, 48,

124, 125, 129–135, 143, 144]. Likewise, a significant part of

work has been performed in relation the mobilization of

neutral lipids in non-oleaginous yeasts (inmost cases research

performed with S. cerevisiae—see i.e.: Athenstaedt et al.

[145]; for review see: Mullner and Daum [107]).

Utilization of TAGs and steryl-esters (STEs) (the main lipid

compounds found in both oleaginous and non-oleaginous

microorganisms—see: Ratledge [20]; Mullner and Daum

[107]) from the lipid particles requires the action of TAG

lipase(s) (EC 3.1.1.3) and STE hydrolase(s) (EC 3.1.1.13).

An intra-cellular lipase’s and hydrolase’s system is expressed

either constitutively or inducibly [146, 147], being respon-

sible for the cleavage of the esters and the generation of fatty

acids that will be subsequently catabolized [34, 146, 147].

Moreover, in the case of plant or mammalian cells, in which

storage lipid mobilization is also routinely observed, some

proteins associated with the phospholipid monolayer of lipid

particles, such as perilipins (in the case of mammalian cells)

and oleosins (in the case of the plants), were assumed to be

involved in the mobilization of the neutral lipid core of the

particle by serving as docking and/or activating proteins for

TAG lipases and STE hydrolases [107]. Analysis of the most

abundant yeast lipid particle proteins by mass spectrometry

led to the identification of several polypeptides with unknown

function [145], but in any case, no one of these polypeptides,

was homologous to perilipins and oleosins [107].

The presence of lipases and STE hydrolases is a prereq-

uisite in order for onset of storage lipid turnover to be given

[107, 146, 147], while carbon starvation conditions are also

required [25, 32, 124, 131, 134, 143, 144, 146]. In a series of

investigations it has been demonstrated that the phenomenon

of cellular lipid degradation in the oleaginous microorgan-

isms was independent of the culture ‘‘pre-history’’ (meaning,

in fact, independent of the type of the carbon source assimi-

lated by the microorganism in order to proceed with reserve

lipid accumulation), since such turnover has been observed

Table 2. Composition of lipid produced by Yarrowia lipolytica ACA-DC 5010 during growth on various carbon sources and fermentation

configurations

Fatty acid composition of stearin (% w/w)

C16:0 C18:0 C18:1 C18:2

25 52 2 T.

Fatty acid composition of microbial lipid produced (% w/w)

S0 (g/L) Glol0 (g/L) Glc0 (g/L) C16:0 C18:0 C18:1 C18:2 Reference

10 – – 13 83 3 T. Papanikolaou et al. [25] a)

– 30 – 15 10 45 20 Papanikolaou et al. [133] a)

– – 30.0 11 6 53 10 Papanikolaou et al. [102] a)

– 50 – 15 11 47 21 Papanikolaou and Aggelis [74] b)

10 11 – 16 72 7 2 Papanikolaou et al. [40] c)

10 10 – 15 76 5 2 Papanikolaou et al. [26] a)



11 – 21 15 68 7 3 Papanikolaou et al. [102] a)



T. <0.5% w/w. Representation of cellular fatty acid composition in the fermentation time in which the maximum concentration of microbial

lipid had been achieved. S0, initial concentration of fatty material (stearin, that is a fully saturated tallow derivative composed of free-fatty

acids); Glol0, initial concentration of glycerol (utilization of bio-diesel derived waste glycerol as substrate); Glc0, initial concentration of

glucose (utilization of commercial-glucose as substrate). Fatty acids and glucose (or glycerol) were simultaneously consumed by the

microorganism.a) Batch flask culture.b) Single-stage continuous culture.c) Batch bioreactor culture.



Table 3. Fatty acid composition of substrate fat (S) and cellular lipids (C) during growth of oleaginous yeasts or molds on fatty materials

utilized as substrates

Fatty acid composition (% w/w)

Substrate C16:0 C18:0 C18:1 C18:2 C18:3 C20:5 C22:6 Strain Reference

Methyl-stearate S – 100 – – – – – Torupopsis sp. Matsuo et al. [21]

C 4 40 48 5 1 – –

Vinyl-esters S 35 65 – – – – – Torupopsis sp. Matsuo et al. [21]

C 26 37 22 8 T. – –

Methyl-esters S 30 70 – – – – – Candida tropicalis Matsuo et al. [21]

C 35 17 32 6 2 – –

Methyl-esters S 30 70 – – – – – Trichosporon sp. Matsuo et al. [21]

C 39 18 32 5 1 – –

Pure stearic acid S – 100 – – – – – Rhodospodidium toruloides Gierhart [117]

C 13 49 26 4 T. – –

Corn oil S 12 2 22 62 1 – – Candida lipolytica Glatz et al. [119]

C 11 2 36 41 T. – –

Palm oil S 30 2 45 11 2 – – Candida lipolytica Glatz et al. [119]

C 27 8 47 10 T. – –

Olive oil S 13 3 69 11 T. – – Candida lipolytica Glatz et al. [119]

C 12 3 70 13 T. – –

Palmitate soaps S 52 5 31 7 T. – – Candida lipolytica Montet et al. [120]

C 34 3 38 17 2 – –

Refined soybean oil S 10 4 22 56 8 – – Aspergillus flavus Koritala et al. [37]

Ca) 9 2 9 66 12 – –

Soybean oil S 10 4 22 56 8 – – Candida lipolytica Koritala et al. [37]

Ca) 7 7 21 60 9 – –

EPOb) S 7 2 11 69 7 – – Langermania gigantea Aggelis et al. [122]C 7 1 9 74 7 – –

EPO S 7 2 11 69 7 – – Rhodotorula sp. Aggelis et al. [122]

C 30 5 36 25 T. – –

EPO S 7 2 11 69 7 – – Candida lipolytica Aggelis et al. [122]

C 8 1 13 76 T. – –

Crude fish oil S 16 6 22 3 1 9 13 Geotrichum sp. Guo et al. [38]

C 20 7 21 5 2 7 16

Crude fish oil S 16 6 22 3 1 9 13 Candida guillermondii Guo and Ota [108]

C 20 7 29 6 2 4 10

Refined sardine oil S 17 5 15 2 – 22 13 Geotrichum sp. Kinoshita and Ota [39]

C 7 3 13 4 – 26 21

Tuna head oil S 18 4 26 1 – 7 27 Geotrichum sp. Kinoshita and Ota [39]

C 11 2 31 5 8 37

Tuna head oil S 18 4 26 1 – 7 27 Candida guilliermondii Kinoshita and Ota [39]

C 22 2 38 4 2 14

Stearin/HOROc) S 15 26 38 10 3 – – Yarrowia lipolytica Papanikolaou et al. [25]

50/50 C 14 50 24 4 – – –

Stearin S 25 52 2 T. – – – Yarrowia lipolytica Papanikolaou et al. [40]

C 16 80 1 – – – –

Stearin/HORO S 20 38 22 5 1 – – Yarrowia lipolytica Papanikolaou and Aggelis [125]

70/30 C 21 68 8 1 – – –


40/60 C 16 57 23 4 T. – –

Olive oil S 11 5 79 4 T. – – Yarrowia lipolytica Najjar et al. [128]

Ca) 3 1 83 8 T. – –

Olive oil S 11 5 79 4 T. – – Yarrowia lipolytica Najjar et al. [128]

Cd) 30 24 35 5 T. – –

Waste cooking olive oil S 16 2 72 7 T. – – Aspergillus sp. Papanikolaou et al. [143]

C 9 19 65 6 T. – –

Waste cooking olive oil S 16 2 72 7 T. – – Aspergillus niger Papanikolaou et al. [143]

C 14 3 78 5 T. – –

T. <0.6% w/w.a) Representation of the TAG fraction of total cellular lipids produced.b) EPO is evening primrose oil.c) Stearin: Industrial derivative of tallow composed of fully saturated free fatty acids; HORO: Fully hydrolyzed oleic rapeseed oil.d) Representation of the free-fatty acids fraction of total cellular lipids.



even though SCO accumulation previously occurred through

de novo [30, 32, 67, 144, 148] or ex novomechanism [25, 40,

124, 129, 131, 134, 143]. In any case, the released cellular

fatty acids will be catabolized via the process of b-oxidation

and the produced acetyl-CoAwill be further converted via the

Krebs cycle and the anaplerotic by-pass of glyoxylic acid [40,

67, 97, 148]; in the case in which de novo lipid accumulation

had been previously performed, during lipid turnover period,

sugar (that could potentially exist into the culture medium),

is no longer assimilated, while the function of Krebs cycle

through the utilization of NADþ- (and potentially NADPþ)-

isocitrate dehydrogenase had already been suppressed due to

extra-cellular nitrogen limitation [47, 99]. In general, the

activity of glyoxylic acid by-pass enzymes (carnitine acetyl-

transferase and iso-citrate lyase) increases considerably in

cells growing on C2 compounds (e.g. ethanol) or on sub-

strates leading to C2 unit’s formation (i.e. previously accu-

mulated TAGs, extra-cellular TAGs, hydrocarbons or free

fatty acids), while this activity is indeed reduced during

growth on glucose or other sugars [67, 142, 148, 149].

Therefore, even though microbial growth had been per-

formed on sugars or similarly metabolized compounds,

SCO turnover should be principally performed through the

glyoxylic acid by-pass pathway, while cellular nitrogen

(obtainable via AMP-desaminase) should secure the biosyn-

thesis of new lipid-free material [47]. On the other hand,

degradation of accumulated lipid when ex novo lipid

accumulation had been previously performed is conducted

indisputable principally through the glyoxylic acid anapler-

otic by-pass pathway; in fact, reserve lipid turnover generally

Table 4. Fatty acid composition of initial (S) and remaining (SR) substrate fat during growth of oleaginous yeasts or molds on fatty materials

utilized as substrates

Fatty acid composition (% w/w)

Substrate C16:0 C18:0 C18:1 C18:2 C18:3 C20:5 C22:6 Strain Reference

Refined soybean oil S 10 4 22 56 8 – – Candida lipolytica Koritala et al. [37]

SRa) 12 5 23 51 7 – –

SRb) 8 4 24 58 6 – –

Refined soybean oil S 10 4 22 56 8 – – Amylomyces rouxii Koritala et al. [37]

SRa) 12 4 22 55 8 – –

SRb) 4 1 24 59 8 – –

Refined soybean oil S 10 4 22 56 8 – – Cunninghamella

blakesleeana

Koritala et al. [37]

SRa) 18 7 24 46 6 – –

SRb) 5 2 24 61 8 – –

EPOc) S 7 2 11 69 7 – – Langermania gigantea Aggelis et al. [122]

SR 9 2 10 72 7 – –

EPO S 7 2 11 69 7 – – Rhodotorula sp. Aggelis et al. [122]

SR 7 2 10 72 8 – –

EPO S 7 2 11 69 7 – – Candida lipolytica Aggelis et al. [122]

SR 7 2 11 71 9 – –

Refined sardine oil S 17 5 15 2 – 22 13 Geotrichum sp. Kinoshita and Ota [39]

SR 5 4 10 2 – 37 19

Stearin/HOROd) S 20 38 22 5 1 – – Yarrowia lipolytica Papanikolaou and Aggelis [125]

70/30 SR 22 74 4 T. – – –


60/40 SR 15 76 8 T. – – –


40/60 SR 17 71 11 T. T. – –

Sunflower oil S 5 5 34 53 n.r. – – Yarrowia lipolytica Kamzolova et al. [142]

SR 5 24 27 35 n.r. – –

Waste cooking olive oil S 16 2 72 7 T. – – Aspergillus niger Papanikolaou et al. [143]

SR 5 4 88 3 T. – –

Waste cooking olive oil S 16 2 72 7 T. – – Penicillium expansum Papanikolaou et al. [143]

SR 11 2 83 4 T. – –

T. <0.5% w/w; n.r.: not reported.a) Free-fatty acid fraction of the residual oil.b) TAG fraction of the residual oil.c) EPO is evening primrose oil.d) Stearin: Industrial derivative of tallow composed of fully saturated free-fatty acids; HORO: Fully hydrolyzed oleic rapeseed oil.



occurs when the extra-cellular flow rate of aliphatic chains is

considerably decreased. This fact can be either due to the

apparent absence of substrate fat from the culture medium

[134, 143] or due to the presence of fat that is substantially

rich in the fatty acid C18:0 and cannot be easily catabolized

by the yeast strains due to discrimination against this fatty

acid [25, 40, 121, 124, 125]. In the case of ex novo lipid

accumulation, culture conditions could be still favorable for

microbial growth since extra-cellular nitrogen may be pre-

sented into the medium even during reserve lipid degradation

[40]. Additionally, biosynthesis of carnitine acetyl-transferase

and iso-citrate lyase had been previously induced due to the

cultivation on fatty materials [142, 143, 150]. Moreover, in

recent investigations it has been demonstrated that lack of

some nutritional components such as iron and magnesium

from the fermentation medium, can partially or even com-

pletely suppress the degradation of microbial lipids [47]. On

the other hand, utilization of multiple-limited media did not

negatively influence the process of storage lipid accumu-

lation, and therefore utilization of such types of media

presents obvious interest for the process of SCO fermentation

in industrial scale [47].

Microbial lipid turnover has been extensively studied inC.

echinulata and Y. lipolytica. It has been demonstrated that C.

echinulata tended to selectively degrade its stored TAGs

instead of the other neutral lipid fractions (e.g. diacylgly-

cerols, free-fatty acids, STE etc) [32]. Moreover, the intra-

cellular saturated and monounsaturated fatty acids were rap-

idly and selectively consumed, while the polyunsaturated

ones (e.g. D9,12C18:2, D6,9,12C18:3) were not. Therefore,

after accomplishment of SCO turnover, the remaining cel-

lular lipid was drastically more unsaturated compared with

the originally accumulated lipid [32]. On the contrary, Y.

lipolytica tended to dissimilate for growth and energy require-

ments the unsaturated fatty acids regardless of their concen-

trations into the stored lipid reserves, and thus, after storage

lipid turnover, the remaining cellular lipid was almost

completely saturated [125]. Mobilization of microbial

lipophilic compounds has also been studied in oleaginous

bacteria; in this case the tested strains had previously

accumulated both TAGs and poly-hydroxy-alkanoates

(PHAs), and as in the case of oleaginous yeasts and molds

turnover of storage materials occurred due to shifting from

carbon excess to carbon limited conditions [146]. In this case,

it has been revealed that simultaneous consumption of

TAGs and PHAs at the stationary growth phase was

performed [146].

Reserve lipid turnover was in general accompanied by

generation of lipid-free material (Xf) providing evidence that

accumulated lipids act as precursor in the biosynthesis of cell

materials. Holdsworth and Ratledge [144] have proceeded

with a two-stage fermentation of the oleaginous yeast C.

curvata growing on glucose. In the first stage, lipid accumu-

lation was enhanced (growth on glucose, initial molar ratio

C/N ¼ 60), while, in the second stage, no extra-cellular car-

bon source was employed. The microorganism used its

accumulated lipids; in this case, the obtained yield YXf/L

(g of lipid-free biomass formed per 1 g of cellular lipid

consumed) was about 1.9 g/g [144] that can be considered

as somehow elevated, since the maximum theoretical

yield of conversion of TAGs into lipid-free biomass is around

1.7 g/g [30, 32, 144]. The same yield presented drastically

lower values (0.6–0.8 g/g) during growth of Y. lipolytica

ACA-DC 50109 on industrial fats composed of mixtures

of saturated and unsaturated fatty acids [25, 40, 124], whilst

growth of Mucor circinelloides on vegetable oils was accom-

panied by a yield YXf/L 0.63–0.66 g/g [134]. Finally, growth

of the mold C. echinulata on glucose enriched tomato

waste hydrolysate was accompanied by cellular lipid degra-

dation with yield YXf/L value 0.95 g/g [32], with the respect-

ive value for M. isabellina cultured on glucose being around

1.0 � 0.1 g/g [47] and on starch being slightly lower (0.7–

0.8 g/g) [48].

3 Concluding remarks—future perspectives

The production of SCOs presents continuous expansion the

last years. Amongst microbial lipids, yeast lipids present a

high importance in both academic and industrial point of

view. Due to their unicellular nature and their potentiality to

grow on a plethora of hydrophilic or hydrophobic substrates,

the oleaginous yeasts are considered as perfect ‘‘tools’’ for

studying phenomena of advanced lipid biochemistry and

biotechnology [4, 8, 132, 135]. Interestingly, the last years

there have been works indicating interesting analogies

between lipid metabolism in the yeast cells and the gastro-

intestinal tract and vascular system [128, 151].

Fundamental differences in biochemical level exist

between de novo lipid accumulation from hydrophilic sub-

strates and ex novo lipid accumulation from hydrophobic

substrates. In the former case, lipid production is a secondary

anabolic activity occurring after nitrogen had been depleted

from the growth medium. On the other hand, ex novo lipid

production is a growth associated process occurring simul-

taneously with cell growth, being entirely independent from

nitrogen exhaustion from the culture medium. The major

industrial applications related with the de novo accumulation

of storage lipid in yeasts refer to the production of ‘‘non-

specified’’ lipids that will be further converted into FAMEs

(2nd generation bio-diesel), since the conventional utilization

of edible oils as starting materials for the synthesis of bio-

diesel has resulted in significant increase in their price the last

years [1]. Moreover, through the de novo lipid accumulation,

it is possible to produce yeast lipids that have a fatty acid

composition similar to that of cocoa-butter [3, 4, 19, 20]. The

prospective (significant) rise in the price of cocoa-butter

indicates again the potentiality of the utilization of oleaginous

yeasts to produce cocoa butter-like cellular lipids. Needless

to indicate that for both the above mentioned potentialities,

the utilization of low- or even negative-cost hydrophilic



materials as substrates should be envisaged (hemicelluloses

hydrolysates, bio-diesel derived waste glycerol, whey, sugar-

based wastes, etc.), in order for an economically viable bio-

process to be performed.

The process of lipid production from fatty materials is

critically influenced by the fatty acid composition of the fat

used as substrate. ‘‘New’’ fatty acid profiles (in both extra-

and intra-cellular level) that did not previously exist in the

substrate fat may be produced during fat fermentation led by

the oleaginous yeasts. An important application of the ex

novo lipid production refers to the valorization of fatty wastes

and the production of lipid-rich microbial biomass (contain-

ing in several cases valuable PUFAs) that will further be used

as food or feed supplement [33, 38, 40, 108, 119, 120, 124,

131, 143]. On the other hand, themost important application

of the ex novo lipid accumulation process in the oleaginous

microorganisms refers to the ‘‘improvement’’ and the ‘‘up-

grade’’ of the fatty materials utilized as substrates in order to

produce ‘‘tailor-made’’ lipids of high-added value. Such cases

refer to the valorization of low-cost saturated fatty acids so as

to produce intra-cellular substitutes of cocoa-butter or other

high-value exotic fats like shea butter, sal fat etc (for review

see: Papanikolaou and Aggelis [8]) or the valorization of low-

or negative-cost unsaturated fatty substances in order to

produce lipids containing themedically important g-linolenic

acid [129, 130, 152].

The number of research investigations on the topic of

SCO fermentation presents a significant expansion the last

years. Research teams work on the identification and selec-

tion of ‘‘new’’ naturally occurring oleaginous strains capable

of storing huge lipid quantities inside their cells or mycelia.

Optimization of culture conditions related with the maximi-

zation of SCO production is being currently performed.

These studies present interest for future experiments.

Likewise, studies that concern advanced lipid biochemistry,

identification of limiting steps related with the accumulation

of storage lipid inside the cells or mycelia of the oleaginous

microorganisms and extensive analysis of microbial lipid

produced are also performed, while, equally, these studies

present interest for future experimental work. The same is

stressed in relation with investigations dealing with cloning of

specific genes coding for lipid-accumulating key enzymes

[153], kinetic modeling [29, 102, 124, 129, 130] and meta-

bolic flux analysis and finally proteomic—genomic analysis

[154] for several oleaginous microorganisms.

Financial support concerning the results achieved by our research

team has been kindly provided by: Agricultural University of

Athens; University of Patras; Dracoil SA; Project BIOSIS

(INTERREG III GREECE, ITALY); State Scholarship

Foundation (Athens, Greece); General Secretary of Research and

Technology (Ministry of Development, Greek Government).

The authors have declared no conflict of interest.

References

[1] Ratledge, C., Cohen, Z., Microbial and algal lipids: Do theyhave a future for biodiesel or as commodity oils? LipidTechnol. 2008, 20, 155–160.

[2] Ratledge, C., Wynn, J., The biochemistry and molecularbiology of lipid accumulation in oleaginous microorgan-isms. Adv. Appl. Microbiol. 2002, 51, 1–51.

[3] Davies, R. J., Holdsworth, J. E., Synthesis of lipids in yeasts:Biochemistry, physiology and production. Adv. Appl. LipidRes. 1992, 1, 119–159.

[4] Ratledge, C., Microbial lipids, in: Kleinkauf, H., Dohren,H. (Eds.), Biotechnology, 2nd Edn., Vol. 7, Wiley-VCH,Weinheim (Germany) 1997, pp. 135–197.

[5] Fakas, S., Certik, M., Papanikolaou, S., Aggelis, G., et al.,g-Linolenic acid production by Cunninghamella echinulatagrowing on complex organic nitrogen sources. Bioresour.Technol. 2008, 99, 5986–5990.

[6] Fakas, S., Papanikolaou, S., Galiotou-Panayotou, M.,Komaitis, M., Aggelis, G., Organic nitrogen of tomatowaste hydrolysate enhances glucose uptake and lipidaccumulation in Cunninghamella echinulata. J. Appl.Microbiol. 2008, 105, 1062–1070.

[7] Papanikolaou, S., Aggelis, G., Biotechnological valorizationof biodiesel derived glycerol waste through production ofsingle cell oil and citric acid by Yarrowia lipolytica. LipidTechnol. 2009, 21, 83–87.

[8] Papanikolaou, S., Aggelis, G., Yarrowia lipolytica: A modelmicroorganism used for the production of tailor-made lip-ids. Eur. J. Lipid Sci. Technol. 2010, 112, 639–654.

[9] Li, Y. H., Zhao, Z. B., Bai, F. W., High density cultivationof oleaginous yeastRhodosporidium toruloides Y4 in fed-batchculture. Enzyme Microb. Technol. 2007, 41, 312–317.

[10] Angerbauer, C., Siebenhofer, M., Mittelbach, M., Guebitz,G.M., Conversion of sewage sludge into lipids by Lipomycesstarkeyi for biodiesel production. Bioresour. Technol. 2008,99, 3051–3056.

[11] Zhao, X., Kong, X., Hua, Y., Feng, B., Zhao, Z. B.,Medium optimization for lipid production through co-fer-mentation of glucose and xylose by the oleaginous yeastLipomyces starkeyi. Eur. J. Lipid Sci. Technol. 2008, 110,405–412.

[12] Zhu, L. Y., Zong,M. H., Wu, H., Efficient lipid productionwith Trichosporon fermentans and its use for biodiesel prep-aration. Bioresour. Technol. 2008, 99, 7881–7885.

[13] Subramaniam, R., Dufreche, S., Zappi, M., Bajpai, R.,Microbial lipids from renewable resources: Productionand characterization. J. Ind. Microbiol. Biotechnol. 2010,37, 1271–1287.

[14] Wu, S., Hu, C., Jin, C., Zhao, X., Zhao, Z. B., Phosphate-limitation mediated lipid production by Rhodosporidium tor-uloides. Bioresour. Technol. 2010, 101, 6124–6129.

[15] Wu, S., Hu, C., Zhao, X., Zhao, Z. B., Production of lipidfrom N-acetylglucosamine by Cryptococcus curvatus. Eur. J.Lipid Sci. Technol. 2010, 112, 727–733.

[16] Wu, H., Li, Y., Chen, L., Zong, M. H., Production ofmicrobial oil with high oleic acid content by Trichosporoncapitatum. Appl. Energy 2011, 88, 138–142.

[17] Ratledge, C., Boulton, C. A., Fats and Oils, in: Moo-Young, M., Blanch, H. N., Drew, S., Wang, D. I. C.(Eds.), Comprehensive Biotechnology, the Principles,



Applications and Regulations of Biotechnology in Industry,Agriculture and Medicine, Pergamon Press, New York(USA) 1985, pp. 983–1003.

[18] Ratledge, C., Biochemistry, stoichiometry, substrates andeconomics, in: Moreton, R. S. (Ed.), Single Cell Oil,Longman Scientific & Technical, Harlow (UK) 1988,pp. 33–70.

[19] Ratledge, C., Single cell oils – have they a biotechnologicalfuture? Trends Biotechnol. 1993, 11, 278–284.

[20] Ratledge, C., Yeasts, moulds, algae and bacteria as sourcesof lipids, in: Kamel, B. S., Kakuda, Y. (Eds.), TechnologicalAdvances in Improved and Alternative Sources of Lipids,Blackie Academic and Professional, London (UK) 1994,pp. 235–291.

[21] Matsuo, T., Terashima, M., Hashimoto, Y., Hasida, W.,US Patent 4308 350, 1981.

[22] Davies, R. J., Yeast oil from cheese whey – process develop-ment, in: Moreton, R. S. (Ed.), Single Cell Oil, LongmanScientific & Technical, Harlow (UK) 1988, pp. 99–146.

[23] Moreton, R. S., Physiology of lipid accumulating yeasts, in:Moreton, R. S. (Ed.), Single Cell Oil, Longman Scientific &Technical, Harlow (UK) 1988, pp. 1–32.

[24] Aggelis, G., Stathas, D., Tavoularis, N., Komaitis, M.,Composition of lipids produced by some strains ofCandida species. Production of single-cell oil in a chemostatculture. Folia Microbiol. 1996, 41, 299–302.

[25] Papanikolaou, S., Chevalot, I., Komaitis, M., Aggelis, G.,Marc, I., Kinetic profile of the cellular lipid composition inan oleaginous Yarrowia lipolytica capable of producing acocoa-butter substitute from industrial fats. Antonie VanLeeuwenhoek 2001, 80, 215–224.

[26] Papanikolaou, S., Muniglia, L., Chevalot, I., Aggelis, G.,Marc, I., Accumulation of a cocoa-butter-like lipid byYarrowia lipolytica cultivated on agro-industrial residues.Curr. Microbiol. 2003, 46, 124–130.

[27] Beopoulos, A., Cescut, J., Haddouche, R., Uribelarrea,J. L., et al., Yarrowia lipolytica as a model for bio-oil pro-duction. Prog. Lipid Res. 2009, 48, 375–387.

[28] Fakas, S., Papanikolaou, S., Batsos, A., Galiotou-Panayotou, M., et al., Evaluating renewable carbon sourcesas substrates for single cell oil production byCunninghamellaechinulata and Mortierella isabellina. Biomass Bioenergy 2009,33, 573–580.

[29] Fakas, S., Bellou, S., Makri, A., Aggelis, G., Single cell oiland gamma-linolenic acid production by Thamnidium ele-gans grown on raw glycerol, in: Aggelis, G. (Ed.), MicrobialConversions of Raw Glycerol, Nova Science Publishers Inc.,New York (USA) 2009, pp. 85–99.

[30] Fakas, S., Papanikolaou, S., Galiotou-Panayotou, M.,Komaitis, M., Aggelis, G., Biochemistry and biotechnologyof single cell oil, in: Pandey, A., Larroche, C., Soccol, C. R.,Dussard, C. G. (Eds.), New Horizons in Biotechnology,AsiaTech Publishers Inc., New Delhi (India) 2009,pp. 38–60.

[31] Fakas, S., Papanikolaou, S., Galiotou-Panayotou, M.,Komaitis, M., Aggelis, G., Lipids of Cunninghamellaechinulata with emphasis to g-linolenic acid distributionamong lipid classes. Appl. Microbiol. Biotechnol. 2006, 73,676–683.

[32] Fakas, S., Galiotou-Panayotou, M., Papanikolaou, S.,Komaitis, M., Aggelis, G., Compositional shifts in lipid

fractions during lipid turnover in Cunninghamella echinulata.Enzyme Microb. Technol. 2007, 40, 1321–1327.

[33] Bati, N., Hammond, E. G., Glatz, B. A., Biomodification offats and oils: Trials with Candida lipolytica. J. Am. Oil Chem.Soc. 1984, 61, 1743–1746.

[34] Alvarez, H. M., Kalscheuer, R., Steinbuchel, A.,Accumulation of storage lipids in species of Rhodococcusand Nocardia and effect of inhibitors and polyethylene gly-col. Fett-Lipid 1997, 99, 239–246.

[35] Athenstaedt, K., Jolivet, P., Boulard, C., Zivy, M., et al.,Lipid particle composition of the yeast Yarrowia lipolyticadepends on the carbon source. Proteomics 2006, 6, 1450–1459.

[36] Thorpe, R. F., Ratledge, C., Fatty acid distribution intriglycerides of yeasts grown on glucose and n-alkanes.J. Gen. Microbiol. 1972, 75, 151–163.

[37] Koritala, S., Hesseltine, C. W., Pryde, E. H., Mounts,T. L., Biochemical modification of fats by microorganisms:A preliminary study. J. Am. Oil Chem. Soc. 1987, 64, 509–513.

[38] Guo, X., Tomonaga, T., Yanagihara, Y., Ota, Y., Screeningfor yeasts incorporating the exogenous eicosapentaenoicand docosahexaenoic acids from crude fish oil. J. Biosci.Bioeng. 1999, 87, 184–188.

[39] Kinoshita, H., Ota, Y., Concentration of docosahexaenoicacid from fish oils using Geotrichum sp. FO347-2. Biosci.Biotechnol. Biochem. 2001, 65, 1022–1026.

[40] Papanikolaou, S., Chevalot, I., Komaitis, M., Marc, I.,Aggelis, G., Single cell oil production by Yarrowia lipolyticagrowing on an industrial derivative of animal fat in batchcultures. Appl. Microbiol. Biotechnol. 2002, 58, 308–312.

[41] Dyal, S. D., Narine, S. S., Implications for the use ofMortierella fungi in the industrial production of essentialfatty acids. Food Res. Int. 2005, 38, 445–467.

[42] Aggelis, G., Ratomahenina, R., Arnaud, A., Galzy, P., et al.,Etude de l’influence des conditions de culture sur la teneuren acide gamma linolenique de souches de Mucor.Oleagineux 1988, 43, 311–317.

[43] Aggelis, G., Pina, M., Graille, J., Localisation de l’acidegamma linolenique dans les myceliums et les spores chezdeux Mucorales. Oleagineux 1990, 45, 229–232.

[44] Aggelis, G., Komaitis, M., Dimitroulias, G., Pina, M.,Graille, J., Possibilite de production d’acide gamma linole-nique par culture de Mucor circinelloides CBS 172-27sur quelques huiles vegetales. Oleagineux 1991, 46, 208–212.

[45] Chen, H. C., Chang, C. C., Production of g-linolenic acidby the fungus Cunninghamella echinulata CCRC 31840.Biotechnol. Prog. 1996, 12, 338–341.

[46] Papanikolaou, S., Komaitis, M., Aggelis, G., Single celloil (SCO) production by Mortierella isabellina grown onhigh-sugar content media. Bioresour. Technol. 2004, 95,287–291.

[47] Papanikolaou, S., Sarantou, S., Komaitis, M., Aggelis, G.,Repression of reserve lipid turnover in Cunninghamellaechinulata and Mortierella isabellina cultivated in multiple-limited media. J. Appl. Microbiol. 2004, 97, 867–874.

[48] Papanikolaou, S., Galiotou-Panayotou, M., Fakas, S.,Komaitis, M., Aggelis, G., Lipid production by oleaginousMucorales cultivated on renewable carbon sources. Eur. J.Lipid Sci. Technol. 2007, 109, 1060–1070.



[49] Papanikolaou, S., Fakas, S., Fick, M., Chevalot, I., et al.,Biotechnological valorisation of raw glycerol dischargedafter bio-diesel (fatty acid methyl-esters) manufacturingprocess: Production of 1,3-propanediol, citric acid andsingle cell oil. Biomass Bioenergy 2008, 32, 60–71.

[50] Certik, M., Shimizu, S., Biosynthesis and regulation ofmicrobial poly-unsaturated fatty acid production.J. Biosci. Bioeng. 1999, 87, 1–14.

[51] Sakuradani, E., Shimizu, S., Single cell oil production byMortierella alpina. J. Biotechnol. 2009, 114, 31–36.

[52] Sakuradani, E., Ando, A., Ogawa, J., Shimizu, S., Improvedproduction of various polyunsaturated fatty acids throughfilamentous fungus Mortierella alpina breeding. Appl.Microbiol. Biotechnol. 2009, 84, 1–10.

[53] Damude, H. G., Zhang, H., Farrall, L., Ripp, K. G., et al.,Identification of bifunctional delta 12/omega 3 fatty aciddesaturases for improving the ratio of omega 3 to omega 6fatty acids in microbes and plants. Proc. Natl. Acad. Sci.USA 2006, 103, 9446–9451.

[54] Damude, H. G., Pollak, D.M.W., Xue, Z., Zhu, Q. Q., USPatent WO2007136646 A2, 2007.

[55] Damude, H. G., Zhu, Q. Q., US Patent WO2007127381A2, 2007.

[56] Chang, L., Chen, D., Chen, Y., Nicaud, J. M., et al.,Production of functional g-linolenic acid (GLA) by expres-sion of fungal D12 and D6 desaturase genes in the oleagi-nous yeast Yarrowia lipolytica, in: Ching, J. F. (Ed.),Biocatalysis and Agricultural Biotechnology, CRC Press,Taylor & Francis Group, Boca Raton (USA) 2009,pp. 164–179.

[57] Peng, X.-W., Chen, H.-Z., Microbial oil accumulation andcellulase secretion of the endophytic fungi from oleaginousplants. Ann. Microbiol. 2007, 57, 239–242.

[58] Hui, L., Wan, C., Hai-Tao, D., Xue-Jiao, C., et al., Directmicrobial conversion of wheat straw into lipid by a cellulo-lytic fungus of Aspergillus oryzae A-4 in solid-state fermen-tation. Bioresour. Technol. 2010, 101, 7556–7562.

[59] Evans, C. T., Ratledge, C., A comparison of the oleaginousyeast Candida curvata, grown on different carbon sources incontinuous and batch culture. Lipids 1983, 18, 623–629.

[60] Ykema, A., Verbree, E. C., Kater, M. M., Smit, H.,Optimization of lipid production in the oleaginous yeastApiotrichum curvatum in whey permeate. Appl. Microbiol.Biotechnol. 1988, 29, 211–218.

[61] Ykema, A., Verbree, E. C., Nijkamp, H. J. J., Smit, H.,Isolation and characterization of fatty acid auxotrophs fromthe oleaginous yeast Apiotrichum curvatum. Appl. Microbiol.Biotechnol. 1989, 32, 76–84.

[62] Ykema, A., Verbree, E. C., Verwoert, I. I. G. S., Van derLinden, K. H., et al., Lipid production of revertants of Ufamutants from the oleaginous yeast Apiotrichum curvatum.Appl. Microbiol. Biotechnol. 1990, 33, 176–182.

[63] Brown, B. D., Hsu, K. H., Hammond, E. G., Glatz, B. A.,A relationship between growth and lipid accumulation inCandida curvata D. J. Ferm. Bioeng. 1989, 68, 344–352.

[64] Aggelis, G., Komaitis, M., Enhancement of single-cell oilproduction by Yarrowia lipolytica growing in the presence ofTeuchrium polium L. aqueous extract. Biotechnol. Lett. 1999,21, 747–749.

[65] Papanikolaou, S., Diamantopoulou, P., Chatzifragkou, A.,Philippoussis, A., Aggelis, G., Suitability of low-cost sugars

as substrates for lipid production by the fungus Thamnidiumelegans. Energy Fuel. 2010, 24, 4078–4086.

[66] Chatzifragkou, A., Fakas, S., Galiotou-Panayotou, M.,Komaitis, M., et al., Commercial sugars as substrates forlipid accumulation by Cunninghamella echinulata andMortierella isabellina fungi. Eur. J. Lipid Sci. Technol.2010, 112, 1048–1057.

[67] Vamvakaki, A.-N., Kandarakis, I., Kaminarides, S.,Komaitis, M., Papanikolaou, S., Cheese whey as a renew-able substrate for microbial lipid and biomass production byZygomycetes. Eng. Life Sci. 2010, 10, 348–360.

[68] Minkevich, I. G., Dedyukhina, E. G., Chistyakova, T. I.,The effect of lipid content on the elemental compositionand energy capacity of yeast biomass. Appl. Microbiol.Biotechnol. 2010, 88, 799–806.

[69] Huang, C., Zong, M. H., Wu, H., Liu, Q., Microbial oilproduction from rice straw hydrolysate by Trichosporon fer-mentans. Bioresour. Technol. 2009, 100, 4535–4538.

[70] Pan, L.-X., Yang, D.-F., Shao, L., Li, W., et al., Isolation ofthe oleaginous yeasts from the soil and studies of their lipid-producing capacities. Food Technol. Biotechnol. 2009, 47,215–220.

[71] Sajbidor, J., Certik, M., Dobronova, S., Influence of differ-ent carbon sources on growth, lipid content and fatty acidcomposition in four strains belonging to Mucorales.Biotechnol. Lett. 1988, 10, 347–350.

[72] Meesters, P. A. E. P., Huijberts, G.N.M., Eggink, G., Highcell density cultivation of the lipid accumulating yeastCryptococcus curvatus using glycerol as a carbon source.Appl. Microbiol. Biotechnol. 1996, 45, 575–579.

[73] Meesters, P. A. E. P., van der Wal, H., Weusthuis, R.,Eggink, G., Cultivation of the oleaginous yeastCryptococcus curvatus in a new reactor with improved mixingand mass transfer characteristics (SUPER-R). Biotechnol.Tech. 1996, 10, 277–282.

[74] Papanikolaou, S., Aggelis, G., Lipid production byYarrowia lipolytica growing on industrial glycerol in asingle-stage continuous culture. Bioresour. Technol. 2002,82, 43–49.

[75] Andre, A., Chatzifragkou, A., Diamantopoulou, P., Sarris,D., et al., Biotechnological conversions of bio-diesel derivedcrude glycerol by Yarrowia lipolytica strains. Eng. Life Sci.2009, 9, 468–478.

[76] Makri, A., Fakas, S., Aggelis, G., Metabolic activities ofbiotechnological interest in Yarrowia lipolytica grown onglycerol in repeated batch cultures. Bioresour. Technol.2010, 101, 2351–2358.

[77] Liang, Y., Cui, Y., Trushenski, J., Blackburn, J. W.,Converting crude glycerol derived from yellow grease tolipids through yeast fermentation. Bioresour. Technol.2010, 101, 7581–7586.

[78] Rymowicz, W., Fatykhova, A. R., Kamzolova, S. V.,Rywinska, A., Morgunov, I. G., Citric acid production fromglycerol-containing waste of biodiesel industry by Yarrrowialipolytica in batch, repeated batch, and cell recycle regimes.Appl. Microbiol. Biotechnol. 2010, 87, 971–979.

[79] Chi, Z., Pyle, D. J., Wen, Z., Frear, C., Chen, S.,A laboratory study of producing docosahexanoic acid frombiodiesel-waste glycerol by microalgal fermentation. Proc.Biochem. 2007, 42, 1537–1545.

[80] Pyle, D. J., Garcia, R. A., Wen, Z., Producing docosahex-aenoic acid (DHA)-rich algae from biodiesel-derived crude



glycerol: Effects of impurities on DHA production and algalbiomass composition. J. Agric. Food Chem. 2008, 56, 3933–3939.

[81] Wen, Z., Pyle, D. J., Athalye, S. K., Production of omega-3polyunsaturated fatty acids from biodiesel-derived crudeglycerol by microalgal and fungal fermentation, in:Aggelis, G. (Ed.), Microbial Conversions of Raw Glycerol,Nova Science Publishers Inc., New York (USA) 2009,pp. 41–63.

[82] Andre, A., Diamantopoulou, P., Philippoussis, A., Sarris,D., et al., Biotechnological conversions of bio-diesel derivedwaste glycerol into added-value compounds by higher fungi:Production of biomass, single cell oil and oxalic acid. Ind.Crops Prod. 2010, 31, 407–416.

[83] Chatzifragkou, A., Makri, A., Belka, A., Bellou, S., et al.,Biotechnological conversions of biodiesel derived wasteglycerol by yeast and fungal species. Energy 2011, 36,1097–1108.

[84] Eroshin, V. K., Krylova, N. I., Efficiency of lipid synthesisby yeast. Biotechnol. Bioeng. 1983, 325, 1693–1700.

[85] Yamauchi, H., Mori, H., Kobayashi, T., Shimizu, S., Massproduction of lipids by Lipomyces starkeyi in microcomputeraided fed-batch culture. J. Ferm. Technol. 1983, 61, 275–280.

[86] Emelyanova, E., Lipid and g-linolenic acid production byMucor inaquisporus. Proc. Biochem. 1997, 3, 173–177.

[87] Papanikolaou, S., Microbial conversion of glycerol into 1,3-propanediol: Glycerol assimilation, biochemical eventsrelated with 1,3-propanediol biosynthesis and biochemicalengineering of the process, in: Aggelis, G. (Ed.), MicrobialConversions of Raw Glycerol, Nova Science Publishers Inc.,New York (USA) 2009, pp. 137–168.

[88] Aggelis, G., Two alternative pathways for substrate assim-ilation byMucor circinelloides. Folia Microbiol. 1996, 41, 254–256.

[89] Roux, M. P., Kock, J. L. F., Du Preez, J. C., Botha, A., Theinfluence of dissolved oxygen tension on the production ofcocoa butter equivalents and gamma-linolenic acid byMucor circinelloides. System. Appl. Microbiol. 1995, 18,329–334.

[90] Botha, A., Strauss, T., Kock, J. L. F., Pohl, C. H., Coetzee,D. J., Carbon source utilization and g-linolenic productionby Mucoralean fungi. System. Appl. Microbiol. 1997, 20,165–170.

[91] Immelman,M., Du Preez, J. C., Kilian, S. G., Effect of C:Nratio on gamma-linolenic acid production by Mucor circi-nelloides grown on acetic acid. System Appl Microbiol. 1997,20, 158–164.

[92] Fei, Q., Chang, H. N., Shang, L., Choi, J., et al., The effectof volatile fatty acids as a sole carbon source on lipidaccumulation by Cryptococcus albidus for biodiesel pro-duction. Bioresour. Technol. 2011, 102, 2695–2701.

[93] Du Preez, J. C., Immelman, M., Kock, J. L. F., Kilian,S. G., The effect of acetic acid concentration on the growthand production of gamma-linolenic acid by Mucor circinel-loides CBS 203.28 in fed-batch culture. World J. Microbiol.Biotechnol. 1997, 13, 81–87.

[94] Evans, C. T., Ratledge, C., The role of the mitochondrialNADþ isocitrate dehydrogenase in lipid accumulation bythe oleaginous yeast Rhodosporidium toruloides CBS 14. Can.J. Microbiol. 1985, 31, 479–484.

[95] Boulton, C., Ratledge, C., Regulatory studies on citratesynthase in Candida 107, an oleaginous yeast. J. Gen.Microbiol. 1980, 121, 441–447.

[96] Boulton, C., Ratledge, C., Correlation of lipid accumu-lation in yeasts with possession of ATP-citrate lyase.J. Gen. Microbiol. 1981, 127, 169–176.

[97] Wynn, J. P., Hamid, A. A., Li, Y., Ratledge, C.,Biochemical events leading to the diversion of carbon intostorage lipids in the oleaginous fungi Mucor circinelloidesand Mortierella alpina. Microbiology UK 2001, 147, 2857–2864.

[98] Evans, C. T., Scragg, A. H., Ratledge, C., A comparativestudy of citrate efflux from mitochondria of oleaginous andnon-oleaginous microorganisms. Eur. J. Biochem. 1983,130, 195–204.

[99] Botham, P., Ratledge, C., A biochemical explanation forlipid accumulation in Candida 107 and other oleaginousmicroorganisms. J. Gen. Microbiol. 1979, 114, 361–375.

[100] Galiotou-Panayotou, M., Kalatzi, O., Aggelis, G.,Modelling of simultaneous production of polygalactorunaseand exopolysaccharide by Aureobasidium pullulansATHUM 2915. Antonie Van Leeuwenhoek 1998, 73, 155–162.

[101] Anastassiadis, S., Aivasidis, A., Wandrey, C., Citric acidproduction by Candida strains under intracellular nitrogenlimitation. Appl. Microbiol. Biotechnol. 2002, 60, 81–87.

[102] Papanikolaou, S., Galiotou-Panayotou, M., Chevalot, I.,Komaitis, M., et al., Influence of glucose and saturated free-fatty acid mixtures on citric acid and lipid production byYarrowia lipolytica. Curr. Microbiol. 2006, 52, 134–142.

[103] Papanikolaou, S., Chatzifragkou, A., Fakas, S., Galiotou-Panayotou, M., et al., Biosynthesis of lipids and organicacids by Yarrowia lipolytica strains cultivated on glucose.Eur. J. Lipid Sci. Technol. 2009, 111, 1221–1232.

[104] Gill, C., Hall, M., Ratledge, C., Lipid accumulation in anoleaginous yeast (Candida 107) growing on glucose insingle-stage continuous culture. Appl. Environ. Microbiol.1977, 33, 231–239.

[105] Meyer, K. H., Schweizer, E., Control of fatty-acid synthe-tase levels by exogenous long-chain fatty acids in the yeastsCandida lipolytica and Saccharomyces cerevisiae. Eur. J.Biochem. 1976, 65, 317–324.

[106] Athenstaedt, K., Daum, G., Phosphatidic acid, a key inter-mediate in lipid metabolism. Eur. J. Biochem. 1999, 266, 1–16.

[107] Mullner, H., Daum, G., Dynamics of neutral lipid storagein yeast. Acta Biochim. Polonica 2004, 51, 323–347.

[108] Guo, X., Ota, Y., Incorporation of eicosapentaenoic anddocosahexaenoic acids by a yeast (FO726A). J. Appl.Microbiol. 2000, 89, 107–115.

[109] Wu, S., Zhao, X., Shen, H., Wang, Q., Zhao, Z. B.,Microbial lipid production by Rhodosporidium toruloidesunder sulfate-limited conditions. Bioresour. Technol. 2011,102, 1803–1807.

[110] Smit, H., Ykema, A., Verbree, E. C., Verwoert, I. I. G. S.,Kater, M. M., Producstion of cocoa butter equivalents byyeast mutants, in: Kyle, D. J., Ratledge, C. (Eds.), IndustrialProduction of Single Cell Oils, AOCS Press, Champaign(USA) 1992, pp. 185–195.

[111] Choi, S. Y., Ryu, D. Y., Rhee, J. S., Effects of growth rateand oxygen on lipid synthesis and fatty acid composition of



Rhodotorula gracilis. Biotechnol. Bioeng. 1982, 14, 1165–1172.

[112] Hansson, L., Dostalek, M., Influence of cultivation con-ditions on lipid production by Cryptococcus albidus. Appl.Microbiol. Biotechnol. 1986, 24, 12–18.

[113] Hassan, M., Blanc, P. J., Granger, L. M., Pareilleux, A.,Goma, G., Lipid production by an unsaturated fatty acidauxotroph of the oleaginous yeast Apiotrichum curvatumgrown in single-stage continuous culture. Appl. Microbiol.Biotechnol. 1993, 40, 483–488.

[114] Hassan, M., Blanc, P. J., Pareilleux, A., Goma, G.,Selection of fatty acid auxotrophs from the oleaginousyeast Cryptococcus curvatus and production of cocoa-butterequivalents in batch mode. Biotechnol. Lett. 1994, 16, 819–824.

[115] Hu, C., Zhao, X., Zhao, J., Wu, S., Zhao, Z. B., Effects ofbiomass hydrolysis by-products on oleaginous yeastRhodosporidium toruloides. Bioresour. Technol. 2009, 100,4843–4847.

[116] Zhao, C. H., Zhang, T., Li, M., Chi, Z. M., Single cell oilproduction from hydrolysates of inulin and extract of tubersof Jerusalem artichoke by Rhodotorula mucilaginosa TJY15a.Proc. Biochem. 2010, 45, 1121–1126.

[117] Gierhart, D. I., US Patent 4485 173, 1984.

[118] Beopoulos, A., Mrozova, Z., Thevenieau, F., Le Dall,M. T., et al., Control of lipid accumulation in the yeastYarrowia lipolytica. Appl. Environ. Microbiol. 2008, 74,7779–7789.

[119] Glatz, B. A., Hammond, E. G., Hsu, K. H., Baehman, L.,et al., Production and modification of fats and oils byyeast fermentation, in: Ratledge, C., Rattray, J. B. M.,Dawson, P. S. S. (Eds.), Biotechnology for the Oils andFats Industry, AOCS Press, Champaign (USA) 1984,pp. 163–176.

[120] Montet, D., Ratomahenina, R., Galzy, P., Pina,M., Graille,J., A study of the influence of the growth media on the fattyacid composition in Candida lipolytica DIDDENS andLODDER. Biotechnol. Lett. 1985, 7, 733–736.

[121] Lee, I. M., Hammond, E. G., Cornette, J. L., Glatz, B. A.,Triacylglycerol assembly from binary mixtures of fattyacids by Apiotrichum curvatum. Lipids 1993, 28, 1055–1061.

[122] Aggelis, G., Papadiotis, G., Komaitis, M., Microbial fattyacid specificity. Folia Microbiol. 1997, 42, 117–120.

[123] Aoki, H., Miyamoto, N., Furuya, Y., Mankura, M., et al.,Incorporation and accumulation of docosahexaenoic acidfrom the medium by Pichia methanolica HA-32. Biosci.Biotechnol. Biochem. 2002, 66, 2632–2638.

[124] Papanikolaou, S., Aggelis, G., Modeling lipid accumulationand degradation in Yarrowia lipolytica cultivated on indus-trial fats. Curr. Microbiol. 2003, 46, 398–402.

[125] Papanikolaou, S., Aggelis, G., Selective uptake of fatty acidsby the yeast Yarrowia lipolytica. Eur. J. Lipid Sci. Technol.2003, 105, 651–655.

[126] Mlickova, K., Luo, Y., D’Andrea, S., Pec, P., et al., Acyl-CoA oxidase, a key step for lipid accumulation in the yeastYarrowia lipolytica. J. Mol. Catal. B: Enzym. 2004, 28, 81–85.

[127] Mlickova, K., Roux, E., Athenstaedt, K., D’Andrea, S.,et al., Lipid accumulation, lipid body formation, andacyl-coenzyme A oxidases of the yeast Yarrowia lipolytica.Appl. Environ. Microbiol. 2004, 70, 3918–3924.

[128] Najjar, A., Robert, S., Guerin, C., Violet-Asther, M.,Carriere, F., Quantitative study of lipase secretion, extra-cellular lipolysis, and lipid storage in the yeast Yarrowialipolytica grown in the presence of olive oil: Analogies withlipolysis in humans. Appl. Microbiol. Biotechnol. 2011, 89,1947–1962.

[129] Aggelis, G., Komaitis, M., Papanikolaou, S., Papadopoulos,G., Amathematicalmodel for the study of lipid accumulationin oleaginous microorganisms. I. Lipid accumulation duringgrowth of Mucor circinelloides CBS 172-27 on a vegetable oil.Gracas y Aceites 1995, 46, 169–173.

[130] Aggelis, G., Komaitis, M., Papanikolaou, S.,Papadopoulos, G., A mathematical model for the studyof lipid accumulation in oleaginous microorganisms. II.Study of cellular lipids of Mucor circinelloides during growthon a vegetable oil. Gracas y Aceites 1995, 46, 245–250.

[131] Papanikolaou, S., Chevalot, I., Galiotou-Panayotou, M.,Komaitis, M., et al., Industrial derivative of tallow: A prom-ising renewable substrate for microbial lipid, single-cellprotein and lipase production by Yarrowia lipolytica.Electron. J. Biotechnol. 2007, 10, 425–435.

[132] Sabirova, J. S., Haddouche, R., Van Bogaert, I. N., Mulaa,F., et al., The ‘‘LipoYeasts’’ project: Using the oleaginousyeast Yarrowia lipolytica in combination with specific bac-terial genes for the bioconversion of lipids, fats and oils intohigh-value products. Microb. Biotechnol. 2010, 21, 47–54.

[133] Papanikolaou, S., Muniglia, L., Chevalot, I., Aggelis, G.,Marc, I., Yarrowia lipolytica as a potential producer of citricacid from raw glycerol. J. Appl. Microbiol. 2002, 92, 737–744.

[134] Aggelis, G., Sourdis, J., Prediction of lipid accumulation –degradation in oleaginous micro-organisms growing on veg-etable oils. Antonie Van Leeuwenhoek 1997, 72, 159–165.

[135] Fickers, P., Benetti, P. H., Wache, Y., Marty, A., et al.,Hydrophobic substrate utilisation by the yeast Yarrowialipolytica, and its potential applications. FEMS Yeast Res.2005, 5, 527–543.

[136] Wang, H.-J., Le Dall, M. T., Wache, Y., Laroche, C., et al.,Evaluation of acyl coenzyme A oxidase (Aox) isozyme func-tion in the n-alkane-assimilating yeast Yarrowia lipolytica.J. Bacteriol. 1999, 181, 5140–5148.

[137] Pignede, G., Wang, H.-J., Fudalej, F., Seman, M., et al.,Autocloning and amplification of LIP2 in Yarrowia lipoly-tica. Appl. Environ. Microbiol. 2000, 66, 3283–3289.

[138] Mauersberger, S., Wang, H.-J., Gaillardin, C., Barth, G.,Nicaud, J. M., Insertional mutagenesis in the n-alkaneassimilating yeast Yarrowia lipolytica: Generation of taggedmutations in genes involved in hydrophobic substrate util-ization. J. Bacteriol. 2001, 183, 5102–5109.

[139] Luo, Y. S., Nicaud, J. M., Van Veldhoven, P., Chardot, T.,The acyl-CoA oxidases from the yeast Yarrowia lipolytica:Characterization of Aox2p. Arch. Biochem. Biophys. 2002,407, 32–38.

[140] Fickers, P., Fudalej, F., Nicaud, J. M., Destain, J., Thonart,P., Selection of new over-producing derivatives for theimprovement of extracellular lipase production by thenon-conventional yeast Yarrowia lipolytica. J. Biotechnol.2005, 115, 379–386.

[141] Gierhart, D. I., US Patent 4485 172, 1984.

[142] Kamzolova, S. V., Finogenova, T., Morgunov, I. G.,Microbiological production of citric and isocitric acids fromsunflower oil. Food Technol. Biotechnol. 2008, 46, 51–59.



[143] Papanikolaou, S., Dimou, A., Fakas, S., Diamantopoulou,P., et al., Biotechnological conversion of waste cooking oliveoil into lipid-rich biomass using Aspergillus and Penicilliumstrains. J. Appl. Microbiol. 2011, 110, 1138–1150.

[144] Holdsworth, J. E., Ratledge, C., Lipid turnover in oleagi-nous yeasts. J. Gen. Microbiol. 1988, 134, 339–346.

[145] Athenstaedt, K., Zweytick, D., Jandrositz, A., Kohlwein,S. D., Daum, G., Identification and characterization ofmajor lipid particle proteins of the yeast Saccharomycescerevisiae. J. Bacteriol. 1999, 181, 6441–6448.

[146] Alvarez, H., Kalscheuer, R., Steinbuchel, A., Accumulationand mobilization of storage lipids by Rhodococcus opacusPD630 and Rhodococcus ruber NCIMB 40126. Appl.Microbiol. Biotechnol. 2000, 54, 218–223.

[147] Waltermann, M., Luftmann, H., Baumeister, D.,Kalscheuer, R., Steinbuchel, A., Rhodococcus opacus strainPD630 as a new source of high-value single-cell oil?Isolation and characterization of triacylglycerols and otherstorage lipids. Microbiology UK 2000, 146, 1143–1149.

[148] Holdsworth, J. E., Veenhuis, M., Ratledge, C., Enzymeactivities in oleaginous yeasts accumulating and utilizingexogenous or endogenous lipids. J. Gen. Microbiol. 1988,134, 2907–2915.

[149] Kamzolova, S. V., Yusupova, A. I., Dedyukhina,E. G., Chistyakova, T. I., et al., Succinic acid synthesisby ethanol-grown yeasts. Food Technol. Biotechnol. 2009, 47,144–152.

[150] Wynn, J. P., Ratledge, C., Evidence that the rate-limitingstep for the biosynthesis of arachidonic acid in Mortierellaalpina is at the level of the 18:3 to 20:3 elongase.Microbiology UK 2000, 146, 2325–2331.

[151] Pembroke, T., The yeast Yarrowia lipolytica as a model forobesity. Nat. Inst. Health. Sci. Res. Bull. 2006, 3, 11–13.

[152] Roux-Van der Merwe, M. P., Badenhorst, J., Britz, T. J.,Fungal treatment of an edible-oil-containing industrialeffluent. World J. Microbiol. Biotechnol 2005, 21, 947–953.

[153] Tang, W., Zhang, S., Wang, Q., Tan, H., Zhao, Z. B., Theisocitrate dehydrogenase gene of oleaginous yeast Lipomycesstarkeyi is linked to lipid accumulation. Can. J. Microbiol.2009, 55, 1062–1069.

[154] Liu, H., Zhao, X., Wang, F., Jiang, X., et al., The proteomeanalysis of oleaginous yeast Lipomyces starkeyi. FEMS YeastRes. 2011, 11, 42–51.

[155] Papanikolaou, S., Aggelis, G., Lipids of oleaginousyeasts. Part II: Technology and potential applications.Eur. J. Lipid. Sci. Technol. 2011, 1052–1073.



Lipids of oleaginous yeasts. Part I: Biochemistry of single cell oil production

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