Title
Author(s)
Genetic engineering of Chlorella zofingiensis for enhanced
astaxanthinbiosynthesis and assessment of the algal oil for
biodiesel production
Liu, Jin; 刘进
Citation
Issued Date
URL
Rights
2010
http://hdl.handle.net/10722/131813
The author retains all proprietary rights, (such as patent rights)
and the right to use in future works.
GENETIC ENGINEERING OF CHLORELLA
ZOFINGIENSIS FOR ENHANCED ASTAXANTHIN
BIOSYNTHESIS AND ASSESSMENT OF THE ALGAL
OIL FOR BIODIESEL PRODUCTION
by
LIU Jin
B.Sc. (Sun Yat-sen University, P.R. China)
M.Sc. (Sun Yat-sen University, P.R. China)
A thesis submitted in fulfillment of the requirements
for the degree of Doctor of Philosophy
at The University of Hong Kong
July 2010
Abstract of thesis entitled
GENETIC ENGINEERING OF CHLORELLA
ZOFINGIENSIS FOR ENHANCED ASTAXANTHIN
BIOSYNTHESIS AND ASSESSMENT OF THE ALGAL
OIL FOR BIODIESEL PRODUCTION
Submitted by
LIU Jin
for the degree of Doctor of Philosophy
at The University of Hong Kong
in July 2010
Chlorella zofingiensis is an important microalga for astaxanthin (a
high-value ketocarotenoid) and biodiesel. In this thesis, C. zofingiensis was
genetically engineered for enhanced astaxanthin production. In addition, the
potential of the alga was assessed for biodiesel production.
C. zofingiensis was found to be very sensitive to the herbicide
norflurazon which specifically inhibits the activity of phytoene desaturase (PDS),
a rate-limiting enzyme in carotenoid biosynthesis. Mutated PDS genes resistant to
norflurazon were proposed to be used as dominant selectable markers for genetic
engineering of algae. To address this issue, C. zofingiensis mutants resistant to
norflurazon were made by treating the algal cells with chemical mutagens and
screening with norflurazon. One mutant E17 produced 54% more astaxanthin than
its wild type (WT). A point mutation (C to T) was found to occur in its PDS gene,
leading to an amino acid change (leucine to phenylalanine) at position 516. The
mutated PDS exhibited 30-fold higher resistance to norflurazon and 30% greater
desaturation activity than its WT one. The PDS-L516F gene was delivered into
WT C. zofingiensis via the biolistic approach. Transgenic C. zofingiensis showed
norflurazon resistance and accumulated up to 46.3% more astaxanthin than WT,
suggesting that the mutated PDS gene could serve as a selectable marker for
genetic engineering of C. zofingiensis and concurrently for enhanced production
of astaxanthin.
C. zofingiensis cells cultivated under different growth modes showed
differential lipid and fatty acid profiles. Compared with photoautotrophic cells, a
900% increase in lipid yield was achieved in heterotrophic cells. Furthermore
heterotrophic cells accumulated predominantly neutral lipids (NL) that accounted
for 79.5% of total lipids, with 88.7% of NL being triacylglycerol (TAG); whereas
photoautotrophic cells contained mainly glycolipids (GL) and phospholipids (PL).
Together with the much higher content of oleic acid (C18:1), oils from
heterotrophic C. zofingiensis appear to be more feasible for biodiesel production.
A number of nutritional and environmental factors were found to
influence the accumulation of fatty acids in C. zofingiensis. The optimal
nutritional and environmental conditions for total fatty acid (TFA) production by
C. zofingiensis were 5 mM nitrate, 5 mM phosphate, 25-100 μM ferrous ion,
temperature 25 °C and pH 6.5.
High light and glucose were revealed to up-regulate the expression of
both biotin carboxylase (BC) and stearoyl ACP desaturase (SAD) genes that are
involved in fatty acid biosynthesis and therefore enhanced the accumulation of
TFA including oleic acid. Unlike high light or glucose, salt stress, however, could
only trigger the up-regulation of SAD gene; accordingly, the TFA content was
slightly affected while the biosynthesis of oleic acid was promoted.
In conclusion, a selectable marker and transgenic C. zofingiensis system
have been developed for enhanced production of astaxanthin. C. zofingiensis was
shown to have great potential for biodiesel production. The study provides novel
information on both carotenoid and fatty acid biosynthesis, which is important for
the exploitation of the alga as a source of natural astaxanthin as well as biodiesel.
GENETIC ENGINEERING OF CHLORELLA
ZOFINGIENSIS FOR ENHANCED ASTAXANTHIN
BIOSYNTHESIS AND ASSESSMENT OF THE ALGAL
OIL FOR BIODIESEL PRODUCTION
by
LIU Jin
B.Sc. (Sun Yat-sen University, P.R. China)
M.Sc. (Sun Yat-sen University, P.R. China)
A thesis submitted in fulfillment of the requirements
for the degree of Doctor of Philosophy
at The University of Hong Kong
July 2010
DECLARATION
I declare that this thesis represents my own work, except where due
acknowledgement is made, and that it has not been previously included in a thesis,
dissertation or report submitted to this University or to any other institution for a
degree, diploma or other qualification.
Signed…………………………………..…………
LIU Jin
i
ACKNOWLEDGEMENTS
First and foremost, I would like to thank my supervisor Prof. Steven Feng
Chen for his helpful guidance, invaluable advice and continuous supports for my
research and life during the whole PhD study. I also thank Dr. Junchao Huang for
his great help on my works and constructive suggestions on my manuscripts and
thesis. His patience and kindness make me enjoy the research.
I am greatful to Prof. Gerhard Sandmann of Geothe University and Dr.
Yue Jiang of Hong Kong Baptist University for their advices on my manuscripts.
My thanks also extend to Dr. Mingfu Wang, Dr. Clive S.C. Lo and Dr. W.K. Yip
for their kind supply of laboratory apparatus.
Many thanks are given to my lab-mates and collegues for their kind
assistance and memorable friendship: Dr. Keith K.W. Fan, Mr. Zheng Sun, Ms.
Yujuan Zhong, Dr. Yantao Li, Dr. Guangquan Chen, Dr. Huabing Li, Dr. Yizhong
Cai, Ms. Jieqiong Huangfu, Ms. Lily R.L. Suen, Dr. Ni Sun, Dr. Bevin S.Y. Ho,
Mr. Dennis C.C. Wong, Dr. Yan Wang.
Finally, I would like to express my thanks to my parents, my young
brother and his wife, and my lovely nephew for their infinite physical and
emotional support and encouragement. To them, my appreciation never ends.
ii
PUBLICATIONS
Journal papers
Liu J, Huang J, Fan K-W, Jiang Y, Zhong Y, Sun Z, Chen F (2010) Production
potential of Chlorella zofingienesis as a feedstock for biodiesel. Bioresource
Technology 101: 8658-8663
Liu J, Huang J, Sun Z, Zhong Y, Jiang Y, Chen F (2010) Differential lipid and
fatty acid profiles of photoautotrophic and heterotrophic Chlorella zofingiensis:
Assessment of algal oils for biodiesel production. Bioresource Technology, in
press
Liu J, Zhong Y, Sun Z, Huang J, Sandmann G, Chen F (2010) One amino acid
substitution in phytoene desaturase makes Chlorella zofingiensis resistant to
norflurazon and enhances the biosynthesis of astaxanthin. Planta 232: 61-67
Sun Z, Peng X, Liu J, Fan K-W, Wang M, Chen F (2010) Inhibitory effects of
microalgal extracts on the formation of advanced glycation endproducts
(AGEs). Food Chemistry 120: 261-267
Huang JC, Liu J*, Li YT, Chen F (2008) Isolation and characterization of the
phytoene desaturase gene as a potential selective marker for genetic
engineering of the astaxanthin-producing green alga Chlorella zofingiensis
(Chlorophyta). Journal of Phycology 44: 684-690
*Co-first author
Conference papers
Liu J, Huang J, Chen F (2009) Metabolic engineering of Chlorella zofingiensis
(Chlorophyta) for enhanced biosynthesis of astaxanthin. FEBS Journal 276:
S283-S283
Liu J, Huang J, Sandmann G, Chen F (2008) Metabolic engineering for enhanced
astaxanthin biosynthesis in Chlorella zofingiensis (chlorophyta). Journal of
Biotechnology 136: S572-S572
iii
CONTENTS
DECLARATION ................................................................................................. i
ACKNOWLEDGEMENTS ................................................................................. ii
PUBLICATIONS ............................................................................................... iii
CONTENTS........................................................................................................iv
LIST OF ABBREVIATIONS ..............................................................................xi
PART I. LITERATURE REVIEW AND RESEARCH AIM ................................. 1
Chapter 1. Literature review and research aim ............................................ 2
1.1 Introduction ............................................................................................ 2
1.2 Astaxanthin ............................................................................................. 3
1.2.1 Biochemical properties of astaxanthin............................................... 3
1.2.2 Applications of astaxanthin ............................................................... 6
1.2.2.1 Astaxanthin as a coloring agent .................................................. 6
1.2.2.2 Astaxanthin as an antioxidant ..................................................... 7
1.2.3 Potential sources of astaxanthin ........................................................ 9
1.2.3.1 Synthetic astaxanthin .................................................................10
1.2.3.2 Astaxanthin from crustacean by-products ..................................10
1.2.3.3 Astaxanthin from yeast .............................................................. 11
1.2.3.4 Astaxanthin from microalgae .....................................................12
1.2.3.5 Astaxanthin from transgenic plants ............................................14
1.2.3.6 Natural astaxanthin versus synthetic astaxanthin........................14
1.2.4 Genetic modification for enhanced astaxanthin accumulation ..........15
1.2.4.1 Mutagenesis ..............................................................................15
1.2.4.2 Genetic engineering of the carotenoid biosynthetic pathway ......16
1.3 Biodiesel................................................................................................25
1.3.1 Introduction .....................................................................................25
1.3.2 Biodiesel production through transesterification ..............................28
1.3.3 Biodiesel feedstocks ........................................................................32
iv
1.3.3.1 Plant oils ...................................................................................32
1.3.3.2 Animal fats and waste oils .........................................................34
1.3.3.3 Algal oils ...................................................................................34
1.3.4 Potential and prospect of microalgal biodiesel .................................35
1.3.5 Lipid metabolism in microalgae .......................................................42
1.3.5.1 Fatty acid/lipid biosynthesis ......................................................42
1.3.5.2 Factors affecting lipid accumulation and fatty acid composition 45
1.4 The green alga Chlorella zofingiensis .....................................................48
1.4.1 Pigment profiles ..............................................................................49
1.4.2 Astaxanthin biosynthesis ..................................................................53
1.4.3 Lipid and fatty acid profiles .............................................................55
1.5 Research aim .........................................................................................55
PART II. GENETIC ENGINEERING OF CHLORELLA ZOFINGIENSIS FOR
ENHANCED ASTAXANTHIN PRODUCTION ................................................57
Chapter 2. Isolation and characterization of the phytoene desaturase gene
from Chlorella zofingiensis ............................................................................58
2.1 Abstract .................................................................................................58
2.2 Introduction ...........................................................................................58
2.3 Materials and methods ...........................................................................60
2.3.1 Algal strain and culture conditions ...................................................60
2.3.2 Genomic DNA and RNA isolation ...................................................60
2.3.3 Cloning of PDS cDNA and its corresponding gene ..........................61
2.3.4 Functional analysis of PDS cDNA ...................................................61
2.3.5 RT-PCR assay ..................................................................................62
2.3.6 Preparation of enzyme and substrate ................................................62
2.3.7 In vitro PDS assay ...........................................................................63
2.3.8 Pigment analysis ..............................................................................63
2.4 Results and discussion ...........................................................................64
2.4.1 Cloning of C. zofingiensis PDS gene................................................64
v
2.4.2 Functional analysis of C. zofingiensis PDS cDNA in E. coli ............66
2.4.3 C. zofingiensis PDS gene is up-regulated by high light and glucose .70
2.4.4 PDS-L516R is resistant to the herbicide norflurazon ........................71
Chapter 3. Isolation and characterization of Chlorella zofingiensis mutants
with enhanced biosynthesis of astaxanthin ..................................................75
3.1 Abstract .................................................................................................75
3.2 Introduction ...........................................................................................76
3.3 Methods and materials ...........................................................................76
3.3.1 Algal strain and culture conditions ...................................................76
3.3.2 Mutagenesis.....................................................................................77
3.3.4 Isolation of Chlorella mutants ..........................................................77
3.3.5 Cell dry weight determination ..........................................................77
3.3.6 Astaxanthin induction ......................................................................78
3.3.7 Extraction and analysis of pigments .................................................78
3.3.8 RNA isolation and RT-PCR assay ....................................................78
3.4 Results ...................................................................................................79
3.4.1 Carotenoid biosynthesis blocked by norflurazon ..............................79
3.4.2 Isolation of Chlorella mutants resistant to norflurazon .....................80
3.4.3 Growth and astaxanthin accumulation of mutants and WT ...............82
3.4.4 Accumulation of TC and astaxanthin induced by glucose .................84
3.4.5 Expression analysis of carotenogenic genes .....................................86
3.5 Discussion .............................................................................................86
Chpater 4. Molecular characterization of the Chlorella zofingiensis mutant
E17 .................................................................................................................90
4.1 Abstract .................................................................................................90
4.2 Introduction ...........................................................................................91
4.3 Methods and materials ...........................................................................91
4.3.1 Algal strain and culture conditions ...................................................91
4.3.2 Cell dry weight determination ..........................................................92
4.3.3 Extraction and analysis of pigments .................................................92
vi
4.3.4 Chlorophyll fluorescence measurement............................................92
4.3.5 RNA isolation and RT-PCR assay ....................................................92
4.3.6 PDS expression in E. coli.................................................................92
4.3.7 Enzyme and substrate preparation and in vitro PDS assay ................93
4.4 Results ...................................................................................................93
4.4.1 The growth and carotenogensis of E17.............................................93
4.4.2 Enhanced production of TC and astaxanthin by E17 under high light
stress or glucose induction ........................................................................95
4.4.3 Characterization of E17 PDS ...........................................................98
4.4.4 Transcription analysis of PDS, BKT and CHYb genes .................... 100
4.5 Discussion ........................................................................................... 101
Chapter 5. Transformation of Chlorella zofingiensis with a phytoene
desaturase gene for enhanced astaxanthin accumulation .......................... 104
5.1 Abstract ............................................................................................... 104
5.2 Introduction ......................................................................................... 104
5.3 Materials and methods ......................................................................... 106
5.3.1 Algal strain and culture conditions ................................................. 106
5.3.2 Construction of the transformation vector pBlue-PDS-L516F and
transformation protocol .......................................................................... 107
5.3.3 Genomic DNA and RNA isolation ................................................. 108
5.3.4 PCR determination of transformants .............................................. 108
5.3.5 Cell dry weight determination ........................................................ 108
5.3.6 Extraction and analysis of pigments ............................................... 109
5.3.7 RT-PCR assay ................................................................................ 109
5.4 Results ................................................................................................. 109
5.4.1 Analysis of C. zofingiensis transformants ....................................... 109
5.4.2 Resistance of transformants to norflurazon .................................... 111
5.4.3 Enhanced biosynthesis of astaxanthin in P6 induced by glucose..... 113
5.4.4 Transcription analysis of carotenogenic genes ................................ 115
5.5 Discussion ........................................................................................... 116
vii
PART III. POTENTIAL ASSESSMENT OF CHLORELLA ZOFINGIENSIS AS A
BIODIESEL FEEDSTOCK.............................................................................. 119
Chapter 6. Differential lipid and fatty acid profiles of photoautotrophic and
heterotrophic Chlorella zofingiensis: assessment of algal oils for biodiesel
production ................................................................................................... 120
6.1 Abstract ............................................................................................... 120
6.2 Introduction ......................................................................................... 121
6.3 Methods and materials ......................................................................... 122
6.3.1 Algal strain and culture conditions ................................................. 122
6.3.2 Determination of glucose concentration, nitrate concentration, cell dry
weight and specific growth rate .............................................................. 123
6.3.3 Lipid extraction and analysis ......................................................... 123
6.3.4 Fatty acid analysis ......................................................................... 124
6.4.5 Calculation of iodine value ............................................................ 124
6.5 Results ................................................................................................. 125
6.5.1 Growth characteristics and fatty acid accumulation ........................ 125
6.5.2 Lipid class composition ................................................................. 127
6.5.3 Fatty acid composition of individual lipid classes .......................... 129
6.6 Discussion ........................................................................................... 131
Chapter 7. Sugar-based growth and lipid accumulation for oil production
in heterotrophic Chlorella zofingienesis...................................................... 134
7.1 Abstract ............................................................................................... 134
7.2 Introduction ......................................................................................... 135
7.3 Methods and materials ......................................................................... 135
7.3.1 Algal strain and culture conditions ................................................. 135
7.3.2 Pretreatment of molasses ............................................................... 136
7.3.3 Batch and fed-batch culture ........................................................... 136
7.3.4 Determination of glucose and nitrate concentration, cell dry weight
and specific growth rate.......................................................................... 137
viii
7.3.5 Lipid extraction and analysis ......................................................... 137
7.3.6 Fatty acid analysis ......................................................................... 137
7.3.7 RNA isolation and RT-PCR assay .................................................. 137
7.4 Results ................................................................................................. 138
7.4.1 Heterotrophic growth and lipid production of C. zofingiensis with
various carbon sources ........................................................................... 138
7.4.2 Fatty acid profiles of dark-grown C. zofingiensis cultures .............. 140
7.4.3 Sugars up-regulate the transcription of BC and SAD genes of C.
zofingiensis ............................................................................................ 142
7.4.4 Fed-batch fermentation enhances lipid production by C. zofingiensis
............................................................................................................... 143
7.4.5 Assessment of cane molasses as the carbon source for lipid production
by C. zofingiensis ................................................................................... 145
7.5 Discussion ........................................................................................... 146
Chapter 8. Production of fatty acids by heterotrophic Chlorella zofingiensis:
influences of nutritional and environmental factors .................................. 149
8.1 Abstract ............................................................................................... 149
8.2 Introduction ......................................................................................... 150
8.3 Methods and materials ......................................................................... 150
8.3.1 Algal strain and culture conditions ................................................. 150
8.3.2 Determination of cell dry weight.................................................... 151
8.3.3 Fatty acid analysis ......................................................................... 151
8.4 Results and discussion ......................................................................... 151
8.4.1 Nitrate ........................................................................................... 151
8.4.2 Phosphate ...................................................................................... 154
8.4.3 Ferrous ion .................................................................................... 155
8.4.4 Cultivation temperature ................................................................. 156
8.4.5 Initial pH of culture medium .......................................................... 157
Chapter 9. Isolation and characterization of biotin carboxylase gene and
stearoyl ACP desaturase gene from Chlorella zofingiensis ........................ 159
ix
9.1 Abstract ............................................................................................... 159
9.2 Introduction ......................................................................................... 160
9.3 Methods and materials ......................................................................... 160
9.3.1 Algal strain and culture conditions ................................................. 160
9.3.2 Genomic DNA and RNA isolation ................................................. 161
9.3.3 Cloning of BC cDNA, SAD cDNA and their corresponding genes .. 161
9.3.4 RT-PCR assay ................................................................................ 163
9.3.5 Fatty acid analysis ......................................................................... 163
9.4 Results ................................................................................................. 163
9.4.1 Cloning and characterization of the BC and SAD gene from C.
zofingiensis ............................................................................................ 163
9.4.2 High light irradiation up-regulates the transcripts of BC and SAD and
enhances the biosynthesis of TFA and oleic acid ..................................... 167
9.4.3 Salt stress induces the up-regulation of SAD gene and the
accumulation of oleic acid ...................................................................... 169
9.4.5 Glucose triggers the up-regulation of BC and SAD genes and induces
the enhanced biosynthesis of TFA and oleic acid .................................... 170
9.5 Discussion ........................................................................................... 171
PART IV. RESEARCH SUMMARY AND RECOMMENDATION FOR
FUTURE WORK ............................................................................................. 174
Chapter 10. Research summary and recommendation for future work ... 175
10.1 Introduction ....................................................................................... 175
10.2 Research summary ............................................................................. 175
10.3 Recommendation for future work....................................................... 179
10.3.1 Future work for astaxanthin production by C. zofingiensis ........... 179
10.3.2 Future work for biodiesel production by C. zofingiensis ............... 180
REFERENCES................................................................................................. 183
x
LIST OF ABBRECIATIONS
ACCase
acetyl-CoA carboxylase
ACP
acyl carrier protein
BC
biotin carboxylase
BKT
carotenoid ketolase
CHYb
carotenoid hydroxylase
CN
cetane number
CP
cloud point
CPTA
2-(4-chloro-phenylthio)-triethylamine
DAG
diacylglycerol
DMAPP
dimethylallyl diphosphate
DPA
diphenylamine
DTT
dithiothreitol
DUS
the degree of fatty acid unsaturation
EB
ethidium bromide
EMS
ethyl methanesulphonate
FAME
Fatty acid methyl ester
FP
flash point
FPP
farnesyl diphosphate
GC
gas chromatography
GGPP
geranylgeranyl pyrophosphate
GGPS
geranylgeranyl pyrophosphate synthase
GPP
geranyl diphosphate
GL
glycolipid
HPLC
high performance liquid chromatography
IPP
isopentenyl diphosphate
KAS
3-ketoayl ACP synthase
KV
kinematic viscosity
xi
LCY
lycopene cyclase
MAG
monoacylglycerol
MNNG
N-methyl-N′-nitro-N-nitrosoguanidine
MUFA
monounsaturated fatty acid
NL
neutral lipid
NPQ
non-photochemical quenching
NXS
neoxanthin synthase
PDS
phytoene desaturase
PL
phospholipid
PP
pour point
PSY
phytoene synthase
PUFA
polyunsaturated fatty acid
RACE
rapid amplification of cDNA ends
RNS
reactive nitrogen species
ROS
reactive oxygen species
RT-PCR
reverse transcription polymerase chain reaction
SAD
stearoyl ACP desaturase
SE
steroid ester
SFA
saturated fatty acid
TAG
triacylglycerol
TC
total carotenoids
TFA
total fatty acids
TLC
thin layer chromatography
UV
ultraviolet
WT
wild type
ZEP
zeaxanthin epoxidase
xii
PART I
LITERATURE REVIEW AND RESEARCH AIM
1
Chapter 1. Literature review and research aim
Chapter 1
Literature review and research aim
1.1 Introduction
Algae are a large and diverse group of unicellular or multicellular
organisms, ranging from the microscopic cyanobacteria (commonly referred to as
blue-green algae), which are closely related to Gram-negative bacteria, to the
giant kelps with the height of over 10 m (Graham et al., 2009). Like plants, algae
are photoautotrophic, yet some are also able to grow heterotrophically at the
expense of organic carbon sources such as sugars; in addition, some algae can be
mixotrophic, with the ability of utilizing both organic carbon substrate and light
and CO2 for growth (Radmer, 1996). Algae are dominant in water, common on
land and can also be found in unusual environments, such as on snow and in
desert soils (Lee, 2008).
Microalgae represent one of the most promising sources for numerous
products
including
carotenoids,
polyunsaturated
fatty
acids,
proteins,
polysaccharides, vitamins, and other biologically active compounds, and thus are
used in a wide variety of technological applications for the development of feed
and food products (Pulz & Gross, 2004; Gouveia et al., 2009). They are also
found to be useful in bioremediation applications for environmental clean-up
(Suresh & Ravishankar, 2004; Munoz & Guieysse, 2006). Recently, microalgae
have been considered with a great potential as a sustainable feedstock superior to
oil crops for biodiesel production (Chisti, 2007).
This chapter reviews the background of astaxanthin, approaches through
genetic engineering for enhanced astaxanthin production, current status of
biodiesel and the production potential of biodiesel using microalgae as feedstocks.
2
Chapter 1. Literature review and research aim
1.2 Astaxanthin
Astaxanthin is a red ketocarotenoid that belongs to the family of the
xanthophylls. It can be found in some microalgae, bacteria, yeasts, and many
marine animals (Lorenz & Cysewski, 2000; Zhang et al., 2009). Because of its
strong pigmentation function, powerful antioxidative activity and beneficial
effects on human health, astaxanthin has important applications in feed, food,
nutraceutical, and pharmaceutical industries (Guerin et al., 2003; Fraser &
Bramley, 2004).
1.2.1 Biochemical properties of astaxanthin
Carotenoids are a group of structurally diverse terpenoid pigments with
the isoprene as basic units. The structure of carotenoids is derived from lycopene
and the majority is the 40-carbon chain conjugated by double bonds (Figure 1.1).
Carotenoids are split into two classes, carotenes which are purely hydrocarbons
(e.g., lycopene and β-carotene) and xanthophylls which are oxygenated
derivatives of carotenes (e.g., zeaxanthin, canthaxanthin and astaxanthin) (Jin et
al., 2003).
3
Chapter 1. Literature review and research aim
Lycopene
β-carotene
OH
Zeaxanthin
OH
O
O
Canthaxanthin
O
OH
OH
O
Astaxanthin
Figure 1.1 Chemical structure of some carotenoids (Sandmann, 2001).
Asatxanthin is a hydrophobic carotenoid formed via the hydroxylation
and oxidation of β-carotene, with a chemical formula of C40H52O4 and a molecular
weight of 596.86 (Britton et al, 2004). It exists in geometric cis and trans isomers;
the latter is thermodynamically more stable than the former and found
predominantly in nature (Britton, 1995). As for the trans astaxanthin, there are
three forms of stereoisomers: two enantiomers (3R, 3′R and 3S, 3′S) and a meso
form (3R, 3′S) (Higuera-Ciapara et al., 2006). The isomers have the same
molecular and structural formula with the only difference in position of the
4
Chapter 1. Literature review and research aim
rotating functional group (Figure 1.2). Among these isomers, the 3S, 3′S is the
most abundant in nature, predominantly synthesized by green microalgae (Lorenz
& Cysewski, 2000). In contrast, synthetic astaxanthin is a racemic mixture of
these three isomers (Turujman et al., 1997).
Figure 1.2 Astaxanthin configurational isomers and its geometric cis isomer
(Higuera-Ciapara et al., 2006).
5
Chapter 1. Literature review and research aim
Astaxanthin exists in free form or esterified in its one or both hydroxyl
groups with various fatty acids such as palmitic, stearic, and oleic. Synthetic
astaxanthin is in free form while the natural one found in algae is preferred in
esterified form (Miao et al., 2006; Peng et al., 2008; Holtin et al., 2009). In
addition, astaxanthin found in marine animals may interact with proteins or
lipoproteins to form carotenoproteins or carotenolipoproteins (Ando & Tanaka,
1996; Britton et al., 1997).
1.2.2 Applications of astaxanthin
1.2.2.1 Astaxanthin as a coloring agent
With strong coloring ability, astaxanthin serves as the feed supplement
and food additive for aquaculture. The use of astaxanthin in aquaculture species,
for example salmon and lobster, has been extensively studied and well
documented in the past years (Torrissen, 1986; Laird et al., 2001; Wade et al.,
2005; Bjerkeng et al., 2007; Williams, 2007; Niamnuy et al., 2008; Choubert &
Baccaunaud, 2010). Human and animals cannot synthesize carotenoids de novo,
instead, they have to obtain carotenoids through their food chain or feeds. The
dietary carotenoids give such organisms as salmonids and crustacean the
reddish-orange color characteristic that is regarded by consumers as one of the
key quality attributes (Laird et al., 2001). Astaxanthin is the major carotenoid
found in marine animals, for example in crab the red pigment accounts for more
than 80% of total carotenoids (Shahidi & Synowiecki, 1991). Because of the
higher color intensity and better absorption by the digestive tract of salmonids,
astaxanthin is preferred over canthaxanthin in aquatic farming (Torrisen, 1986;
Storebakken & No, 1992). In addition to pigmentation, astaxanthin has been
shown to benefit the growth and survival of larval fish and shrimp (Storebakken &
Goswami 1996; Niu et al., 2009). Astaxanthin is also used in the tropical marine
6
Chapter 1. Literature review and research aim
ornamental industry and poultry, aiming to pigment ornamental fish or egg yolk
(Elwinger et al., 1997; Ako & Tamaru, 1999; Fredriksson et al., 2006).
1.2.2.2 Astaxanthin as an antioxidant
During normal metabolic processes for energy production, oxygen is
reduced, producing several oxygen-derived free radicals such as hydroxyls and
peroxides, as well as reactive oxygen species (ROS) which are considered playing
important roles in hormone biosynthesis, cell signaling and aging, and microbial
killing (Rada & Leto, 2008). ROS levels can increase dramatically when exposing
to environmental stress, for example UV or heat. Without efficient removal, the
overproducing ROS become a problem, resulting in oxidative stress that causes
DNA damage, protein and lipid oxidation, and inactivation of specific enzymes
(Stadtman, 1992; Papas, 1999). Such oxidative damage has been linked to various
diseases,
for
example,
retinopathy,
carcinogenesis,
arteriosclerosis,
and
age-related macular degeneration (Papas, 1999; Muller et al., 2007).
In addition to the enzymatic antioxidants generated by bodies (e.g., super
oxidate dismutase, catalase, and peroxidase), dietary antioxidants such as
carotenoids can serve as potent free-radical scavengers to cope with the oxidative
stress and benefit human health. The powerful antioxidant activity of astaxanthin
has been demonstrated in numerous studies in the past years (Kurashige, 1990;
Palozza & Krinsky, 1992; Lawlor & O'Brien, 1995; Shimidzu et al., 1996; Naguib,
2000; Kupcinskas et al., 2008; Liu et al., 2009a). It was reported that astaxanthin
has stronger anti-oxidative activity than other carotenoids and vitamin E
(Kurashige, 1990; Palozza & Krinsky, 1992; Naguib, 2000). The strong
anti-oxidative power of astaxanthin is indicated by the oxygen radical absorbance
capacity (ORAC) value as shown in Figure 1.3. It is the powerful anti-oxidative
properties that make astaxanthin play important roles in following human health
conditions:
7
Chapter 1. Literature review and research aim
(1) reducing the DNA damage (Santocono et al., 2006, 2007; Tripathi & Jena,
2009);
(2) protecting eyes and skin from UV-light mediated photo-oxidation (Lyons
& O'Brien, 2002);
(3) protecting membranes of cell and mitochondria from oxidative damage
(Barros et al., 2002; Goto et al., 2001; Liu et al., 2009a; Wolf et al., 2010);
(4) inhibiting lipid peroxidation that may cause plaque formation in the
circulatory system (Goto et al., 2001; McNulty et al., 2007);
(5) attenuating inflammation by quenching ROS (Bennedsen et al., 2000;
Lockwood et al., 2006; Pashkow et al., 2008);
(6) crossing the blood-brain barrier in mammals and alleviates oxidative stress,
and may help maintain neuroligic health (Hussein et al., 2005; Liu et al.,
2008);
(7) benefiting liver function by pretecting liver cells against oxidative damage
(Gradelet et al., 1998; Curek et al., 2010);
(8) preventing the initiation of tumorigenesis in the mouth, oral cavity,
prostate, large bowel, bladder and breast (Tanka et al., 1994, 1995;
Lockwood et al., 2006);
(9) boosting immune system by enhancing the production of antibody and
increase the total number of T-cells (Jyonouchi et al., 1993, 1994, 1995);
(10) benefiting heart health by modifying blood levels of LDL and HDL
cholesterol (Miki et al., 1998; Yoshida et al., 2010).
8
Chapter 1. Literature review and research aim
Figure 1.3 ORAC values of carotenoids and vitamin E. ORAC is a method for
measuring antioxidant ability of foods and other chemicals using Trolox as the
standard. A higher ORAC value represents a higher antioxidant ability (Ip, 2005).
1.2.3 Potential sources of astaxanthin
Being widely used in aquatic farming as a feed additive, astaxanthin
market in United States is estimated to be US$200 millions per year at a price of
around US$2500/kg (Lorenz & Cyswski, 2000). Commercial astaxanthin can be
produced synthetically or extracted from crustacean byproducts and some
microorganisms.
9
Chapter 1. Literature review and research aim
1.2.3.1 Synthetic astaxanthin
Nowadays, commercial astaxanthin for aquaculture is mainly produced
synthetically from petrochemical sources, whereas natural astaxanthin contributes
only to a minor portion of the market (Guerin et al., 2003). Synthetic astaxanthin
contains a mixture of isomers (3S, 3′S), (3S, 3′R), and (3R, 3′R) at the ratio of
1:2:1 (Alga Technologies, 2004). DSM and BASF are the world’s leading
suppliers of synthetic astaxanthin. Although the astaxanthin market is occupied by
synthetic products, the growing demand of customers for natural foods has urged
the production of astaxanthin from natural sources such as crustacean by-products,
yeast, microalgae and transgenic plants.
1.2.3.2 Astaxanthin from crustacean by-products
Crustacean by-products such as heads, shells and tails generated from
food processing or conditioning consist mainly of chitin, proteins, fatty acids and
carotenoids (Heu et al., 2003). The carotenoid composition and content in
crustacean by-products have been well studied (Mandeville et al., 1991; Shahidi
& Synowiecki, 1991; Olsen & Jacobsen, 1995; Sachindra & Mahendrakar, 2005;
Sachindra et al., 2007; Babu et al., 2008; Handayani et al., 2008), indicating the
potential of these by-products as natural astaxanthin source. However, these
by-products generally contains only a small amount of astaxanthin (Table 1.1) but
high contents of ash and chitin that significantly decrease fish’s digestibility
(Higuera-Ciapara et al., 2006), making the use of crustacean by-products in fish
feeding less feasible and economical. Astaxanthin found in crustacean by-products
is mainly in esterified form (Table 1.1).
10
Chapter 1. Literature review and research aim
Table 1.1 Carotenoid contents in various sources of crustacean by-products.
Adapted from Higuera-Ciapara et al. (2006)
Total
astaxanthin
(μg/100 g)
Source
Shrimp
(P. borealis)
Shrimp
(P. borealis)
Crawfish
(P. clarkii)
Backs snow crab
(Ch. opilio)
a
Astaxanthin (%)
Other carotenoids
Free
Monoester
Diester
14.77
4.0
19.7
74.3
zeaxanthin
4.97a
8
22.5
69.5
Not detected
15.3
40.3
11.96
21.2
49.4
astacene
5.1
56.6
Lutein, zeaxanthin,
astacene
mg/100 g wet basis
1.2.3.3 Astaxanthin from yeast
The red yeast Xanthophyllomyces dendrorhous (previously referred as
Phaffia rhodozyma) as the astaxanthin producer has been intensively studied in
the past years (Cruz & Parajo, 1998; Parajo et al., 1998; An et al., 2001; Ramirez
et al., 2001; Visser et al., 2003; Zheng et al., 2006; Liu & Wu, 2007; Lee et al.,
2008). X. dendrorhous can grow fast and achieve high cell biomass through
utilizing a variety of carbon sources such as glucose, xylose and even molasses.
However, the cellular astaxanthin content is relatively low, varying from 0.14 to
0.79 mg g-1 dry weight depending on different strains of X. dendrorhous (Ip,
2005). Genetic modification strategies have been employed to enhance the
astaxanthin content in X. dendrorhous cells (An et al., 1989; An et al., 1996;
Ramirez et al., 2000; Ukibe et al., 2008). Ukibe et al. (2008) addressed that the
isolated mutants of X. dendrorhous could produce 1.5-3.8 fold more astaxanthin
than the wild type cells. Currently X. dendrorhous is commercial in a fine powder
form as a natural source of astaxanthin for fish feeding. The thick cell walls of
yeast, however, hinder the assimilation of astaxanthin by fish and thus cell wall
11
Chapter 1. Literature review and research aim
disruption is needed (Storebakken et al., 2004). The X. dendrorhous derived
astaxanthin is exceptionally in the isomer form of 3R, 3′R (Johnson & An, 1991).
1.2.3.4 Astaxanthin from microalgae
Microalgae represent the most potential sources of natural astaxanthin
and have attracted thorough investigations in terms of strain screening, culture
medium optimization, stress induction, cultivation strategy modification and
genetic improvement for astaxanthin accumulation and production during the past
decades (Boussiba et al., 1992; Olaizola, 2000; Orosa et al., 2000; Hata et al.,
2001; Orosa et al., 2001; Ip and Chen, 2005; Steinbrenner & Sandmann, 2006; Hu
et al., 2008; Sandesh Kamath et al., 2008; Sun et al., 2008; Zhang et al., 2009).
Heamatococcus pluvialis has been considered as the most promising microalga
for commercial astaxanthin production, in that it is able to accumulate astaxanthin
up to 4% of its dry biomass, the highest content in nature (Boussiba, 2000).
Heamatococcus algal meal has been approved as a color additive in salmonid
feeds and as a dietary-supplement ingredient for human consumption in Japan,
USA and some other countries. In the large-scale, enclosed photobioreactor or
outdoor system, a two-step process is employed for the production of
astaxanthin-rich Heamatococcus cells (Figure 1.4): cell biomass accumulation and
astaxanthin induction. First, vegetable cells accumulate and achieve a sufficient
density under optimal growth conditions; the cell abundant culture then is
subjected to stress conditions (e.g., deprivation of nitrate, high light intensity, and
salt stress) for astaxanthin induction (Fabregas et al., 2001; Zhang et al., 2009).
12
Chapter 1. Literature review and research aim
Figure 1.4 Typical flow sheet
for the commercial production of
Haematococcus derived natural astaxanthin (Lorenz & Cysewski, 2000).
Although H. pluvialis can accumulate a high content of astaxanthin, it
grows relatively slow with a low cell biomass yield and is susceptible to
contamination and adverse environment (Lorenz & Cysewski, 2000; Olaizola,
2000; Hata et al., 2001; Ip, 2005). In addition, extremely high light illumination is
required for astaxanthin induction and accumulation in this alga and thus
hindering its commercial application (Fabregas et al., 2001; Imamoglu et al.,
2009). Recently, the green alga Chlorella zofingiensis has been regarded as a
potential alternative host for mass production of astaxanthin due to its fast growth,
low sensitivity to contamination and unfavorable environments, and astaxanthin
13
Chapter 1. Literature review and research aim
accumulation under heterotrophic conditions with glucose as the sole carbon and
energy source (Rise et al., 1994; Bar et al., 1995; Orosa et al., 2001; Del Campo et
al., 2004; Ip & Chen, 2005; Sun et al., 2008). In addition, another green microalga
Chlorococcum sp. was reported to produce astaxanthin in photoautotrophic
cultures as well as in heterotrophic and mixotrophic cultures on glucose (Ma,
2001).
1.2.3.5 Astaxanthin from transgenic plants
With the exception of Adonis which produces astaxanthin in its flower
petals (Seybold & Goodwin, 1959), plants are unable to synthesize astaxanthin
because they are devoid of the β-carotene ketolase that induce keto-moieties to the
4, 4’ position of β-ionone rings of β-carotene and zeaxanthin. However, the
production of astaxanthin can be achived through the functional expression of a
heterologous β-carotene ketolase gene in plants. There are numerous studies
addressing the accumulation of astaxanthin in transgenic plants including tobacco
(Mann et al., 2000; Ralley et al., 2004; Gerjets et al., 2007; Hasunuma et al.,
2008), Arabidopsis (Zhong et al., 2008), potato (Gerjets & Sandmann, 2006;
Morris et al., 2006), carrot (Jayaraj et al., 2008) and even tomato (unpublished
data from our laboratory). Although astaxanthin is produced in low amount in
transgenic plants, the accumulated astaxanthin adds nutritional value to the edible
parts of crop plants for human health and may reach commercial use in the near
future.
1.2.3.6 Natural astaxanthin versus synthetic astaxanthin
The difference between natural and synthetic astaxanthin lies in their
stereochemical orientation. Synthetic astaxanthin is a mixture of three isomeric
forms with 50% being 3R, 3′S, while natural astaxanthin from microalgae is in the
14
Chapter 1. Literature review and research aim
form of 3S, 3′S. The 3S, 3′S form of astaxanthin is reported to give a stronger
pigmentation to rainbow trout than other forms and is thus preferred as feed
additives for fish farming (Osterlie et al., 1999). Haematococcus astaxanthin is
considered to play important roles in human health and nutrition, while for other
isomers no significant biological effect has been established (Guerin et al., 2003).
Moreover, unlike synthetic astaxanthin that is present in free form, natural
astaxanthin usually exists mono-esterified or di-esterified with fatty acids (Miao et
al., 2006; Peng et al., 2008; Holtin et al., 2009). The esterified astaxanthin is
inherently more stable than the free one, thus giving a greatly longer shelf life
without being oxidized. As consumers become more and more aware of the
putative benefits of natural astaxanthin, and as the commercial production is
optimized with lowered costs, the natural astaxanthin will beat petroleum derived
synthetic astaxanthin and finally dominate the market.
1.2.4 Genetic modification for enhanced astaxanthin accumulation
1.2.4.1 Mutagenesis
Mutagenesis is a process by which the genetic information of an
organism is changed in a stable manner, either in nature or experimentally by the
use
of
chemicals
or
radiation.
Ethyl
methanesulphonate
(EMS)
and
N-methyl-N′-nitro-N-nitrosoguanidine (MNNG) are the most preferred chemical
mutagens. Ultraviolet (UV) light is also frequently used. Mutagenesis has been
widely used to modify X. dendrorhous for improved astaxanthin production (An et
al., 1989; Lewis et al., 1990; An et al., 1996; Bon et al., 1997; Ramirez et al., 2000;
Ukibe et al., 2008). X. dendrorhous mutants could produce 1-4 fold more
astaxanthin as compared with the parent ones (Lewis et al., 1990; Bon et al., 1997;
Ukibe et al., 2008). Mutagenesis has also been employed to enhance the
biosynthesis of carotenoids including astaxanthin in various algal species,
15
Chapter 1. Literature review and research aim
especially in H. pluvialis (Tjahjono et al., 1994; Zhang & Lee, 1997; Chen et al.,
2003; Ishikawa et al., 2004; Hu et al., 2008; Sandesh Kamath et al., 2008).
Commonly, after treatment of mutagens, specific inhibitors (mainly herbicide, e.g.,
fluridone, norflurazon, nicotine and diphenylamine) to carotenogenic enzymes are
used to select mutants with enhanced biosynthesis of carotenoids. Hu et al. (2008a)
reported an astaxanthin-overproducing mutant of H. pluvialis that could
accumulate astaxanthin about two times of that by the wild type (WT).
1.2.4.2 Genetic engineering of the carotenoid biosynthetic pathway
Within the past few years, genetic engineering of carotenoid metabolic
pathway has been widely employed in bacteria, algae and plants to increase the
amount of pre-existing carotenoids (Shewmaker et al., 1999; Steinbrenner &
Sandmann, 2006), to alter the carotenoid contents (Albrecht et al, 1999; Romer et
al., 2000; Gerjets et al., 2007), and to biosynthesize new products (Harker &
Hirschberg, 1997; Mann et al., 2000; Ye et al., 2000; Ralley et al., 2004; Morris et
al., 2006; Gerjets & Sandmann, 2006; Hasunuma et al., 2008; Jayaraj et al., 2008;
Zhong et al., 2008). Several prerequisites are required for the engineering of the
carotenoid metabolic pathway: (1) a clearly elucidated map for carotenoid
biosynthesis; (2) a collection of cloned genes encoding the enzymes required in
the carotenoid biosynthetic pathway; (3) knowledge about the cellular localization
of carotenoids and the enzymes involved in the carotenoid biosynthesis.
1.2.4.2.1 A clearly elucidated map for carotenoid biosynthesis
Up till now, major advances have been made in the elucidation of
carotenoid biosynthetic pathway in bacteria, algae and higher plants. It is widely
accepted that in these organisms the biosynthesis of primary carotenoids follows a
similar pathway, which generally involves three types of reactions: condensation
of two molecules of geranylgeranyl pyrophosphate (GGPP) resulting in the first
16
Chapter 1. Literature review and research aim
C40 carotene phytoene, four step-wise desaturation reactions converting colorless
phytoene to pink-colored lycopene, and cyclization of this pigment to introduce
β-ionone or ε-ionone groups at both ends, leading to the formation of β-carotene
and α-carotene respectively (Sandmann, 2002). A map for the biosynthesis of
β-carotene and α-carotene is outlined in Figure 1.5. Xanthaphylls, deriving from
oxidation of carotenes (β-carotene and α-carotene), are different in diverse types
of organisms, for example higher plants generally lack loroxanthin, astanxanthin
and canthaxanthin which are produced by certain green algae under specific
environmental stimuli (Jin et al., 2003). Basically several hydroxylation,
oxygenation and epoxidation reactions are involved in the synthesis of
xanthophylls, which is showed in Figure 1.6. Hydroxylation of the C-3 and C-3′
positions of β-carotene and α-carotene result in the formation of zeaxanthin and
lutein via β-cryptoxanthin and α-cryptoxanthin, respectively. The subsequent
epoxidation of zeaxanthin leads to the production of violaxanthin which is further
converted to neoxanthin. While in some algae, additional xanthaphyll biosynthetic
pathway is present, for example, oxygenation and hydroxylation of β-carotene
brings on the synthesis of astaxanthin, a commercially high-value ketocarotenoid.
1.2.4.2.2 A collection of cloned genes encoding the enzymes required in the
carotenoid biosynthetic pathway
In the past few years, genes encoding most of the enzymes involved in
the carotenoid biosynthesis from different prokaryotic and eukaryotic organisms
have been cloned and identified (Table 1.2; reviews see Armstrong, 1997;
Cunningham & Gantt, 1998; Jin et al, 2003). The availability of these genes has
greatly facilitated the genetic manipulation of carotenoid biosynthesis for special
purposes, such as the production of a new pigment astaxanthin in tobacco flowers
and leaves (Mann et al, 2000; Ralley et al., 2004). Since the genes from bacteria
share low level identity at nucleotide sequences with those from plants thus
eliminating the co-suppression commonly present in transgenic plants, the
17
Chapter 1. Literature review and research aim
CH2OPP
CH2OPP
DMAPP
IPP
+ IPP
CH2OPP
GPP
+ IPP
CH2OPP
FPP
+ IPP
CH2OPP
GGPP
+ GGPP
Phytoene
Phytofluene
ζ-carotene
Neurosporene
Lycopene
α-carotene
β-carotene
Figure 1.5 Carotenoid biosynthetic pathway to form β-carotene and α-carotene
(Sandmann, 2002). Abbreviations: DMAPP, dimethylallyl diphosphate; IPP,
isopentenyl diphosphate; GPP, geranyl diphosphate; FPP, farnesyl diphosphate;
GGPP, geranylgeranyl diphosphate.
18
Chapter 1. Literature review and research aim
Lycopene
α-carotene
β-carotene
Echinenone
Adonirubin
Cryptoxanthin
Lutein
Zeaxanthin
Loroxanthin
Antheraxanthin
Canthaxanthin
Adonixanthin
Astaxanthin
Violaxanthin
Neoxanthin
Figure 1.6 Schematic diagram of pathway of xanthophyll biosynthesis. In the
box is the astaxanthin biosynthetic pathway which is present in certain algae.
Two possible ways are indicated. Adapted from Jin et al. (2003).
bacteria-derived genes have been successfully employed to transform plants for
enhanced production of carotenoids. For example, overexpression of the phytoene
synthase gene from bacterium Erwinia uredovora (crtB) leaded to 2-4-fold higher
total carotenoids in fruits of transgenic tomato plants (Fraser et al., 2002);
constitutive expression of crtI (phytoene desaturase gene from E. uredovora) in
rice transformants resulted in the accumulation of α-carotene, β-carotene, lutein
and zeaxanthin in endosperms that do not normally produce carotenoid pigments
(Ye et al., 2000); simultaneous expression of crtZ and crtW from Agrobacterium
aurantiacum as polyprotein made tobacco accumulate new carotenoids including
astaxanthin, canthaxanthin and 4-ketozeaxanthin (Ralley et al., 2004). The
cyanobacterium derived crtO gene was also expressed in potato, resulting in the
19
Chapter 1. Literature review and research aim
accumulation of echinenone, 3'-hydroxyechinenone, and 4-ketozeaxanthin in
leaves, as well as astaxanthin in the tuber (Gerjets & Sandmann, 2006). In
addition, astaxanthin-producing transgenic potato and carrot plants were
successfully obtained through the expression of a β-carotene oxygenase gene from
H. pluvialis, making these economical crops more nutritionally attractive (Morris
et al., 2006; Gerjets et al., 2008). All the mentioned transgenic plants are
generated through Agrobacterium-mediated gene transfer method, yielding low
amounts of astaxanthin due to the presence of large quantities of astaxanthin
intermediates. The employment of plastid transformation strategy overcame this
problem and made transgenic tobacco plants accumulate high levels of
astaxanthin (more than 70% of total carotenoids) in leaves (Hasunuma et al.,
2008). In addition to higher plants, the green alga H. pluvialis has also been
successfully genetically modified with accelerated astaxanthin production
(Steinbrenner & Sandmann, 2006).
20
Chapter 1. Literature review and research aim
Table 1.2 Genes involved in the carotenoid biosynthesis
Gene
Enzymatic function
Species
crtE/GGPPS
GGPP synthase
Erwinia uredovora (Misawa et al., 1990); Pepper (Kuntz et al., 1992); Arabidopsis (Lange & Ghassemian, 2003)
crtB/PSY
Phytoene synthase
E. ruedovora (Misawa et al., 1990); Arabidopsis (Lange & Ghassemian, 2003); Tomato (Fraser et al., 2000); H.
pluvialis (Steinbrenner & Linden, 2001)
crtI/crtP/PDS
Phytoene desaturase
E. ruedovora (Misawa et al., 1990); Synechococcus (Chamovitz et al., 1991); Chlamydomonas reinhardtii
(McCarthy et al., 2004), H. pluvialis (Harker & Hirchberg, 1997); C. zofingiensis (Huang et al., 2008); Dunaliella
salina (Zhu et al., 2005); Arabidopsis (Lange & Ghassemian, 2003); Tomato (Pecker et al., 1992)
crtY/crtL/LCY
Lycopene cyclase
E. ruedovora (Misawa et al., 1990); Synechococcus (Cunningham et al., 1994); C. reinhardtii (Lohr et al., 2005); H.
pluvialis (Steinbrenner & Linden, 2003); D. salina (Ramos et al., 2008); Arabidopsis (Cunningham et al., 1996;
Cunningham & Gantt, 2001); Tomato (Rogen et al., 1999 & 2000)
crtZ/CHYb
crtW/crtO/BKT
β-carotene
E. ruedovora (Misawa et al., 1990); C. reinhardtii (Lohr et al., 2005); H. pluvialis (Linden, 1999); C. zofingiensis
hydroxylase
(Li et al., 2008); Arabidopsis (Sun et al., 1996); Tomato (Hirchberg, 1998)
β-carotene oxygenase
A. aurantiacum (Misawa et al., 1995); Synechocystis (Fernandez-Gonzalez et al., 1997); C. reinhardtii (unpublished
data); H. pluvialis (Kajiwara et al., 1995; Huang et al., 2006a); C. zofingiensis (Huang et al., 2006b);
ZEP
Zeaxanthin epoxidase
Arabidopsis (Lange & Ghassemian, 2003); Pepper (Bouvier et al., 1996)
NXS
Neoxanthin synthase
Potato (Al-Babili et al., 2000); Tomato (Bouvier et al., 2000)
21
Chapter 1. Literature review and research aim
1.2.4.2.3 Cellular localization of carotenoids and the enzymes involved in the
carotenoid biosynthesis
Carotenoids, relatively hydrophobic molecules, are typically associated
with membranes and/or non-covalently bound to specific proteins (Armstrong,
1997). Considerable progress has been made in the cellular localization of
carotenoids and the enzymes that are required for the synthesis of these
carotenoids in the recent years.
In higher plants, carotenoids are localized in the plastid where the
carotenoid biosynthetic enzymes are present as membrane associated or bound
proteins. Phytoene synthase, responsible for the formation of the first carotenoid
phytoene from GGPP through a two-step reaction, was purified from pepper
chromoplast stroma in soluble form (Dogbo et al., 1988). The solubilization of
this enzyme, however, required treatment with high ionic strength buffer or mild
non-ionic detergent (Fraser et al., 2000), suggesting that phytoene synthase
acitivity is membrane-associated. As reported by Al-Babili et al. (1996), both
soluble and membrane-bound phytoene desaturase was detected by western blot
analysis in the isolated chromoplasts: the soluble one in stroma showed no
enzymatical activity and got activated once bound to membrane. This is further
supported by in vitro protein import assays in pea chloroplasts (Bonk et al., 1997).
In this research, the suborganellar localization of three other carotenogenic
enzymes was also investigated. Similar to phytoene desaturase, after imported into
chloroplasts, lycopene cyclase remained soluble in the stroma, forming a
high-molecular-mass complex with chloroplast 60-kDa chaperonin (Cpn60);
while phytoene synthase transiently formed a complex with Cpn60 and then
associated rapidly to thylakoid membranes upon import. As expected, the
geranylgeranyl diphosphate synthase stayed in soluble and free form in the stroma.
The in vitro import assay of carotenogenic enzymes from citrus was also carried
out in a very recent report which demonstrated the subcellular localization and
transit peptide cleavage of these proteins (Inoue et al., 2006). Consistent with the
22
Chapter 1. Literature review and research aim
results stated above, phytoene synthase was found to be peripherally associated
with the membrane and phytoene desaturase mainly stayed in the stroma.
Lycopene cyclase, however, was targeted both to the soluble and to the membrane
compartments, which slightly differed from the previous report of Bonk et al.
(1997). In addition, carotenoid β-ring hydroxylase was exclusively inserted into
the chloroplast internal membranes, the first demonstration of plastid cellular
localization of this enzyme in plants (Inoue et al., 2006).
Similar to plants, in algae all primary carotenoids and some xanthaphylls
such as lutein and zeaxanthin are synthesized within plastids. Additional
carotenoids, e.g. canthaxanthin and astaxanthin which are generally not present in
plants, however, are found to accumulate in lipid vesicles outside the plastid in H.
pluvialis (Boussiba, 2000). The cellular compartmentaton of H. pluvialis phytoene
desaturase was investigated through using immunogold labeling of ultra sections
and western blot analysis of cell fractions which revealed that this active enzyme
was localized exclusively in the chloroplast, or more specifically speaking in close
contact to thylakoids (Grunewald et al., 2000), ruling out the possible biosynthesis
of lycopene, the direct β-carotene precursor, outside chloroplast. β-carotene
oxygenase, the enzyme catalyzing the introduction of keto groups at position C-4
of the β-ionone ring of β-carotene and zeaxanthin, localized both in the
chloroplast and cytoplasmic membrane-derived lipid vesicles but only functions
in the latter compartment (Grunewald et al., 2001). Based on the information,
there might be a transport process of intermediate carotenoids over compartment
borders from chloroplast as the site of synthesis to the lipid vesicles located in the
cytoplasm as the site of accumulation. To determine what carotenoids may serve
as the intermediates, further research was carried out by Grunewald & Hagen
(2001). Treated with diphenylamine (DPA, a specific inhibitor to β-carotene
oxgenation), the majority of β-carotene was found to accumulate in lipid vesicles;
while
few
lycopene
was
observed
when
treated
with
2-(4-chlorophenylthio)-triethylamine (CPTA, specifically inhibiting lycopene
cyclization), suggesting that β-carotene is the intermediate exported from the
23
Chapter 1. Literature review and research aim
chloroplast during accumulation of secondary carotenoids in H. pluvialis. It was
hypothesized that the nuclear-encoded β-carotene oxgenase could be transported
first into the chloroplast and might then be exported out together with the
substrate β-carotene into cytoplasm where it was sequestered by cytoplasmic
membrane-derived lipid vesicles (Grunewald et al., 2001). Since the β-carotene
hydroxylase owns a dual function in H. pluvialis being responsible for the
formation of zeaxanthin from β-carotene and of astaxanthin from canthaxanthin, it
may locate and function in both the chloroplasts and the lipid vesicles.
Generally, enzymes should locate specific cellular compartments where
substrates and/or cofactors are present to perform their function. Thus the
information about cellular compartmentation described above is of great
importance to genetic manipulation of carotenoid biosynthesis. Since carotenoids
are produced and accumulated in plastids, the heterogenous genes which are
employed to genetically engineer carotenoid biosynthesis in certain organism
should be fused to a short specific protein sequence that ensures the
plastid-targeting of enzymes encoded by these genes. Overexpression of a
bacterial phytoene synthase (crtB) gene resulted in 50-fold higher carotenoid
accumulation in canola mature seed (Shewmaker et al., 1999). crtI, another
bacteria gene encoding a enzyme responsible for a four-step desaturation reaction,
has been widely used in plants to elevate carotenoid level or produce new
pigments in carotenoid-free tissues (Romer et al., 2000; Ye et al., 2000). In these
genetic manipulations or others (Gerjets & Sandmann, 2006; Morris et al., 2006),
the pea ribulose bisphosphate carboxylase small subunit transit sequence was used
to target the proteins of interest into plastids. Tomato phytoene synthase-1 transit
sequence was also employed for the charomoplast targeting of CRTB protein to
alter the carotenoid content (Fraser et al., 2002). What’s more, the crtO gene from
green algae H. pluvialis was transferred to tobacco and the CRTO polypeptide was
targeted by the transit peptide of PDS from tomato into chromoplasts where this
enzyme catalyzes the oxygenation reaction, leading to the accumulation of
astaxanthin and other ketocarotenoids in nectary tissues and thus the color of the
24
Chapter 1. Literature review and research aim
nectary changes from yellow to red (Mann et al., 2000).
1.3 Biodiesel
1.3.1 Introduction
Fossil-based fuels including oil, coal and gas play a pivotal role in
modern world energy market. These fossil fuels, according to world energy
outlook 2007, will remain the major sources of energy and are expected to meet
about 84% of energy demand in 2030. However, fossil fuels are non-renewable
and will be finally diminished. It has been recently estimated that the global oil,
coat and gas lasts only around for 35, 100 and 37 years respectively, based on a
modified Klass model (Shafiee & Topal, 2009). In order to sustain a stable energy
supply in the future, it is necessary to develop other sources of energy, e.g.,
renewable energy. Renewable energy is derived from natural processes that are
replenished constantly, including hydropower, wind power, solar energy,
geothermal energy, biodiesel, etc. An estimated $120 billion was invested in
renewable energy worldwide in 2008, around 2 times of the 2006 investment
(Figure 1.7).
25
Chapter 1. Literature review and research aim
Figure 1.7 Global investment in renewable energy, 2004-2008. Source:
REN21, 2009.
It is well known that transport is almost totally dependent on
petroleum-based fuels, which will be depleted within no more than 40 years. An
alternative fuel to petrodiesel must be technically feasible, easily available,
economically competitive, and environmentally acceptable (Demirbas, 2008).
Biodiesel is such a candidate fuel for powering the transport vehicles. Biodiesel
refers to a biomass-based diesel fuel consisting of long-chain alkyl (methyl,
propyl or ethyl) esters. In addition to being comparable to petrodiesel in most
technical aspects, biodiesel has several following distinct advantages over
petrodiesel (Knothe, 2005):
(1)
derivation from renewable domestic resources, thus reducing dependence
on and preserving petroleum;
(2)
biodegradability;
(3)
reduction of most exhaust emissions (except nitrogen oxides, NOx).
26
Chapter 1. Literature review and research aim
(4)
higher flash point, leading to safer handling and storage;
(5)
excellent lubricity.
Like petrodiesel, biodiesel operates in compression ignition engines.
Biodiesel is miscible with petrodiesel in all ratios. Currently, the blends of
biodiesel and petrodiesel instead of net biodiesel have been widely used in many
countries and no engine modification is required (Singhania et al., 2008). These
blends of biodiesel with petrodiesel are usually denoted by acronyms, for example
B20 which indicates a blend of 20% biodiesel with petrodiesel (Knothe, 2005).
The global markets for biodiesel are entering a period of rapid and transitional
growth. In the year 2007, there were only 20 oil producing nations supplying the
needs of over 200 nations; by the year 2010, more than 200 nations will become
biodiesel producing nations and suppliers (Thurmond, 2008). Global biodiesel
production has massively increased to 10.8 million tons per year over the last
eight years (Figure 1.8). Much of the growth is happening in just three countries:
the United States, Brazil and Germany, which together account for over half of
biodiesel (Checkbiotech, 2009). The International Energy Agency’s report
suggests that world production of biodiesel could top 20 million tons per year by
2012 if the recent trends continue.
27
Chapter 1. Literature review and research aim
Figure 1.8 Global biodiesel production, 2001-2008. Adapted from Thurmond
(2008).
1.3.2 Biodiesel production through transesterification
Vegetable oils have long been recognized as fuels for diesel engines
(Shay, 1993). However, vegetable oils are too viscous for most currently used
diesel engines. The widely employed method in industry to reduce oil viscosity is
called transesterification, resulting in biodiesel production (Demirbas, 2005).
Transesterification is a chemical conversion process involving reacting
triglycerides of vegetable oils or animal fats catalytically with a short-chain
alcohol (typically methanol or ethanol) to form fatty acid esters and glycerol
(Figure 1.9). This reaction occurs stepwise with the first conversion of
triglycerides to diglycerides and then to monoglycerides and finally to glycerol.
The complete transesterification of 1 mol of triglycerides requires 3 mol of
alcohol, producing 1 mol of glycerol and 3 mol of fatty esters. Considering that
the reaction is reversible, large excess of alcohol is used in industrial processes to
28
Chapter 1. Literature review and research aim
ensure the direction of fatty acid esters (Fukuda et al., 2001). Methanol is the
preferred alcohol for industrial use because of its low cost, although other
alcohols like ethanol, propanol and butanol are also commonly used (Ataya et al.,
2007).
Figure 1.9 Transesterification of oil to biodiesel. R1-3 indicate hydrocarbon
groups.
In addition to heat, a catalyst is needed to facilitate the transesterification.
The transesterification of triglycerides can be catalyzed by acids (Ataya et al.,
2007; Guan et al., 2009; Miao et al., 2009; Furukawa et al., 2010), alkalis (Leung
& Guo, 2006; Qian et al., 2008) or enzymes (Torres et al., 2005; Shah & Gupta,
2007; Su et al., 2007; Raita et al., 2010). Acid transesterification is considered
suitable for the conversion of feedstocks with high free fatty acids but its reaction
rate is low (Gerpen, 2005). In contrast, alkali-catalyzed transesterification has a
much higher reaction rate, approximately 4000 times faster than the
acid-catalyzed one (Fukuda et al., 2001). In this context, alkalis (sodium
hydroxide and potassium hydroxide) are preferred as catalysts for industrial
production of biodiesel. The production of biodiesel by alkali process is shown in
Figure 1.10. The use of lipases as transesterification catalysts has also attracted
much attention as it produces high purity product and enables easy separation
from the byproduct glycerol (Ranganathan et al., 2008). However, the cost of
enzyme is still relatively high and remains a barrier for its industrial
implementation. In addition, it has been proposed that biodiesel can be prepared
29
Chapter 1. Literature review and research aim
from oil via transesterification with supercritical methanol (Saka & Kusdiana,
2001; Demirbas, 2002). Table 1.3 shows the comparison of various methanolic
transesterification methods, in terms of reaction temperature and time; while Table
1.4 summarizes the advantages and disadvantages of various transesterification
methods.
Oil
Transesterification
Alkali
+
Methanol
Separation
Evaporation of methanol
Upper
phase
Waste
water-alkali
Washing
Biodiesel
Lower
phase
Evaporation of methanol
Saponified
products
Purification
Glycerol
Figure 1.10 Production process of biodiesel using alkali as the catalyst
(Ranganathan et al., 2008).
30
Chapter 1. Literature review and research aim
Table 1.3 Comparison of various methanolic transesterification methods
(Demirbas, 2008)
Method
Reaction temperature (K) Reaction time (min)
Acid or alkali catalytic process
303-338
60-360
Boron trifluoride–methanol
360-390
20-50
Sodium methoxide–catalyzed
293-298
4-6
Non-catalytic supercritical methanol
523-573
6-12
Catalytic supercritical methanol
523-573
0.5-1.5
Table 1.4 Advantages and disadvantages of various transesterification methods
(Huang et al., 2010)
Type of
Advantages
Disadvantages
Chemical
Reaction condition can be well
Reaction temperature is relative high
catalysis
controlled
and the process is complex
Large scale production
The later disposal process is complex
The cost of the production
The process need much energy
process is cheap
Need a installation for methanol recycle
The methanol produced in the
The waste water pollutes the
process can be recycled
environment
transesterification
High conversion of the
production
Enzymatic
Moderate reaction condition
Limitation of enzyme in the conversion
catalysis
The small amount of methanol
of short chain of fatty acids
required in the reaction
Chemicals exist in the process of
Have no pollution to natural
production are poisonous to enzyme
environment
Supercritical
Easy to be controlled
High temperature and pressure in the
fluid techniques
It is safe and fast
reaction condition leads to high
Friendly to environment
production cost and wastes energy
31
Chapter 1. Literature review and research aim
1.3.3 Biodiesel feedstocks
Biodiesel can be produced from a variety of feedstocks, including plant
oils, animal fats and waste oils as well as algae (Demiras, 2008). Each feedstock
has its advantages and disadvantages in terms of oil content, fatty acid
composition, biomass yield and geographic distribution. Depending on the origin
and quality of feedstocks, changes may be required for the production process of
biodiesel.
1.3.3.1 Plant oils
The use of plant oils as biodiesel feedstocks has been long recognized
and well documented in numerous studies (Saka & Kusdiana, 2001; de Oliveira et
al., 2005; Hill et al., 2006; Rashid & Anwar, 2008; Abdullah et al., 2009; Hawash
et al., 2009; Graef et al., 2009; Sahoo & Das, 2009; Patil & Deng, 2009; Jain &
Sharma, 2009; Nakpong & Wootthikanokkhan, 2010). These feedstocks include
the oils from soybean, rapeseed, palm, canola, peanut, cottonseed, sunflower and
safflower. Based on the geographic distribution, soybean is the primary source for
biodiesel in USA, palm oil is used as a significant biodiesel feedstock in Malaysia
and Indonesia, and rapeseed is the most common base oil used in Europe for
biodiesel production (Demiras, 2008). The vast majority of these plants are also
used for food and feed production, which means that possible food versus fuel
conflicts are present. Thus, the use of these plant oils as feedstocks for biodiesel
seems insignificant for the developing countries which are importers of edible oils
(Meher et al., 2008). In addition to these edible oils, various non-edible,
tree-borne oils from jatropha, karanja, jojoba and neem are the potential biodiesel
feedstocks (Meher et al., 2008; Sahoo & Das, 2009; Jain & Sharma, 2009).
Jatropha and karanja are two oilseed plants that are not widely exploited due to
the presence of toxic components in the oils. In India, they are popularly used as
32
Chapter 1. Literature review and research aim
biodiesel feedstocks. The physical properties of biodiesel from various plant
feedstocks are listed in Table 1.5.
Table 1.5 Physical properties of biodiesel from varous feestocks. Sources: Knothe,
2005; Meher et al., 2008
Feedstock; ester
CN
HG (kj/kg)
KV (40 ℃;
mm2/s)
CP (℃)
PP (℃)
FP (℃)
No. 2 Diesel
47.0
45343
2.7
-15.0
-33.0
52
Soybean; methyl
49.6
37372
4.18
-1.1
-3.9
190.6
Palm; ethyl
56.2
39070
4.50 (37.8 ℃)
8
6
—
Rapeseed; methyl
47.9
39870
4.76 (37.8 ℃)
-3
-9
166
Coconut; ethyl
67.4
38158
3.08
5
-3
190
Corn; methyl
65
38480
4.52
-3.4
-3
111
Sunflower; methyl
58
38472
4.39
1.5
3
110
Safflower; ehtyl
62.2
39872
4.31
-6
-6
178
Cottonseed; methyl
51.2
—
6.8 (21 ℃)
—
-4
110
Olive; methyl
61
37287
4.70
-2
-3
>110
Mustard; ethyl
54.9
40679
5.66
1
-15
183
Jatropha; methyl
51
—
4.84
—
—
191
Karanja; methyl
56
—
4.77
—
—
174
Tallow; methyl
61.8
37531
4.99
15.6
12.8
187.8
Grease; ethyl
—
—
6.20
5
-1
—
Used frying oil;
methyl
59
37337
4.50
1
-3
>110
Waste olive oil;
methyl
58.7
—
5.29
-2
-6
—
CN, cetane number; HG, gross heat of combustion; KV, kenematic viscosity; CP,
cloud point; PP, pour point; FP, flash point.
33
Chapter 1. Literature review and research aim
1.3.3.2 Animal fats and waste oils
In addition to the plant oils, animal fats and waste oils are the potential
sources for commercial biodiesel production (Thompson et al., 2010). Among
these feedstocks, tallow, lard, yellow grease and waste cooking oils have received
most interest (Canakci, 2007; Phan & Phan, 2008; Banerjee et al., 2009; da Cunha
et al., 2009; Dias et al., 2009; Diaz-Felix et al., 2009; Oner & Altun, 2009).
However, animal fats and waste oils usually contain large amounts of free fatty
acids, which can be as high as 41.8% (Canakci, 2007). Free fatty acids cannot be
directly converted to biodiesel in alkali-catalyzed transesterificatoin but react with
alkali to form soaps that inhibit the separation of biodiesel from glycerin and wash
water fraction (Huang et al., 2010). A two-step process was developed for these
high fatty acid feedstocks: acid-catalyzed pretreatment and alkali-catalyzed
transesterificaton (Canacki & Van Gerpen, 2003). Because animal fats and waste
oils have relatively high level of saturation (Canakci, 2007), the biodiesel from
these sources exhibits poor cold flow properties (Table 1.5).
1.3.3.3 Algal oils
Algae represent a wide variety of aquatic photosynthetic organisms with
the potential of producing high biomass and accumulating high level of oil. The
production of biodiesel from algal oil has long been recognized and been
evaluated in response to the United States Department of Energy for research in
alternative
renewable
energy
(Sheehan
et
al.,
1998).
Currently,
the
commercialization of algae-derived biodiesel is still in its infancy stage. An
unprecedentedly increasing interest was received, in terms of algal strain selection,
biomass production, lipid yielding, transesterification technologies, fuel properties
and engine tests (Miao & Wu, 2006; Xu et al., 2006; Li et al., 2007; Ross et al.,
2008; Demirbas, 2009; Greenwell et al., 2009; Pruvost et al., 2009; Rodolfi et al.,
2009; Yoo et al., 2009; Brennan & Owende, 2010; Xiong et al., 2010). Algae have
34
Chapter 1. Literature review and research aim
been considered as the only feedstock of biodiesel that has the potential to
displace fossil diesel (Chisti, 2007).
1.3.4 Potential and prospect of microalgal biodiesel
Microalgae can grow autotrophically using CO2 from air and light
through photosynthetic reactions and/or heterotrophically utilizing organic
compounds as carbon and energy sources. Compared with higher plants,
microalgae exhibit higher photosynthetic efficiency and growth rate (Chisti, 2007).
Commonly, microalgae are cultivated photoautotrophically in open ponds or
enclosed bioreactors for biomass production (Molina et al., 1999; Carvalho et al.,
2006; Spolaore et al., 2006). A comparison of open and enclosed culture systems
for microalgae is shown in Table 1.6. However, it is not easy to obtain high cell
densities on a large scale in autotrophic culture systems because of light limitation
or photoinhibition when exposed to high light (Chen, 1996; Harker et al., 1996;
Wen et al., 2003). In contrast, heterotrophic culture provides several advantages
over autotrophic culture, including good control of cultivation parameters,
elimination of light requirement, high cell biomass obtained and low-cost for algal
biomass harvest (Chen, 1996). Heterotrophic algal biomiass prodcution has been
documented in many studies (Hata et al., 2001; Wen & Chen, 2001; Shi & Chen,
2002; Ip & Chen, 2005; Sun et al., 2008; Johnson & Wen, 2009; Liang et al., 2009).
Chlorella protothecoides, a well-studied green microalga, is considered to be
feasible for biodiesel feedstocks under heterotrophic culture conditions (Miao &
Wu, 2004, 2006; Xu et al., 2006; Li et al., 2007; Gao et al., 2010; Xiong et al.,
2010).
35
Chapter 1. Literature review and research aim
Table 1.6 A comparison of open and closed culture systems for microalgae.
Source: Mata et al., 2010
Culture systems
Open ponds
Enclosed bioreactors
Contamination control
Difficult
Easy
Contamination risk
High
Reduced
Sterility
None
Achievable
Process control
Difficult
Easy
Species control
Difficult
Easy
Mixing
Very poor
Uniform
Operation regime
Batch or
Batch or
semi-continuous
semi-continuous
Area/volume ration
Low
High
Algal cell density
Low
High
Investment
Low
Hight
Operation cost
Low
High
Light utilization efficiency
Poor
High
Temperature control
difficult
More uniform
temperature
Productivity
Low
High
Hydrodynamic stress on
algae
Very low
Low-high
Evaporation of growth
medium
High
Low
Gas transfer control
Low
High
O2 inhibition
< bioreactors
Great problem
Scale-up
Difficult
Difficult
In addition to biomass, lipid content is another important factor to assess
the potential of microalgae for biodiesel production. Over the past few decades,
36
Chapter 1. Literature review and research aim
thousands of algae and cyanobacterial species have been screened for high lipid
production, and numerous oleaginous species have been isolated and
characterized. The lipid contents of these oleaginous algae are species- and/or
strains-dependent and may vary greatly, as shown in Figure 1.11. Under optimal
growth conditions, algae commonly synthesize a low content of lipids (i.e.,
averagely 25.5% of dry weight for green algae, Figure 1.11A), with membrane
lipids (e.g., phospholipids and glycolidips) being the main components; whereas
under unfavorable environmental or stress conditions, a great increase in total
lipids was observed (e.g., averagely 45.7% of dry weight for green algae, Figure
1.11A) with neutral lipids in particular triacylglycerols (TAGs) being dominant
(Hu, 2004). TAGs are considered to be superior to phospholipids or glycolipids
for biodiesel feedstocks because of their higher percentage of fatty acids and lack
of phosphate (Pruvost et al., 2009). Unlike higher plants in which individual
classes of lipids may be synthesized and localized in a specific cell, tissue or
organ, algae produce these different lipids in a single cell (Hu et al., 2008b). The
synthesized TAGs are deposited in lipid bodies located in cytoplasm of algal cells
(Rabbani et al., 1998; Damiani et al., 2010).
37
Chapter 1. Literature review and research aim
Figure 1.11 Cellular lipid content in various classes of microalgae and
cyanobacteria under normal growth and stress conditions. (A) Green
microalgae; (B) diatoms; (C) oleaginous algae from other eukaryotic algal
taxa; (D) cyanobacteria. Open circles: cellular lipid contents obtained under
normal growth or nitrogen-replete conditions; closed circles: cellular lipid
contents obtained under nitrogen-depleted or other stress conditions. The
differences in cellular lipid content between cultures under normal growth and
stress growth conditions were statistically significant for all three groups (A, B
and C) of algae examined using Duncan’s multiple range test with the ANOVA
procedure. Source: Hu et al., 2008b.
The important properties of biodiesel such as cetane number, viscosity,
cold flow, oxidative stability, are largely determined by the composition and
structure of fatty acid esters which in turn are determined by the characteristics of
fatty acids of biodiesel feedstocks, for exmaple carbon chain length and
unsaturation degree (Knothe, 2005b). Fatty acids are either in saturated or
unsaturated form, and the unsaturated fatty acids may vary in the number and
position of double bones on the acyl chain. Based on the number of double bones,
unsaturated fatty acids are clarified into monounsaturated fatty acids (MUFAs)
and polyunsaturated fatty acids (PUFAs). The fatty acid profile of a great many
algal species has been investigated and is shown in Table 1.7. The synthesized
fatty acids in algae are commonly in medium length, ranging from 16 to 18
carbons, despite the great variation in fatty acid composition. Specifically, the
major fatty acids are C16:0, C18:1 and C18:2 or C18:3 in green algae, C16:0 and
C16:1 in diatoms and C16:0, C16:1, C18:1 and C18:2 in cyanobacteria. It is
worthy to note that these data are obtained from algal species under specific
conditions and vary greatly when algal cells are exposed to different
environmental or nutritional conditions such as temperature, pH, or nitrogen
concentration (Wada & Murata, 1990; Chen & Johns, 1991; Tatsuzawa &
38
Chapter 1. Literature review and research aim
Takizawa, 1996; Khozin-Goldberg et al., 2002; Liu et al., 2005). Generally,
saturated fatty esters possess high cetane number and superior oxidative stability;
whereas unsaturated, especially polyunsaturated, fatty esters have improved
low-temperature properties (Knothe, 2008). In this regard, it is suggested that the
modification of fatty esters, for example the enhanced proportion of oleic acid
(C18:1) ester, can provide a compromise solution between oxidative stability and
low-temperature properties and therefore promote the quality of biodiesel (Knothe,
2008, 2009). Thus, microalgae with high oleic acid are suitable for biodiesel
production.
Currently the commercial production of biodiesel is mainly from plant
oils and animal fats. However, the plant oil derived biodiesel cannot realistically
meet the demand of transport fuels because large arable lands are required for
cultivation of oil plants, as demonstrated in Table 1.8. Based on the oil yield of
different plants, the cropping area needed is calculated and expressed as a
percentage of the total U.S. cropping area. If soybean, the popular oil crop in
United States is used for biodiesel production to meet the existing transport fuel
need, 5.2 times of U.S. cropland will need to be employed. Even the high-yielding
oil plant palm is planted as the biodiesel feedstock, more than 50% of current U.S.
arable lands have to be occupied. The requirement of huge arable lands and the
resulted conflicts between food and oil make the biodiesel from plant oils
unrealistic to completely replace the petroleum derived diesel in the foreseeable
future. It is another case, however, if microalgae are used to produce biodiesel. As
compared with the conventional oil plants, microalgae possess significant
advantages in biomass production and oil yield and therefore the biodiesel
productivity. In terms of land use, microalgae need much less than oil plants, thus
eliminating the competition with food for arable lands (Table 1.8). Based on the
above mentioned information, microalgae appear to be the only source of
biodiesel that has the potential to replace fossil derived diesel.
39
Chapter 1. Literature review and research aim
Table 1.7 Fatty acid composition of some algal species (% of total fatty acids)
Fatty C14:0 C16:0 C16:1 C16:2 C16:3 C17:0 C18:0 C18:1 C18:2 C18:3 C18:4 C18:5 C20:0 C20:1 C20:4 C20:5 C22:5 C22:6
acids
Chlorophyta
C.p.
1.31
12.94
0.89
2.76
60.84 17.28
0.35
0.42
C.v.
18.0
5.0
12.0
2.1
9.2
43.0
10.0
C.s.
25.4
3.1
10.7
4.1
1.4
12.4
34.4
7.1
H.p.a 1.25
22.49 0.64
0.19
3.15
19.36 26.9
17.04
0.2
0.13
0.89
0.57
C.r.
4.0
36.1
1.8
4.4
13.3
17.8
20.5
2.1
C.sp
2.0
39.5
2.1
1.5
30.2
2.7
22.1
P.i.
9.1
0.7
0.6
2.1
15.1
9.3
1.6
1.2
58.9
Bacillariophyta
N.l.
16.9
28.5
23.9
0.7
5.1
3.4
4.1
5.0
11.7
Ch.sp 23.6
9.2
36.5
6.9
2.6
2.0
3.0
1.4
0.6
4.1
8.0
1.0
B.a.
32.0
5.0
27.0
2.0
8.0
26.0
Cyanophyta
Syn.
52.0
3.0
1.0
3.0
9.0
29.0
3.0
N.f.
0.65
21.27 14.91
6.2
22.59 15.03 19.35
N.c.b
23.5
22.5
5.6
21.1
14.1
Others
G.c.
22.0
4.4
4.0
6.6
3.9
5.5
39.2
13.3
N.sp
6.9
19.9
27.4
1.7
3.5
0.7
4.2
34.9
S.sp
3.2
9.4
0.7
0.5
1.5
1.5
5.4
10.6
43.1
1.8
0.1
18.8
Abbreviation of algal species: C.p., Chlorella protothecoides (Li et al., 2007); C.v., Chlorella vulgaris (Harris et al., 1965); C.s., Chlorella sorokiniana (Chen & Johns,
1991); H.p., Haematococcus pluvialis (Damiani et al., 2010); C.r. and C.sp, Chlamydomonas reinhardtii and Chlamydomonas sp. (Tatsuzawa and Takizawa, 1996);
P.i., Parietochloris incise (Khozin-Goldberg et al., 2002); N.l., Nitzschia laevis (Chen, 2007); Ch.sp, Chaetoceros sp. (Renaud et al., 2002); B.a., Biddulphia aurica
(Orcuut & Patterson, 1975); Syn., Synechocystis PCC6803 (Wada & Murata, 1990); N.f., Nostoc flagelliforme (Liu et al., 2005); N.c., Nostoc commune (Pushparaj et
al., 2008); G.c., Glossomastrix chrysoplasta (Kawachi et al., 2002); N.sp, Nannochloropsis sp (Sukenik, 1999); S.sp, Scrippsiella sp (Mansour et al., 1999). a from
neutral lipids, some short-chain fatty acids in low content are not listed. b unknown fatty acids are not listed.
40
Chapter 1. Literature review and research aim
Table 1.8 Comparation of microaglae with other biodiesel feedstocks. Source:
Chisti, 2007; Mata et al., 2010
Plant source
Oil yeild
Land area needed
Percentage of existing
(L/ha year)
(M ha) a
US cropping areaa
Corn
172
3480
1912
Hemp
363
1650
906
Soybean
636
940
516
Jatropha
741
807
443
Camelina
915
650
357
Canola
974
610
335
Sunflower
1070
560
307
Castor
1307
450
247
Palm oil
5366
110
60.4
Microalgae b
58,700
9.0
4.9
Microalgae c
97,800
5.4
3.0
Microalgae d
136,900
3.9
2.1
a
for meeting all transport fuel needs of the United States. b 30% oil by dry weight.
c
50% oil by dry weight. c 70% oil by dry weight.
However, the cost of microalgal biodiesel still remains high, which is
mainly attributed to the high cost of microalgal oil. The microalgal oil is estimated
to cost $2.8/L, much higher than that of crude fossil oil and plant oil (Chisti,
2007). To make algae-derived biodiesel cost-competitive with petroleum,
microalgal oil should be at a price like this: Calgal oil = 6.9 × 10-3 Cpetroleum, where
Calgal oil ($ per liter) is the price of microalgal oil and Cpetroleum ($ per barrel) is the
price of crude fossil oil (Chisti, 2007). For example, if the price of crude oil
approaches to $100/barrel, it appears to be economical for microalgal oil to
replace crude petroleum at the cost of $0.69/L. Production cost of microalgal
biodiesel may be brought down substantially by improving capabilities of
41
Chapter 1. Literature review and research aim
microalgae through metabolic and genetic engineering or by using a biorefinery
based production strategy.
1.3.5 Lipid metabolism in microalgae
Although lipid metabolism, in particular the biosynthesis of fatty acids and
TAGs, is poorly understood in algae, it is generally considered that the basic
pathways for fatty acid and TAGs biosynthesis are similar to those demonstrated
in higher plants.
1.3.5.1 Fatty acid/lipid biosynthesis
Similar to plants, microalgae synthesize fatty acids in the chloroplast
using a single set of enzymes. A simplified schedule for fatty acid biosynthesis is
shown in Figure 1.12. The formation of malonyl CoA from acetyl CoA is
generally regarded as the first reaction of the fatty acid biosynthetic pathway,
which is catalyzed by acetyl CoA carboxylase (ACCase). The malonyl group of
malonyl CoA is transferred to a protein co-factor on the acyl carrier protein (ACP),
resulting in the formation of malonyl ACP that involves in subsequent
condensation and elongation reactions. The first condensation reaction is
catalyzed by 3-ketoayl ACP synthase III (KAS III), forming a four-carbon product.
KAS I and KAS II catalyze the subsequent condensations and finally the saturated
16:0- and 18:0-ACP are produced. To produce unsaturated fatty acids, the double
bonds are introduced by the enzyme stearoyl ACP desaturase (SAD). Unlike
plants, some microalgae produce long-chain acyl ACPs (C20-C22) that derive from
the further elongation and/or desaturation of C18 (Figure 1.13). The final fatty acid
composition of individual algae is determined by the activities of enzymes that
use these acyl ACPs as substrates at the termination phase of fatty acid
biosynthesis. These fatty acids are then used as the precursors for the synthesis of
42
Chapter 1. Literature review and research aim
cellular membranes and neutral storage lipids like TAGs.
Acetyl-CoA
ACCase
KAS III
Malonyl-CoA
C4:0-ACP
Chloroplast
KAS I
C16:0-ACP
KAS II
C18:0-ACP
SAD
C18:1-ACP
C16-C18-CoA
Figure 1.12 A simplified schedule for fatty acid biosynthesis (Ohlrogge &
Jaworski, 1997).
It has been proposed that the biosynthesis of TAG occurs in cytosol via
the direct glycerol pathway (Figure 1.14). Generally, acyl-CoAs sequentially react
with the hydroxyl groups in glycerol-3-phosphate to form phosphatidic acid.
These two reactions are catalyzed by glycerol-3-phospate acyl transferase and
lysophosphatidic acid acyl transferase respectively. Dephosphorylation of
phosphatidic acid results in the release of diacylglycerol which accepts a third
acyl from CoA to form TAG. This final step is catalyzed by diacylglycerol
acyltransferase, an enzymatic reaction that is unique to TAG synthesis. In addition,
an alternative pathway that is independent of acyl-CoA may also be present in
43
Chapter 1. Literature review and research aim
microalgae for TAG biosynthesis (Dahlqvist et al., 2000). This pathway employs
phospholipids as acyl donors and diacylglycerols as the acceptors and might be
activated when algal cells are exposed to stress conditions because algae usually
undergo rapid degradation of the photosynthetic membranes and concurrent
accumulation of cytosolic TAG-enriched lipid bodies (Hu et al., 2008b).
C18:0
△ 9 desaturase
C18:1
△ 12 desaturase
C18:2 (n-6) △ 15 desaturase
△ 9, 12
C18:3 (n-3)
△ 9, 12, 15
C20:2 (n-6)
△ 11, 14
C18:3 (n-6) △ 15 desaturase
△ 6, 9, 12
C18:4 (n-3)
△ 6, 9, 12, 15
△ 8 desaturase
C20:3 (n-6) △ 17 desaturase C20:4 (n-3)
△ 8, 11, 14
△ 8, 11, 14, 17
Elongase
△ 6 desaturase △ 6 desaturase
Elongase
Elongase
C20:3 (n-3)
△ 11, 14, 17
Elongase
△ 5 desaturase
△ 8 desaturase
△ 5 desaturase
C20:4 (n-6) △ 17 desaturase C20:5 (n-3)
△ 5, 8, 11, 14
△ 5, 8, 11, 14, 17
Elongase
Elongase
C22:4 (n-6)
△ 7, 10, 13, 16
C22:5 (n-3)
△ 7, 10, 13, 16, 19
C22:5 (n-6)
△ 4, 7, 10, 13, 16
C22:6 (n-3)
△ 4, 7, 10, 13, 16, 19
△ 4 desaturase
△ 4 desaturase
Figure 1.13 The elongation and desaturation of acyl chain of fatty acids
(Guschina & Harwood, 2006).
44
Chapter 1. Literature review and research aim
Figure 1.14 The biosynthesis of TAG in microalgae (Huang et al., 2010)
1.3.5.2 Factors affecting lipid accumulation and fatty acid composition
The lipid content and fatty acid composition are species/strain-specific
and can be greatly affected by chemical and physical stimuli. The main chemical
stimuli are nutrient concentration, salinity and pH of medium and the main
physical stimuli are light intensity and temperature.
1.3.5.2.1 Nutrients
Of all the nutrients surveyed, nitrogen is the most critical one affecting
lipid metabolism in algae. The influence of nitrogen source and concentration on
lipid and fatty acid production has been investigated in a number of microalgae
(Takagi et al., 2000; Li et al., 2005; Li et al., 2008a; Solovchenko et al., 2008;
Hsieh & Wu, 2009; Pruvost et al., 2009; Li et al., 2010). Nitrate was suggested to
45
Chapter 1. Literature review and research aim
be superior to other nitrogen sources such as urea and ammonium for algal lipid
production (Li et al., 2008a). Generally, low concentration of nitrogen in the
medium favors the accumulation of lipids particularly TAGs and total fatty acids.
But in some cases, nitrogen starvation caused decreased synthesis of lipids and
fatty acids (Saha et al., 2003). As for the effect of nitrogen on fatty acid
composition, there is no general trend available and it appears to be species/strain
dependent. For example, in cyanobacteria, increased levels of C16:0 and C18:1
and decreased C18:2 level were observed in response to nitrogen deprivation
(Piorreck & Pohl, 1984). In the marine alga Pavlova viridis, nitrogen depletion
resulted in an increase in saturated, monounsaturated fatty acids and C22:6 (n-3)
contents (Li et al., 2005). Nitrogen starvation brought about a strong increase in
the proportion of C20:4 (n-6) in the green algal Parietochloris incisa
(Solovchenko et al., 2008).
Other types of nutrient that affect lipid production include phosphorus,
silicon, sulfur and iron. In response to phosphorus deprivation, the enhanced
accumulation of lipids was observed in Ulva pertusa, Scenedesmus sp.LX1,
Monodus subterraenus, Phaeodactylum tricornutum, Chaetoceros sp., Isochrysis
galbana and Pavlova lutheri (Reitan et al., 1994; Floreto et al., 1996;
Khozin-Goldberg & Cohen, 2006; Li et al., 2010), while reduced lipid production
was demonstrated in Nannochloris atomus and Tetraselmis sp. (Reitan et al.,
1994). Among the marine species surveyed by Reitan et al. (1994), phosphorus
deprivation caused increased contents of C16:0 and C18:1 and decreased
polyunsaturated fatty acids; whereas in U. pertusa, phosphorus deprivation
decreased the proportion of C16:0 (Floreto et al., 1996). Silicon is the key nutrient
for lipid metabolism of diatoms. The deficiency of silicon increased the
accumulation of neutral lipids, as well as of saturated and monounsaturated fatty
acids in Cyclotella cryptic (Roessler, 1988). Studies also suggested that sulfur
limitation could promote lipid production in microalgae (Otsuka, 1960; Sato et al.,
2000). In addition, iron at a relative high concentration could induce the
accumulation of lipids in the green alga Chlorella vulgaris (Liu et al., 2008).
46
Chapter 1. Literature review and research aim
1.3.5.2.2 Light
Light has a marked effect on the lipid production and fatty acid
composition in microalgae (Nichols, 1965; Koskimies-Soininena & Nyberg, 1987;
Sukenik et al., 1989; Napolitano, 1994; Brown et al., 1996; Zhekisheva et al.,
2002, 2005; Khotimchenko & Yakovleva, 2005; Damiani et al., 2010). Generally,
low light intensity favors the formation of polar lipids such as the membrane
lipids associated with the chloroplast; whereas high light intensity benefits the
accumulation of neutral storage lipids in particular TAGs. In H. pluvialis, for
example, high light resulted in a great increase of both neutral and polar lipids, but
the increase extent of neutral lipids was much greater than that of polar lipids,
leading to the dominant proportion of neutral lipids in the total lipids (Zhekisheva
et al., 2002, 2005). Although the effect of light intensity on fatty acid composition
differs among the algal species and/or strains, there seems be a general trend, with
a few exceptions, that the increase of light intensity contributes to the enhanced
proportions of saturated and monounsaturated fatty acids and the concurrently the
reduced proportion of polyunsaturated fatty acids (Sukenik et al., 1989;
Zhekisheva et al., 2002, 2005; Eabregas et al., 2004).
1.3.5.2.3 Temperature
Temperature can affect lipid and fatty acid composition of many
organisms including algae (Sato & Murata, 1980; Lynch & Thompson, 1982;
Henderson & Mackinlay, 1989; Wada & Murata, 1990; Thompson et al., 1992;
Renaud et al., 1995, 2002). In response to temperature shift, algae commonly alter
the physical properties and thermal responses of membrane lipids to maintain
fluidity and function of membranes (Somerville, 1995). In general, increased
temperature causes increased fatty acid saturation and at the same time decreased
fatty acid unsaturation. For example, C14:0, C16:0, C18:0 and C18:2 increased
and C18:3 (n-3), C18:4, C20:5 and C22:6 decreased in Rhodomonas sp., and
47
Chapter 1. Literature review and research aim
C16:0 increased and C18:4 decreased in Cryptomonas sp. when temperature
increased (Renaud et al., 2002). As for the effect of temperature on lipids, it
differs with a species-dependent manner. In response to increasing temperature,
increased lipid content was observed in Nanochloropsis salina (Boussiba et al.,
1987) and Ochromonas danica (Aaronson, 1973); decreased lipid content was
observed in Chroomonas salina (Henderson & Mackinlay, 1989), Nitzschia
paleacea (Renaud et al., 1995) and Chaetoceros sp. (Renaud et al., 2002); while in
Chlorella sorokiniana (Patterson, 1970) and Isochrysis sp. (T.ISO) (Renaud et al.,
2002), lipid content showed no significant change.
1.3.5.2.4 Salinity
Salinity stress can influence cell membrane permeability and fluidity.
Cells adapt themselves to high salinity through the compositional changes in
sterols and polar lipids (Parida & Das, 2005). The effect of salinity on lipid profile
and fatty acid composition was reported in some species of microalgae such as
Botryococcus braunii, Chaetoceros cf. wighamii, Cladophora vagabunda,
Crypthecodinium cohnii, Dunaliella tertiolecta, Nitzschia laevis (Elenkov et al.,
1996; de Castro Araujo & Garcia, 2005; Takagi et al., 2006; Rao et al., 2007;
Chen et al., 2008). Generally, salinity stress favors the accumulation of lipids and
causes enhanced fatty acid saturation and reduced fatty acid unsaturation in
microalgae.
1.4 The green alga Chlorella zofingiensis
The green algae (Chlorophyta) are named for their typical color, the
bright grass-green characteristic of land plants. They are a large group of algae
consisting of more than 6000 species distributed in different habitats (Thomas,
2002). Most of green algae live in fresh water, with a minor proportion locate in
48
Chapter 1. Literature review and research aim
marine water, damp soil, land plants or even snow and ice (Lee, 2008). Green
algae exhibit a variety of morphologies. They can be motile or no-motile,
unicellular or colonial flagellates, usually but not always with two flagella per cell
(Wikipedia, 2010).
C. zofingiensis is a fresh water microalga belonging to Class
Chlorophyceae, Order Chlorococcales and Family Chlorellaceae (Pickett-Heaps,
1975). C. zofingiensis cells are non-motile and in unicellular and spherical form,
with the cell size ranging from 2 μm to 15 μm in diameter. Through the formation
of autospores, C. zofingiensis asexually reproduces daughter cells from
non-motile parental cells (Lee, 2008).
C. zofingiensis can grow well photoautotrophically, heterotrophically and
mixotrophically (Orosa et al., 2000, 2001; Ip et al., 2004; Ip & Chen 2005; Sun et al.
2008). Under optimal growth conditions, C. zofingiensis accumulated mainly
chlorophylls and primary carotenoids such as lutein; while under stress conditions
like high light, nitrogen starvation, or high carbon/nitrogen ratio in the dark, C.
zofingiensis predominantly produced astaxanthin and thus exhibited a deep red
color (Orosa et al., 2001; Ip & Chen, 2005). So far, studies about C. zofingiensis
have primarily focused on astaxanthin production, while the investigations about
lipids and fatty acids have been rarely touched.
1.4.1 Pigment profiles
Both primary and secondary carotenoids have been found in C.
zofingiensis. Like plants and other algae, C. zofingiensis synthesizes and
accumulates primary carotenoids (e.g., β-carotene, lutein and zeaxanthin) in
chloroplast (Rise et al., 1994; Grunewald et al., 2001). In contrast, secondary
carotenoids such as astaxanthin, canthaxanthin and adonixanthin are found to
accumulate in lipid bodies outside the chloroplast (Rise et al., 1994; Del Campo et
al., 2004). Commonly, the accumulation of secondary carotenoids is considered
49
Chapter 1. Literature review and research aim
related to the stress conditions, under which algal cells get protected against
oxidative damages by these anti-oxidative carotenoids through quenching the
excessive ROS and other free radicals (Rise et al., 1994; Bar et al., 1995). For
example, when growing under high light and nitrogen starvation, C. zofingiensis
produced mainly secondary carotenoids, with around 70% being astaxanthin (Rise
et al., 1994).
The photoautotrophic cultivation of C. zofingiensis generally gives a low
cell biomass and thus a low astaxanthin yield. For example, as reported by Orosa
et al. (2000), the cell biomass and carotenoid concentrations in photoautotrophic
culture were limited to 0.72 g L-1 and 2.81 mg L-1 , respectively. Furthermore, the
attenuated light absorption caused by mutual shading of cells in large-scale
cultures can severely affect the productivity of algal biomass and products (Chen,
1996). To overcome such problems, mixotrophy and heterotrophy have been
employed for C. zofingiensis growth and astaxanthin production (Ip et al., 2004;
Ip & Chen, 2005). In heterotrophic culture, C. zofingiensis utilized organic
substrate (e.g., glucose) as the sole energy and carbon source and achieved a much
higher cell density as compared with in photoautotrophic culture (Orosa et al.,
2000; Ip et al., 2004). Ip (2005) intensively investigated the heterotrophic
production potential of astaxanthin by Chlorella zofingiensis: (1) a high initial
carbon to nitrogen ratio (e.g., 180) was found to enhance astaxanthin biosynthesis
in algal cells; (2) through the manipulation of the medium nutrients and
cultivation environments, the maximal yield of astaxanthin (9.9 g L-1) was
obtained when the algal cells were grown on the medium consisting of 0.44 g L-1
nitrate, 0.16 g L-1 phosphate and 8 mg L-1 ferrous ion at 30 and pH 5.5; (3)
external supplementation of ROS or reactive nitrogen species/reactive nitrogen
intermediate further induced astaxanthin synthesis in the algal cells; (4)
endogenous arousal of oxidative stresses caused by salt or by inhibition of
antioxidative enzymes also triggered increased astaxanthin production. By using a
well designed fed-batch fermentation strategy, up to 32.4 mg L-1 astaxanthin was
obtained in the algal cells, indicating the potential of employment of heterotrophic
50
Chapter 1. Literature review and research aim
C. zofingiensis cells for commercial production of astaxanthin on a large scale
(Sun et al., 2008). A general pigment profile of C. zofingiensis under heterotrophic
conditions is shown in Figure 1.15. Astaxanthin occurred predominantly in the
form of mono- and di-esters.
51
Chapter 1. Literature review and research aim
0.020
12
13
21
18
24
25
AU
0.015
0.010
32
16
1
27
26
20
0.005
3
9
2
14
5 6
10
78
11
10.00
11.00
4
15
19
17
37
22
23
28
30
33
29 31
34
38
35 36
39
0.000
8.00
9.00
12.00
13.00
14.00
15.00
16.00
17.00 18.00
Minutes
19.00
20.00
21.00
22.00
23.00
24.00
25.00
26.00
27.00
Figure 1.15 A HPLC chromatogram showing a general pigment profile of C. zofingiensis under heterotrophic conditions. 1 Neoxanthin; 2
Violaxanthin; 3 Antheraxanthin; 4,5,7,8,28,29,30,35,36 Unknown or degraded lutein and chlorophylls; 6 Astaxanthin; 9,10 Adonixanthin;
11 Phoenicoxanthin (adonirubin); 12 Lutein; 13,14,15 Zeaxanthin; 16,17 Canthaxanthin; 18 Chlorophyll b; 19 Hydroxyechinenone; 20,22
β−Cryptoxanthin; 21 Chlorophyll a; 23 Echinenone; 24,26 Astaxanthin mono-ester; 25,27 Adonixanthin mono-ester; 31 α−Carotene;
32,33,34 β−Carotene; 37,39 Astaxanthin di-ester; 38 Adonixanthin di-ester. Source: Sun, 2009.
52
Chapter 1. Literature review and research aim
1.4.2 Astaxanthin biosynthesis
β-carotene is considered to be the major precursor for astaxanthin.
Astaxanthin is biosynthesized from β-carotene via two proposed pathways: one
starting from the oxygenation and then the hydroxylation producing the
intermediates of echinenone, canthaxanthin and adonirubin, and the other from the
hydroxylation and then the oxygenation via β-cryptoxanthin, zeaxanthin and
adonixanthin as intermediates (Figure 1.16). It has been demonstrated that
astaxanthin biosynthesis in the green alga H. pluvialis followed the pathway as
indicated by solid dash in Figure 1.16 (Breitenbach et al., 1996; Fraser et al., 1998;
Linden, 1999). In C. zofingiensis, however, there appears to be another case. The
ketolase from H. pluvialis can efficiently convert canthaxanthin to astaxanthin,
but shows a poor ability in conversion of zeaxanthin to astaxanthin (Fraser et al.,
1997, 1998). In contrast, the C. zofingiensis ketolase exhibits a bi-functional
activity: efficient introduction of keto groups to both β-carotene and zeaxanthin
(Huang et al., 2006). Considering that substantial canthaxanthin (around 30% of
total ketocarotenoids) accumulated and might represent the end product of
oxygenated β-carotene in C. zofingiensis, the carotenoid hydroxylase from this
alga might not accept canthaxanthin as a substrate (Rise et al., 1994). Based on
the above results as well as the accumulated adonixanthin in both transformed E.
coli and induced C. zofingiensis cells, a possible astaxanthin biosynthetic pathway
different from that in H. pluvialis was proposed as indicated by dash arrow in
Figure 1.16: oxygenation of zeaxanthin rather than hydroxylation of
canthaxanthin (Huang et al., 2006; Wang & Chen, 2008).
53
Chapter 1. Literature review and research aim
Phytoene
PDS
ζ-carotene
β-carotene
BKT
CHY
β-cryptoxanthin
Echinenone
CHY
BKT
Zeaxanthin
Canthaxanthin
BKT
CHY
Adonirubin
Adonixanthin
BKT
CHY
Astaxanthin
Figure 1.16 Proposed pathways of astaxanthin biosynthesis in algae. PDS,
phytoene desaturase; BKT, carotenoid ketolase; CHY, carotenoid hydroxylase.
Adapted from Huang et al. (2006).
Although high light, salinity and glucose are known to enhance the
biosynthesis of astaxanthin in C. zofingiensis, the underlying mechanism of their
reaction remains largely unknown at molecular level before the availability of
genes involved in astaxanthin biosynthesis (Bar et al., 1995; Ip, 2005). Certain
key genes, namely phytoene desaturase (PDS), carotenoid ketolase (BKT) and
carotenoid hydroxylase (CHYb) have been isolated and characterized from C.
zofingiensis recently (Huang et al., 2006, 2008). High light up-regulates the
transcripts of PDS, CHYb, and BKT and greatly enhances the biosynthesis of
zeaxanthin, canthaxanthin, and astaxanthin; while salinity stress only up-regulates
54
Chapter 1. Literature review and research aim
the transcript of BKT and enhances the biosynthesis of canthaxanthin and
astaxanthin (Li et al., 2009). This discrepancy might result from the generation of
different intracellular ROS, which stimulated the up-regulation of specific
carotenoid genes (Ip, 2005; Li et al., 2009). In dark-grown C. zofingiensis, glucose
sensing (phosphorylation of glucose) directly up-regulates the transcription of
CHYb, while the mitochondrial alternative pathway is closely related to the
regulation of BKT, through which the biosynthesis of astaxanthin is regulated (Li
et al., 2008b). Moreover, the involvement of transcriptional control on the
carotenoid biosynthesis in C. zofingiensis by PDS, CHYb and BKT is suggested
(Li et al., 2008b, 2009; Liu et al., 2010).
Taken together, the availability of carotenoid genes, the demonstrated
astaxanthin biosynthetic pathway and the possible mechanism of astaxanthin
regulation provide a solid foundation for the genetic engineering of C. zofingiensis
with an attempt to enhance astaxanthin production.
1.4.3 Lipid and fatty acid profiles
Some Chlorella species such as C. vulgaris and C. protothecoides have
been analyzed in terms of lipids and fatty acids (Miao & Wu, 2004; Cleber Bertoldi
et al., 2006; Liu et al., 2008). No such information so far, however, is available for
C. zofingiensis. Considering its fast growth and versatile cultivation modes, C.
zofingiensis may have the potential as a biodiesel feedstock candidate, which will
require further investigations, for example the lipid and fatty acid analyses.
1.5 Research aim
C. zofingiensis represents a fast-growing green microalga that can grow
under photoautotrophic, mixotrophic and heterotrophic conditions. It shows great
potential as a source of the high-value carotenoid astaxanthin and may also be a
55
Chapter 1. Literature review and research aim
potential source of lipids for biodiesel production. However, the relatively low
cellular content of astaxanthin and the shortage of information about lipid and
fatty acid profiles in C. zofingiensis hamper its commercial application. So the
aim of this research was to improve the cellular accumulation of astaxanthin in C.
zofingiensis and to assess the potential use of this alga as a biodiesel feedstock.
Following shows the major work of my study:
(1)
The PDS gene from C. zofingiensis was isolated and characterized.
(2)
C. zofingiensis mutants with enhanced astaxanthin accumulation were
generated by treatment of chemical mutagens coupled with norflurazon
selection.
(3)
The molecular characterization of one mutant E17 was conducted.
(4)
The transformation of C. zofingiensis with a mutated PDS gene from E17
and the analysis of obtained transformants with promoted astaxanthin
accumulation were performed.
(5)
The lipid and fatty acid profiles of photoautotrophic and heterotrophic C.
zofingiensis were compared.
(6)
The lipid production and fatty acid profile of C. zofingiensis cultured in
the dark with various carbon sources were investigated.
(7)
The optimized conditions for fatty acid production by heterotrophic C.
zofingiensis were established.
(8)
The key genes BC and SAD involved in fatty acid biosynthesis were
isolated and characterized from C. zofingiensis.
56
PART II
GENETIC ENGINEERING OF CHLORELLA
ZOFINGIENSIS FOR ENHANCED ASTAXANTHIN
PRODUCTION
57
Chpater 2. Isolation and characterization of the phytoene desaturase gene from Chlorella
zofingiensis
Chapter 2
Isolation and characterization of the phytoene desaturase
gene from Chlorella zofingiensis
2.1 Abstract
Phytoene desaturase (PDS) is a rate-limiting enzyme in carotenoid
biosynthesis. Algal PDS is inhibited by some herbicides, leading to the bleaching
of cells due to the destruction of chlorophylls. Specific point mutations in PDS
confer resistance to the herbicide norflurazon, suggesting that mutated PDS could
be used as a dominant selectable marker for genetic engineering of algae, for
which very few selectable markers are available. In this chapter, the isolation and
characterization of the PDS gene from Chlorella zofingiensis were reported. The
open reading frame of this PDS gene, interrupted by six introns, encoded a
polypeptide of 558 amino acid residues. The deduced protein sequence showed
significant homology with phytoene desaturases from other algae, cyanobacteria
and higher plants. Expression of the PDS gene in Escherichia coli demonstrated
that the enzyme was able to convert phytoene to ζ-carotene. The PDS gene in C.
zofingiensis was shown to be up-regulated by high light and glucose treatment.
With a single amino acid change (L516R), the mutated PDS exhibited 35-fold
greater resistance to norflurazon but also a lower desaturation activity than the
unaltered enzyme. The results indicated that certain point mutation could make
Chlorella PDS herbicide-resistant and potentially useful in genetic engineering of
C. zofingiensis.
2.2 Introduction
As the biosynthesis of astaxanthin is observed only in a limited number
58
Chpater 2. Isolation and characterization of the phytoene desaturase gene from Chlorella
zofingiensis
of organisms (e.g., some marine bacteria, the red yeast Xanthophyllomyces
dendrorhous, and some green algae) (Johnson & Schroeder, 1995), potential
production of astaxanthin from microorganisms and transgenic plants has been the
subject of intensive investigations in recent years (Gong & Chen, 1997; Mann et
al., 2000; Stalberg et al., 2003; Ip & Chen, 2005; Steinbrenner & Sandmann,
2006). The unicellular green alga Haematococcus pluvialis has the highest
astaxanthin accumulation (up to 4% of dry biomass) (Boussiba, 2000). However,
its slow growth rate restricts its application. The green microalga Chlorella
zofingiensis is a promising host for boosting astaxanthin production by metabolic
engineering because the alga can grow fast (with a specific growth rate of over
0.031 h-1) and produce astaxanthin (with a astaxanthin yield up to 10.3 mg L-1) in
the dark with glucose as sole carbon and energy source (Ip & Chen, 2005).
Moreover C. zofingiensis has a similar mechanism of storing astaxanthin as H.
pluvialis, suggesting that C. zofingiensis might be genetically modified to
accumulate much higher amounts of astaxanthin.
While it is now relatively easy to generate a transgenic plant, there are
still significant technical challenges to develop functional transgenic systems for
many commercially important algae. Endogenous promoters and terminators
proved critical for the expression of heterologous genes in algae (Cerutti et al.,
1997; Ohresser et al., 1997; Poulsen & Kroger, 2005; Steinbrenner & Sandmann,
2006). Thus, in order to improve the astaxanthin production in C. zofingiensis by
genetic engineering, it is critical that an efficient selectable marker be developed.
Phytoene desaturase (PDS) is the rate-limiting enzyme in carotenoid biosynthesis
(Chamovitz et al., 1993). The plant and algal PDS carries out the first two-step
desaturation of phytoene leading to the formation of ζ-carotene (Pecker et al.,
1992; Sandmann, 1994). PDS is inhibited by some herbicides, which causes
chlorophyll destruction and thus the cell bleaching. However, certain
point-mutations in PDS were found to confer resistance to the herbicide
norflurazon (Chamovitz et al., 1993; Arias et al., 2006). Thus, an endogenous PDS
gene could be modified and used as a dominant selectable marker for nuclear
transformation of commercially important algae for which stable transformation is
still problematic.
Plant and algal PDS genes are highly conserved and have similar
catalytic properties (Pecker et al., 1992; Linden et al., 1995). The aim of the
59
Chpater 2. Isolation and characterization of the phytoene desaturase gene from Chlorella
zofingiensis
present study is to isolate and characterize the PDS gene from C. zofingensis. In
addition, a modified PDS gene resistant to norflurazon was generated by
site-directed mutagenesis. The obtained herbicide-resistant PDS gene could be
very useful for genetic engineering of carotenoid biosynthesis in C. zofingiensis.
2.3 Materials and methods
2.3.1 Algal strain and culture conditions
The green microalga C. zofingiensis (ATCC 30412) was obtained from
the American Type Culture Collection (ATCC, Rockville, MD, USA). This algal
strain was maintained and cultured at 25 °C under continuous illumination of 25
µmol photon m-2 s-1 in Kuhl medium (Kuhl & Lorenzen, 1964) consisting of (per
liter) 1.01 g KNO3; 0.62 g NaH2PO4∙H2O; 0.089 g Na2HPO4∙2H2O; 0.247g
MgSO4∙7H2O; 14.7 mg CaCl2∙2H2O; 6.95 mg FeSO4∙7H2O; 0.061 mg H3BO3;
0.169 mg MnSO4∙H2O; 0.287 mg ZnSO4∙7H2O; 0.0025 mg CuSO4∙5H2O; and
0.01235 mg (NH4)6MO7O24∙4H2O. The pH of the medium was adjusted to pH 6.5
prior to autoclaving. To investigate the effect of high light and glucose on
expression of the PDS gene, photoautotrophically grown cells in exponential
growth phase were exposed to continuous high light (120 μmol photon m-2 s-1) or
provided with glucose (50 mM) in the dark for 0 to 48 h.
2.3.2 Genomic DNA and RNA isolation
DNA was extracted using a modified cetyltrimethylammonim bromide
(CTAB) method (Stewart & Via, 1993). RNA was isolated from aliquots of about
108 cells using the TriPure isolation reagent (Roche, Mannheim, Germany)
according to the manufacturer’s manual. The concentration of DNA and total
RNA was determined spectrophotometrically at 260 nm.
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Chpater 2. Isolation and characterization of the phytoene desaturase gene from Chlorella
zofingiensis
2.3.3 Cloning of PDS cDNA and its corresponding gene
Degenerate primers dF and dR were designed based on the conserved
amino acid sequences (GKVAAWK and LQWKEHS, respectively) of the PDS
proteins from Chlamydomonas reinhardtii, H. pluvialis, Synechococcus sp. PCC
7942, Dunaliella salina and Arabidopsis thaliana (Chamovitz et al., 1991;
Scolnik & Bartley, 1993; Harker & Hirschberg, 1997; McCarthy et al., 2004; Zhu
et al., 2005). They were used for the amplification of a partial PDS cDNA from C.
zofingiensis, which would provide sequence information for designing specific
primers for rapid amplification of 5′ and 3′ cDNA ends (RACE). The primer sets
used in this study are listed in Table 2.1. RACE was performed using the method
described by Huang and Chen (2006). Genomic walking of the PDS gene was
performed according to the approach described in the Universal GenomeWalker
kit (Clontech, Palo Alto, CA, USA).
2.3.4 Functional analysis of PDS cDNA
The PDS open reading frame was digested by EcoRI and BamHI and
inserted into the corresponding sites of pUC18 (Stratagene, La Jolla, CA, USA) as
an in-frame fusion to the lacZ gene, resulting in plasmid pUC-czPDS. The
mutated PDS cDNA (with codon position 516 changed from a leucine codon to an
arginine codon) was obtained by site-directed mutagenesis using the Quickchange
mutagenesis kit (Stratagene). The mutation in the PDS cDNA (termed as
czPDS-L516R) was verified by sequencing.
E. coli strain JM109 was used as a host for functional expression
experiments by co-transformation of pACCRT-EB that harbors the carotenoid
biosynthesis genes for producing phytoene (Misawa et al., 1995) with plasmid
pUC-czPDS or pUC-czPDS-L516R. Cells were grown in LB medium
supplemented with 100 μg mL-1 ampicillin, 50 μg mL-1 chloramphenicol and 1
mM isopropylthiogalactose at 28 °C for 2 days. Pigments were extracted and
analyzed according to Huang et al. (2006). E. coli cells were collected by
centrifugation and then freeze-dried. Extraction was carried out with a mixture of
dichloromethane and methanol (25:75, V/V) until the cell debris was almost
61
Chpater 2. Isolation and characterization of the phytoene desaturase gene from Chlorella
zofingiensis
colorless. The combined extracts were evaporated by nitrogen gas to dryness and
dissolved in acetone for the subsequent high-performance liquid chromatography
(HPLC) analysis.
2.3.5 RT-PCR assay
Total RNA (1 µg) was extracted from C. zofingiensis cells illuminated
with high light (120 µmol photons m-2 s-1) or treated with 50 mM glucose in the
dark for 0 to 48 h. The RNA was reverse transcribed to cDNA with SuperScript III
First-Strand Synthesis System for RT-PCR (Invitrogen, Carlsbad, CA) primed
with oligo(dT) according to the manufacturer's instructions. The C. zofingiensis
actin gene was used to normalize the level of RNA template used in the reaction
(Huang et al., 2006). Amplification of the cDNA was done by conventional PCR
[94 °C for 2 min followed by 26 cycles (for PDS gene) or 22 cycles (for actin
gene) of 94 °C for 15 s, 56 °C for 15 s, 72 °C for 30 s]. PCR products were
separated on 2% agarose gels and stained with ethidium bromide (EB) for
photography (Biorad, Hercules, CA, USA).
2.3.6 Preparation of enzyme and substrate
Plasmids pUC-czPDS and pUC-czPDS-L516R were respectively
transformed into E. coli 109 for PDS expression. Transformed cells were grown in
SOB medium containing 50 μg mL-1 ampicillin at 37 °C with vigorous shaking,
and 1 mM IPTG was added when the optical density at 600 nm reached 0.5. Cells
were harvested after a 5-h
induction period, re-suspended in 0.1 × sodium
phosphate buffer (pH 7.2, containing 1 mM DTT), and then passed through a
French pressure cell (Spectronics Instruments, Rochester, NY, USA) at an internal
pressure of 20 MPa. 10 μg mL-1 DNase was added to the broken cell extract and
the mixture was incubated on ice for 15 min. Cell debris was removed from the
suspension by centrifugation at 10,000 g and the resultant supernatant was
adjusted to 1 mg mL-1 of crude protein and used as the source of PDS enzyme.
Protein concentration was determined according to Bradford (1976). The substrate
62
Chpater 2. Isolation and characterization of the phytoene desaturase gene from Chlorella
zofingiensis
phytoene was extracted with acetone from E. coli JM109/pACCRT-EB
freeze-dried cells according to procedures described by Breitenbach et al. (2001).
2.3.7 In vitro PDS assay
The reaction mixture contained 1 mL of enzyme extract, 5 μL of the
substrate phytoene (1 μg) in acetone, and 5 μL of decyl plastoquinone (10 mM
solution
in
methanol,
Sigma,
St
Louis,
MO,
USA)
and
0.25
mg
L-α-phosphatidylcholine (Sigma) in a sodium phosphate buffer suspension. The
assays were carried out at 28 °C with vigorous shaking for various periods (0.5 to
12 h) to investigate the optimized incubation time for the enzymatic reaction. The
reaction was terminated by the addition of 1 mL of methanol. To survey the
herbicide resistance of phytoene desaturases, norflurazon (5 μL in methanol,
Sigma) was added to the reaction mixture and incubated on ice for 15 min prior to
mixing it with phytoene. Five final concentrations of norflurazon ranging from
0.05 μM to 0.8 μM for the unaltered PDS enzyme, and from 0.5 μM to 8 μM for
the engineered PDS-L516R enzyme, were tested. The residual phytoene and
enzymatically formed ζ-carotene were extracted from the incubation mixture with
diethyl ether/petroleum ether (1:9, v/v), evaporated to dryness under a nitrogen
stream and re-suspended in acetone for subsequent HPLC analysis.
2.3.8 Pigment analysis
Extracted pigments were separated on a Waters Spherisorb® 5 μm ODS2
4.6 250 mm analytical column with a Waters HPLC system (Waters, Milford, MA,
USA). The mobile phase consists of solvent A (acetonitrile/methanol/0.1 M
Tris-HCl (pH 8.0), 84:2:14, by vol.) and solvent B (methanol/ethyl acetate, 68:32,
v/v). Pigments were eluted at a flow rate of 1.2 mL min-1 with a linear gradient
from 100% solvent A to 100% solvent B over a 15 min period, followed by 10
min of solvent B. The absorption spectra of the pigments were shown between
250 and 700 nm. Individual carotenoids were identified by their absorption
spectra and their typical retention times compared to standard samples of pure
63
Chpater 2. Isolation and characterization of the phytoene desaturase gene from Chlorella
zofingiensis
carotenoids (Sigma).
2.4 Results and discussion
2.4.1 Cloning of C. zofingiensis PDS gene
With the primers dF and dR (Table 2.1), a 140-bp fragment of the PDS
gene was amplified (Figure 2.1, lane 1). Searches of the GenBank database using
the BLAST program demonstrated that the nucleotide sequence of this fragment
shared about 88% homology with that of C. reinhardtii and H. pluvialis. Based on
this sequence information, two pairs of specific primers (F1, R1, F2, and R2 in
Table 2.1) were designed for 5′ and 3′ RACE using the method described by
Huang et al. (2006), which generated 2.1-kb fragment (Figure 2.1, lane 2). The
sequence of the fragment was determined as a fusion of the 5′ and 3′ ends of a
putative PDS cDNA. The coding region of the cDNA was amplified using the
primers F3 and R3, which annealed to the start and stop codon regions,
respectively. The PCR product (Figure 2.1, lane 3) was sequenced and contained a
1677-bp open reading frame encoding a deduced PDS with 558 amino acid
residues (Figure 2.2A, GeneBank accession No. EF621405). Upstream of the
translation start codon is a 117-bp 5′ untranslated region and between the stop
codon and poly (A) tail is a 3′ untranslated region of 509 bp nucleotides. TGTAA,
the sequence considered as a potential polyadenylation signal in green algae
(Schmitt et al., 1992), however, is not found in the 3′ untranslated region of this
PDS gene. The GC content of the PDS coding region is 50.8%, which is much
lower than GC content of PDS genes from C. reinhardtii (63.7%) and H. pluvialis
(58.6%).
Protein sequence alignments showed that the PDS of C. zofingensis
shared a high homology with that of other algae, cyanobacteria, and higher plants,
and particularly with the green algae C. reinhardtii (74.6%) and H. pluvialis
(70.0%) (Figure 2.3). No significant identity, however, is observed between the C.
zofingensis PDS and its counterparts from bacteria and fungi, except in the
N-terminal region which contains the dinucleotide binding motif (data not shown).
Similar to H. pluvialis, the C. zofingensis PDS contains an N-terminal extension
64
Chpater 2. Isolation and characterization of the phytoene desaturase gene from Chlorella
zofingiensis
relative to the bacterial and fungal polypeptides. This extended sequence may
serve as a transit peptide to direct the transport of PDS into plastids, since the PDS
is nuclear-encoded but functions exclusively in the chloroplast (Grunewald et al.,
2000). Furthermore, a transit peptide of 40 amino acids in the N-terminal
extension of the PDS was predicted by using the ChloroP 1.1 software
(http://www.cbs.dtu.dk/services/Chlorop).
Table 2.1 Primer sets and PCR product characteristics
Aim
Oligonucleotide sequence 5′-3′
Partial PDS fragment
Product size (kb)
0.14
dF
GGCAAGGTNGCYGCNTGGAA
dR
GAGTGCTCCTTCCACTGCA
5′ and 3′ RACE
2.1
F1
AGGCCTGCACATCTTCTTTGGT
R1
CCAGTCACCATCCTCATCCTTC
F2
GATGAATGTATTTGCTGAACTGGGC
R2
CCTTCCATGCGGCAACCTTGC
PDS coding region*
1.7
F3
gcgaattccgATGCAACAGGCTCTAGGGCAG
R3
ggggatccACTGTGCTGAGCTTGC
3′ PDS walking
0.8
F4
GTGGCAAGCTGGCTACTGAGG
Ap1
GTAATACGACTCACTATAGGGC
F5
CGTCACTGGAAGGTTGCACAC
Ap2
ACTATAGGGCACGCGTGGT
PDS gene
6.3
F6
GGCGCATAGGATTGACAAGCTT
R6
GGGTGCCGCCGATCTGTGG
PDS expression
0.29
F2
R7
GGCCAGTGCCTTAGCCATAGCG
Site-directed mutation
F8 (R8)
CACCAAACAAAAGTACCgTGCATCCATGGAA
GGTGCC (complement reverse)
F: forward; R: reverse. R1, Ap1, R2 and Ap2 were also used for 5′ PDS walking.
*EcoRI and BamHI sites (lower cases underlined) were added for cloning the
gene into the corresponding cut sites of pUC 18 vector.
65
Chpater 2. Isolation and characterization of the phytoene desaturase gene from Chlorella
zofingiensis
To characterize the genomic PDS sequence, 5′ and 3′ genomic walking
by PCR was performed and two fragments were obtained (Figure 2.1, lane 4 and
5). Sequencing results indicated that these two fragments contained the promoter
and terminator regions of the PDS gene, respectively. Based on the sequence
information of the isolated promoter and terminator fragments, the specific
primers F6 and R6 were designed for the amplification of PDS gene and a 6.3-kb
fragment was obtained (Figure 2.1 lane 6). Analysis of this amplified nucleotide
sequence revealed that the product was the genomic sequence of PDS cDNA
(GeneBank accession No. EF621406). The generated PDS gene is flanked by a
2-kb promoter region and a 560-bp terminator region, and its coding region is
interrupted by six introns of 451, 217, 171, 111, 222 and 263 bp, respectively
(Figure 2.2 B). All introns start with GT and end with CAG.
Figure 2.1 PCR-based isolation of C. zofingiensis PDS gene. Specific fragments
were amplified using genomic DNA (lane 1, 4, 5 and 6) or cDNA (lane 2 and 3) as
template. Primers are listed in Table 1. Lane 1, dF+dR; lane 2, F1+R1 (first round)
and F2+R2 (second round); lane 3, F3+R3; lane 4, R1+Ap1 (first round) and
R2+Ap2 (second round); lane 5, F4+Ap1 (first round) and F5+Ap2 (second
round); lane 6: F6+R6. M1, 100 bp ladder; M2, lambda DNA/StyI ladder
(Fermentas).
2.4.2 Functional analysis of C. zofingiensis PDS cDNA in E. coli
In algae the conversion of phytoene, the first C40-carotene in the
carotenoid biosynthesis pathway, to ζ-carotene via phytofluene is catalyzed by
66
Chpater 2. Isolation and characterization of the phytoene desaturase gene from Chlorella
zofingiensis
Figure 2.2 Amino acid sequence alignment of C. zofingiensis PDS with its
counterparts from A. thaliana (AAA20109), C. reinhardtii (AAT38476), D. salina
(AAY26317), H. pluvialis (CAA60479) and Synechococcus sp. PCC 7942
(CAA39004). Amino acid residues which are either well or perfectly conserved in
all sequences are indicated by (.) or (*) above the alignment, respectively. Primes
are underlined with names.
67
Chpater 2. Isolation and characterization of the phytoene desaturase gene from Chlorella
zofingiensis
PDS through a two-step desaturation. To find out the enzymatic activity of the
polypeptide encoded by the C. zofingiensis PDS cDNA, an in vivo functional
expression analysis was performed. E. coli harboring pACCRT-EB produced the
phytoene, which is colorless, while E. coli harboring both pACCRT-EB and
pUC-czPDS exhibited a yellow color, indicating the formation of new pigments in
the bacterium. HPLC analysis of extracts from the different E. coli transformants
is shown in Figure 2.4. The E. coli carrying the plasmid pACCRT-EB accumulates
phytoene (Figure 2.4A, peak 3). The transformants harboring both pACCRT-EB
and pUC-czPDS mainly produced ζ-carotene (Figure 2.4B, peak 1); the level of
residual phytoene was very low in these cells (Figure 2.4B, peak 3), suggesting
that most of the substrate was converted to ζ-carotene. The intermediate product
phytofluene was not observed (Figure 2.4B), possibly because it has a very short
life time as a consequence of rapid desaturation to form the end product of the
reaction. These results suggest that the C. zofingiensis PDS encodes a protein that
rapidly converts phytoene to ζ-carotene.
Figure 2.3 Schematic illustration of C. zofingiensis PDS cDNA (A) and its
correspondent gene (B). The coding region of PDS cDNA is indicated by a black
box; the promoter sequence is denoted by a gray box, the exons are showed by
black boxes and the introns are line-indicated.
68
Chpater 2. Isolation and characterization of the phytoene desaturase gene from Chlorella
zofingiensis
0.08
A
3
AU
0.06
0.04
0.02
0.00
18.00
0.006
19.00
B
20.00
21.00
22.00
23.00 24.00
Minutes
25.00
26.00
27.00
28.00
22.00
23.00 24.00
Minutes
25.00
26.00
27.00
28.00
22.00
23.00 24.00
Minutes
25.00
26.00
27.00
28.00
1
AU
0.004
0.002
3
0.000
18.00
19.00
C
20.00
21.00
1
AU
0.004
3
0.002
2
0.000
18.00
19.00
20.00
21.00
Figure 2.4 HPLC chromatogram of carotenoids extracted from E. coli cells
harboring plasmid pACCRT-EB (A), pUC-czPDS and pACCRT-EB (B) or
pUC-czPDS-L516R and pACCRT-EB (C). Absorbance was recorded at 305 nm.
Peaks are identified as follows: 1 ζ-carotene, 2 phytofluene, 3 phytoene.
69
Chpater 2. Isolation and characterization of the phytoene desaturase gene from Chlorella
zofingiensis
2.4.3 C. zofingiensis PDS gene is up-regulated by high light and
glucose
C. zofingiensis was found to accumulate ketocarotenoids in response to
high light (Del Campo et al., 2004) or high concentrations of glucose (Ip et al.,
2004; Ip & Chen, 2005). Whether the PDS gene is regulated by high light and
glucose is unknown and investigated here. Photoautotrophically grown cells in
exponential growth were either illuminated with high light (120 μmol photon m-2
s-1) or supplemented with glucose (50 mM) and maintained in the dark for 0 to 48
h. Basal expression of the PDS gene in control samples was observed (Figure
2.5A and B, lane 1). Basal expression of the PDS gene is required for the
biosynthesis of carotenoids that serve as antenna pigments for light harvest during
photosynthesis (Demmig-Adams & Adams, 1993). The level of PDS transcript
was elevated upon exposure to high light, reaching its highest levels
approximately 12 h following the high light exposure (Figure 2.5A). The
increased steady-state expression of the PDS gene, in combination with
expression of other genes of the carotenoid biosynthesis pathway, may contribute
to the enhanced biosynthesis of ketocarotenoids. Interestingly, glucose also caused
elevated PDS transcript accumulation (Figure 2.5B). The PDS expression was
drastically up-regulated and reached its maximum upon 24-h glucose induction
(Figure 2.5B, lane 3). Thereafter, the steady-state mRNA level began decreasing,
finally to almost the basal level (Figure 2.5B, lane 4 and 5). The carotenogenic
genes BKT and CHYb that are involved in astaxanthin biosynthesis were also
found to be up-regulated by glucose (Huang et al., 2006; Li et al., 2008b, 2009).
70
Chpater 2. Isolation and characterization of the phytoene desaturase gene from Chlorella
zofingiensis
Figure 2.5 Analysis of the C. zofingiensis PDS expression under high light (120
μmol m-2 s-1) (A) or supplemented with 50 mM glucose (B) for 0 h (lane1), 12 h
(lane 2), 24 h (lane 3), 36 h (lane 4) and 48 h (lane 5) using RT-PCR approach.
Amplified product was separated on a 2% agarose gel, along with the control
(actin) amplification (C).
2.4.4 PDS-L516R is resistant to the herbicide norflurazon
Unlike the crtI-type phytoene desaturase from bacteria and fungi, the
plant and algal PDS is the target of herbicides that inhibit the desaturation activity
of PDS. However, PDS with specific amino acid changes become resistant to the
herbicide norflurazon, as demonstrated in the cyanobacterium Synechococcus
PCC 7942, (Chamovitz et al., 1991; Chamovitz et al., 1993), plants (Arias et al.,
2006), and H. pluvialis (Steinbrenner & Sandmann, 2006). To determine whether
the Chlorella PDS could be modified to become herbicide resistant, site-directed
mutagenesis was used to generate an L516R change. Functional analysis of the
altered protein was carried out in E. coli. As expected, ζ-carotene was produced in
E. coli transformants carrying both pUC-czPDS-L516R and pACCRT-EB (Figure
2.4C). In addition, substantial amounts of the precursor phytoene and the
intermediate phytofluene were present in transformants harboring the mutated
PDS, but not in the tranformants with the unaltered protein (Figure 2.4C, peak 2
and 3). These data indicated that the mutation reduced the activity of phytoene
desaturase, which is consistent with previous studies (Chamovitz et al., 1993;
Michel et al., 2004).
The time course of the formation of phytofluene and ζ-carotene from
phytoene catalyzed by PDS was surveyed to optimize the incubation time for in
71
Chpater 2. Isolation and characterization of the phytoene desaturase gene from Chlorella
zofingiensis
vitro PDS assay. As indicated by Figure 2.6, both the unaltered PDS and
PDS-L516R enzymes produced almost the highest amount of ζ-carotene after
incubation of 6 h. Longer incubation time caused some degradation of the end
product formed. These data suggested that a 6-h incubation time is sufficient for
reliable quantitation of ζ-carotene and thus the PDS activity. PDS-L516R gave the
lower amount of ζ-carotene and concurrent higher amount of the intermediate
phytofluene as compared with the unaltered PDS (Figure 2.6), implying that this
mutated enzyme had an attenuated desaturation activity, which is consistent with
the results from functional analysis in E. coli (Figure 2.4).
Figure 2.6 Time course of the formation of phytofluene (circle) and ζ-carotene
(square) from phytoene catalyzed by the unaltered PDS (solid) and PDS-L516R
(open).
To explore the resistance of the mutated PDS to norflurazon, PDS-L516R
was assayed in vitro. In vitro enzymatic activities of the unaltered PDS and
PDS-L516R were measured by determining the conversion of phytoene to
ζ-carotene after the addition of different concentrations of norflurazon. The results
were expressed as units of activity relative to that of the control without the
herbicide. A plot of the reciprocal of PDS enzymatic activities versus the
concentration of the inhibitor norflurazon is shown in Figure 2.7. The
72
Chpater 2. Isolation and characterization of the phytoene desaturase gene from Chlorella
zofingiensis
concentrations for 50% inhibition (I50) of PDS desaturation activity were
calculated to be 0.064 μM for the unaltered PDS and 2.237 μM for PDS-L516R.
Compared with the unaltered PDS, the mutated one exhibited 35-fold higher
resistance to the bleaching herbicide norflurazon.
Figure 2.7 Plot of the reciprocal of in vitro PDS enzymatic activities versus
concentrations of the bleaching herbicide norflurazon for the E. coli-expressing
unaltered PDS (●) and PDS-L516R (■). RF stands for resistance factor.
Of the enzymes involved in the formation of carotenoids, PDS is the
primary target site of herbicides (Chamovitz et al., 1993; Michel et al., 2004).
Inhibition of this enzyme causes phytoene accumulation at the expense of colored
cyclic carotenoids. Thus in the presence of herbicides, there are not enough
carotenoids formed to ensure efficient photoprotection of the photosynthetic
apparatus. Chlorophylls therefore get degraded upon exposure of the cells to light,
leading to the typical bleaching phenotype and subsequent deaths. In addition to
causing herbicide resistance, the PDS mutations also reduce the enzymatic
activity of phytoene desaturation to varying extents (Chamovitz et al., 1993;
Michel et al., 2004). Since the herbicides do not compete for the binding of
73
Chpater 2. Isolation and characterization of the phytoene desaturase gene from Chlorella
zofingiensis
phytoene on PDS (Sandmann et al., 1989), it is speculated that the binding sites
for the inhibitors and the substrate are either overlapping or in close proximity to
each other (Chamovitz et al., 1993). The exact mechanism of herbicide inhibition,
however, remains incompletely understood. Recently it was demonstrated that
herbicides compete with cofactors of PDS for a binding site in the PDS
polypeptide (Breitenbach et al., 2001). This binding site, in the PDS mutants,
might be modified and thereby hinders the interaction with herbicides. The
modification may also hinder the interaction of PDS with cofactors, resulting in a
reduction in the efficiency of the phytoene desaturation.
In conclusion, the PDS gene from C. zofingiensis encodes a polypeptide
with high enzymatic activity in converting phytoene to ζ-carotene and was shown
to be up-regulated by high light and glucose treatment. The modified PDS-L516R
exhibited much higher resistance to the bleaching herbicide norflurazon than the
unaltered PDS. The results suggested that the PDS-516R gene might be a useful
tool for genetic engineering of carotenoid biosynthesis in C. zofingiensis because
of its algal and endogenous origin and, more generally, may serve as a selectable
marker for the introduction of endogenous DNA into algal genomes.
74
Chpater 3. Isolation and characterization of Chlorella zofingiensis mutants with enhanced
biosynthesis of astaxanthin
Chapter 3
Isolation and characterization of Chlorella zofingiensis
mutants with enhanced biosynthesis of astaxanthin
3.1 Abstract
The green alga Chlorella zofingiensis may serve as a new source of
natural astaxanthin if its astaxanthin content gets increased. This chapter reported
an effective system for generating and isolating astaxanthin-rich C. zofingiensis
mutants. Mutations were generated by the employment of chemical mutagen
MNNG (45 μg mL-1) or EMS (0.36 M) which gave an around 10% survival rate.
Target mutants were isolated by screening the treated cells with 0.5 μM
norflurazon. Over two hundreds mutants were obtained, from which five were
selected for further characterization. No significant differences in cell growth and
pigment profile were found between the mutants and wild type (WT) cells when
the culture medium contained no herbicide. In contrast to WT cells which were
bleached by 0.25 μM norflurazon, all the mutants grew well and synthesized
astaxanthin even when the culture medium contained up to 1 μM norflurazon.
Furthermore, mutants accumulated up to 54% more astaxanthin than WT.
Coordinately, higher transcript levels of carotenoid ketolase (BKT) and carotenoid
hydroxylase (CHYb) genes were found in the mutant cells. Interestingly two of
the five mutants also accumulated higher amounts of total carotenoids (TC).
These results indicates that C. zofingiensis and possibly other green algae can be
genetically modified by mutagenesis followed by specific herbicide screening for
enhanced production of carotenoids including the high-value ketocarotenoid
astaxanthin.
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Chpater 3. Isolation and characterization of Chlorella zofingiensis mutants with enhanced
biosynthesis of astaxanthin
3.2 Introduction
The major constraint of C. zofingiensis as commercial source of natural
astaxanthin is its low cellular content of astaxanthin. One strategy to overcome
this problem is to genetically modify the key enzymes involved in the carotenoid
biosynthesis. Mutagenesis has been found to be an effective approach to enhance
the biosynthesis of carotenoids including astaxanthin in various algal species,
especially in H. pluvialis (Tjahjono et al., 1994; Zhang & Lee, 1997; Chen et al.,
2003; Ishikawa et al., 2004; Hu et al., 2008a; Sandesh Kamath et al., 2008).
Commonly chemical mutagen (e.g., EMS or MNNG) or UV is used to generate
mutants from which target mutants (e.g., cells with higher contents of certain
carotenoids) are selected by using specific inhibitors to carotenogenic enzymes.
While a number of astaxanthin hyper-producing mutants of H. pluvialis have been
obtained, no mutants of other astaxanthin-producing green algae are available. In
addition, there is little information on the mechanisms of metabolic processes of
such mutants. This chapter described for the first time the generation and selection
of C. zofingiensis mutants with over-production of astaxanthin through chemical
mutagenesis followed by herbicide screening. Furthermore, the expression of
carotenogenic genes including phytoene desaturase (PDS), carotenoid ketolase
(BKT) and carotenoid hydroxylase (CHYb) were investigated among WT and
mutants.
3.3 Methods and materials
3.3.1 Algal strain and culture conditions
The maintenance and inoculation of C. zofingiensis were described in
2.3.1.
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Chpater 3. Isolation and characterization of Chlorella zofingiensis mutants with enhanced
biosynthesis of astaxanthin
3.3.2 Mutagenesis
Cells from logarithmic suspension culture were harvested, washed twice
with distilled water and re-suspended in 0.1 M phosphate buffer (pH 7.0) at a
concentration of 108 cells mL-1 for mutagenesis. The cells mentioned above were
treated with ethyl methanesulphonate (EMS, Sigma) in the concentration range of
0 to 0.48 M for 30 min or with N-methyl-N′-nitro-N-nitrosoguanidine (MNNG,
Sigma) at 0 to 60 μg L-1 concentration for 60 min.
3.3.4 Isolation of Chlorella mutants
After
the treatment
of chemical mutagens,
equal volume of
filter-sterilized sodium thiosulfate (5%) was added to terminate the mutagenesis.
All the treated cells were washed twice with distilled water, re-suspended in Kuhl
medium and allowed to grow for 24 h (Hirschberg et al., 1987). For selection, the
mutagenized cells were incubated on Kuhl agar plates containing 0.5 μM
norflurazon at 25 °C under continuous illumination of 25 µmol photon m-2 s-1. The
colonies showing up on the selection plates were picked up based on the colony
characteristics and color and transferred individually into Kuhl liquid medium
containing 0.5 μM norflurazon for cultivation.
3.3.5 Cell dry weight determination
The cultures of WT and mutants were centrifuged at 3,800 g for 5 min.
The pellet was washed three times with distilled water and filtered through a
pre-dried Whatman GF/C filter paper (1.2 μm pore size). The algal cells on the
filter paper discs were dried at 70 °C in a vacuum oven until constant weight and
were cooled down to room temperature in a desiccator before weighting.
77
Chpater 3. Isolation and characterization of Chlorella zofingiensis mutants with enhanced
biosynthesis of astaxanthin
3.3.6 Astaxanthin induction
The cultures of WT and mutants were first grown in Kuhl medium for 4
days. To induce astaxanthin accumulation, the cultures mentioned above were
inoculated into the fresh medium containing 30 g L-1 glucose and cultured in the
absence of light.
3.3.7 Extraction and analysis of pigments
Cell samples were collected by centrifugation and then freeze-dried on a
DW3 freeze-drier (Heto Dry Winner, Denmark). Extraction was carried out with
acetone and liquid nitrogen until the cell debris was almost colorless. The extracts
were filtered through a 0.22 μm Millipore organic membrane. HPLC analysis of
the pigments was described in 2.3.7.
3.3.8 RNA isolation and RT-PCR assay
RNA isolated and reverse transcription were described in 2.3.5. PCR
amplification was carried out using specific primers of PDS, BKT and CHYb
(Table 3.1). C. zofingiensis actin (ACT) primers (Table 3.1) were used to
demonstrate equal amounts of templates and loading. The GenBank accession
numbers for PDS, BKT and CHYb were EF621406, AY772713 and EU016205,
respectively. Amplification was done by conventional PCR [94 °C for 2 m
followed by 24 cycles (for PDS, BKT and ACT genes) or 26 cycles (for CHYb
gene) of 94 °C for 15 s, 58 °C for 20 s, 72 °C for 30 s].
78
Chpater 3. Isolation and characterization of Chlorella zofingiensis mutants with enhanced
biosynthesis of astaxanthin
Table 3.1 Primer sets for gene expression by RT-PCR
Primer (5′-3′)
Gene
PDS
Forward
Reverse
BKT
Forward
Reverse
CHYb
Forward
Reverse
ACT
Forward
Reverse
GATGAATGTATTTGCTGAACTGGGC
GGCCAGTGCCTTAGCCATAGCG
GTGCTCAAAGTGGGGTGGTATG
CCATTTCCCACATATTGCACCT
GCCAGCCATGAAACGTGTG
GTTCCTTCCAGTTATGTACACA
TGCCGAGCGTGAAATTGTGAG
CGTGAATGCCAGCAGCCTCCA
3.4 Results
3.4.1 Carotenoid biosynthesis blocked by norflurazon
C. zofingiensis cells exhibited green phenotype under standard growth
conditions. Exposed to high light irradiation or induced by glucose,
ketocarotenoids mainly astaxanthin and canthaxanthin accumulated within cells,
rendering the cells an orange-red color (Orosa et al., 2000, 2001; Del Campo et al.,
2004; Ip et al., 2004; Ip & Chen, 2005; Li et al., 2009). Norflurazon can
specifically bind to PDS and block phytoene desaturation, giving rise to phytoene
accumulation at an expense of colored cyclic carotenoids (Chamovitz et al., 1993).
Without sufficient carotenoids formed to ensure efficient photoprotection of
photosynthetic apparatus, chlorophylls get degraded upon exposure of cells to
light, leading to the typical bleaching phenotype and subsequent death of cells.
Various concentrations of norflurazon were employed to examine its effect on
carotenoid formation in C. zofingiensis cells. As shown in Figure 3.1, TC content
79
Chpater 3. Isolation and characterization of Chlorella zofingiensis mutants with enhanced
biosynthesis of astaxanthin
in the algal cells exhibited high sensitivity to noflurazon. In the presence of 0.1
μM norflurazon, TC content reduced from 1.685 mg g-1 (control value of culture
with no herbicide) to 0.336 mg g-1. Higher concentrations of norflurazon (0.25 μM,
0.5 μM, and 1 μM) resulted in further lower TC contents, and colorless phytoene
was found to accumulate in these norflurazon-treated cells (data not shown).
Figure 3.1 Effect of norflurazon on TC accumulation in C. zofingiensis cells. Cell
samples were harvested after 4-day cultivation under standard growth condition
supplemented with various concentrations of norflurazon.
3.4.2 Isolation of Chlorella mutants resistant to norflurazon
Appropriate levels of MNNG and EMS were determined by treating C.
zofingiensis cells in various concentrations. The survival rate of C. zofingiensis
treated with MNNG decreased rapidly and reached 11.4% when the concentration
of MNNG increased to 45 μg mL-1 (Figure 3.2A). EMS also reduced the survival
rate in a concentration-dependent manner and gave 9.6% survival rate at 0.36 M
(Figure 3.2B).
The algal cells treated with 45 μg mL-1 MNNG for 1 h or 0.36 M EMS
80
Chpater 3. Isolation and characterization of Chlorella zofingiensis mutants with enhanced
biosynthesis of astaxanthin
Figure 3.2 Effect of MNNG (A) and EMS (B) treatments on the survival rate of
C. zofingiensis cells. The survival rate of each treatment was counted by
comparing with control cells (100% survival).
for 30 min were selected further by spreading onto solid media supplemented with
0.5 μM norflurazon. Green colonies were observed after 2 to 3 weeks of
incubation. More than 100 colonies for each treatment were obtained. Mutants
that were found to show apparent deeper orange color were selected for further
investigation. The selected colonies were transferred into liquid medium
containing 0.5 μM norflurazon and allowed to grow under standard growth
conditions. To eliminate negative effects of herbicide on algal cells, the survived
colonies were inoculated into norflurazon free medium twice for further studies.
81
Chpater 3. Isolation and characterization of Chlorella zofingiensis mutants with enhanced
biosynthesis of astaxanthin
3.4.3 Growth and astaxanthin accumulation of mutants and WT
Under standard growth conditions (without norflurazon), all mutants
showed a similar growth pattern to WT and no significant difference of cell
biomass was observed among WT and mutants (data not shown). Norflurazon
greatly limited the cell growth of WT in a concentration dependent manner
(Figure 3.3A). In contrast, mutants exhibited strong resistance to the herbicide. As
shown in Figure 3.3A, 0.25 μM norflurazon drastically reduced the cell biomass
of WT from 4.64 g L-1 (control value of culture with no herbicide) to 2.82 g L-1
but only slightly affected the cell biomass of mutants. Even in the presence of 1
μM norfluraon, the cell biomass and chlorophyll content of the mutants was only
moderately reduced to 3.54 g L-1 and 2.25 mg g-1 respectively (for E17, Figure
3.3A and Table 3.2). WT cells were totally bleached in the presence of 1 μM
norflurazon, correlated with the drastic decrease of chlorophyll content (Table
3.2).
Table 3.2 Chlorophyll content of C. zofingiensis WT and mutants after 4-day
cultivation with or without norflurazon
Chlorophyll content (mg g-1)
Strains
Norflurazon free
1 μM norflurazon
WT
6.51 ± 0.46
0.41 ± 0.04
M7
5.89 ± 0.53
2.32 ± 0.19
M87
6.24 ± 0.39
2.47 ± 0.28
E5
5.66 ± 0.52
2.11 ± 0.23
E17
6.11 ± 0.33
2.25 ± 0.17
E65
6.03 ± 0.41
2.20 ± 0.35
Norflurazon also exerted a negative effect on astaxanthin formation in
algal cells of both WT and mutants. However, astaxanthin accumulation in WT
82
Chpater 3. Isolation and characterization of Chlorella zofingiensis mutants with enhanced
biosynthesis of astaxanthin
cells was much more sensitive to norflurazon than that in mutants. As illustrated
by Figure 3.3B, in the presence of 0.1 μM norflurazon, astaxanthin content in WT
cells was greatly reduced from 0.235 mg g-1 to 0.014 mg g-1, while in mutants the
astaxanthin contents were only slightly reduced (from 0.273 mg g-1 to 0.212 mg
g-1). Inhibitory concentration that inhibits astaxanthin formation by 50% (I50) was
calibrated by using Dixon plot as [norflurazon (μM)] versus [astaxanthin content
(mg g-1)]-1. The norflurazon-resistant mutants obtained in this study exhibited 21to 26- fold higher resistance than WT (Table 3.3).
Figure 3.3 Effect of norflurazon concentrations on cell biomass (A) and
astaxanthin contents (B) of mutants. Cell samples were harvested after 4-day
cultivation under standard growth condition.
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Chpater 3. Isolation and characterization of Chlorella zofingiensis mutants with enhanced
biosynthesis of astaxanthin
Table 3.3 Half-maximum inhibitory concentration (I50) of norflurazon on
astaxanthin formation in C. zofingiensis WT and mutants
Strains
I50 (μM)
WT
0.021
M7
0.485
M87
0.532
E5
0.448
E17
0.544
E65
0.525
3.4.4 Accumulation of TC and astaxanthin induced by glucose
The algal cells of WT and mutants were first allowed to grow for 4 days
under standard growth conditions, followed by glucose induction for 5 days. The
induced cells were harvested and freeze-dried for pigment extraction and HPLC
analysis. M7, M87 and E5 produced higher amounts of astaxanthin, while the TC
contents of these three mutants were comparable to the WT (Figure 3.4A). The
astaxanthin contents in M7, M87 and E5 were 0.661 mg g-1, 0.65 mg g-1 and 0.621
mg g-1, 48.5%, 46.1% and 39.6% greater than that in the WT (0.445 mg g-1),
respectively. In contrast, E17 and E65 could accumulate greater amounts of both
astaxanthin and TC (Figure 3.4A). Among the mutants investigated in the present
study, E17 had the ability to accumulate highest amounts of TC and astaxanthin,
which were 2.277 mg g-1and 0.685 mg g-1, 30.1% and 53.9% higher than WT,
respectively. Correlated with the higher astaxanthin content, the mutant cells
exhibited deeper yellow color than the WT (Figure 3.4B).
84
Chpater 3. Isolation and characterization of Chlorella zofingiensis mutants with enhanced
biosynthesis of astaxanthin
Figure 3.4 Comparison of WT and mutants cultured with 30 g L-1 glucose in the
dark. (A) TC and astaxanthin accumulated in WT and mutants. (B) Algal cells of
WT and mutants. Cell samples were harvested after glucose induction for 5 days.
Gray and dark columns indicate TC and astaxanthin respectively. Data marked
with the same letters of each total carotenoid content and astaxanthin are not
significantly different (P>0.05).
85
Chpater 3. Isolation and characterization of Chlorella zofingiensis mutants with enhanced
biosynthesis of astaxanthin
3.4.5 Expression analysis of carotenogenic genes
Three genes including PDS, BKT and CHYb that are associated with
general carotenogenesis and specific astaxanthin biosynthesis were selected for
expression analysis. The transcript levels of these genes in WT and mutants were
quantified by RT-PCR and compared. Algal cells induced with glucose for 48 h
were employed. All mutants exhibited higher transcription levels of BKT and
CHYb than WT while the increasing extent differed (Figure 3.5), which was well
consistent with the greater amount of astaxanthin produced by mutants. No
significant difference in PDS transcript, however, was observed between WT and
mutants (M7, M87 and E5) (Figure 3.5). Even in E17 and E65 that produced
notable higher amounts of TC, the PDS expression was merely comparable to WT.
Figure 3.5 Expresson of carotenoid biosynthetic genes in cells of WT and mutants.
Cell samples were induced for 2 days with glucose in the dark. The PCR
amplified products were separated on a 2% agarose gel, along with the control
(ACT) amplification.
3.5 Discussion
In this chapter, chemical mutagens MNNG and EMS were employed to
mutate C. zofingiensis cells for enhanced astaxanthin production. Various
concentrations of the mutagens were surveyed to obtain satisfactory survival rates
86
Chpater 3. Isolation and characterization of Chlorella zofingiensis mutants with enhanced
biosynthesis of astaxanthin
of the algal cells. Cells treated with 45 μg mL-1 MNNG or 0.36 M EMS giving
approximately 10 % survival rate were found practicable to obtain sufficient
mutants from which astaxanthin over-production mutants were selected by
screening with 0.5 μM norflurazon.
Herbicides are a group of diverse chemical compounds with the ability of
disturbing basic metabolic processes essential for algal cells. They have been used
effectively to screen mutants with desired properties, including but not restricted
to herbicide resistance (Hirschberg et al., 1987; Linden et al., 1990; Tjahjono et al.,
1994), enhanced accumulation of carotenoids (Tripathi et al., 2001; Chen et al.,
2003; Sandesh Kamath et al., 2008). Astaxanthin-hyperproducing mutants of H.
pluvilias have been isolated by using chemical mutagenesis followed by screening
with inhibitors of the carotenoid biosynthesis (e.g., diphenylamine, compactin or
nicotine) (Tripathi et al., 2001; Chen et al., 2003). In this chapter, over 200
norflurazon-resistant mutants of C. zofingiensis were obtained, of which 5 mutants
were further investigated. These mutants accumulated higher amounts of
astaxanthin than WT under high light irradiation (Figure 3.4). In addition to
astaxanthin, the EMS-treated mutants E17 and E65 also produced a greater
amount of TC (Figure 3.4A). The enhanced astaxanthin production in the mutants
might be related to the increased expression of both BKT and CHYb genes, based
on the RT-PCR analysis results (Figure 3.5), since astaxanthin formation was
previously shown to be related to the transcript levels of carotenogenic genes (Li
et al., 2008, 2009).
Although
the
EMS-treated
Synechococcus
sp.
mutants
and
MNNG-induced H. pluvialis mutants were suggested to be related to a single-gene
mutation (Linden et al., 1990; Hu et al., 2008a), it remains unknown at this point
whether the five C. zofingiensis mutants obtained in the present study were the
result of a single- or multi-gene mutation. A point mutation might occur in the
PDS gene of the C. zofingiensis mutants, resulting in resistance to the bleaching
herbicide norflurazon (Chamovitz et al., 1993). In addition, it would be likely that
a key regulatory gene involved in the biosynthesis of carotenoids also got mutated,
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Chpater 3. Isolation and characterization of Chlorella zofingiensis mutants with enhanced
biosynthesis of astaxanthin
therefore redirecting the carotenoid flux to astaxanthin and promoting astaxanthin
accumulation in the C. zofingiensis mutants. The underlying mechanism
responsible for the enhanced accumulation of astaxanthin in mutants remains to
be elucidated and further studies will be described in chapter 4.
Astaxanthin has been widely used in aquaculture for pigmentation of
many marine animals such as salmon and trout. Around 100, 000 kg of
astaxanthin per year is demanded, with a price estimated at US$200 million
(Lorenz & Cysewski, 2000). Essentially, today’s commercial astaxanthin for
aquaculture is almost produced by chemical synthesis, but consumer’s growing
need using natural ingredients in all forms of food nutrients provides a good
chance for natural astaxanthin production by green microalgae. H. pluvialis is the
most promising source of astaxanthin for aquaculture in that it has the highest
cellular content of astaxanthin and no further purification of the pigment is
required (Ausich, 1997). However, its slow growth rate, low cell yield and
susceptibility to contamination hinder its commercial application (Olaizola, 2000;
Hata et al., 2001; Ip et al., 2004). Furthermore, light is required for enhancing
astaxanthin production by H. pluvialis cells (Orosa et al., 2000; Hata et al., 2001),
adding cost to commercialization of algal products. Recently C. zofingienesis has
drawn much attention and been proposed as an alternative promising producer of
astaxanthin due to its fast growth and ability to accumulate astaxanthin in the dark,
suggesting that it might be well developed for producing astaxanthin in industrial
fermentation systems (Orosa et al., 2000; Ip et al., 2004; Ip & Chen, 2005). Very
high biomass (53 g L-1) of C. zofingienesis has been achieved by using a fed-batch
fermentation strategy (Sun et al., 2008). Thus, the low cellular astaxanthin content
of C. zofingiensis is the major limitation to its commercial use as natural
astaxanthin. The E17 has the ability to accumulate approximately 54% more
cellular astaxanthin than the WT. The mutants described in this study may be
further modified for enhanced production of astaxanthin by a similar approach
used in this study but selecting by using other herbicides (e.g., nicotine or
diphenylamine) that target different carotenogenic enzymes. In addition, the
88
Chpater 3. Isolation and characterization of Chlorella zofingiensis mutants with enhanced
biosynthesis of astaxanthin
mutants
also
provide
important
materials
for
the
structure-function relationship of the membrane PDS enzyme.
89
investigation
of
Chpater 4. Molecular characterization of the Chlorella zofingiensis mutant E17
Chapter 4
Molecular characterization of the Chlorella zofingiensis
mutant E17
4.1 Abstract
A stable Chlorella zofingiensis mutant (E17) resistant to the herbicide
norflurazon was characterized with respect to growth, astaxanthin biosynthesis,
and phytoene desaturation. The mutant E17 could grow well and produced normal
levels of colored carotenoids in the presence of 0.25 μM norflurazon, in which the
growth of WT cells was greatly limited due to inhibited carotenoid formation.
Induced by high-light irradiation or glucose, E17 produced 44% or 36% more
astaxanthin than WT when cultured in media without norflurazon. A point
mutation (C to T) was revealed to occur in the PDS gene of E17, leading to an
amino acid change (L516F) in its coding region. The mutated PDS exhibited
31-fold resistance to norflurazon when compared to WT as determined by an in
vitro assay. Surprisingly, the mutated PDS exhibited higher efficiency in
converting phytoene to ζ-carotene. No difference in PDS transcripts was found
between E17 and WT cells cultured either in normal or induced conditions. In
contrast, higher transcript levels of BKT and CHYb were found in E17 cells. These
results suggest that a point mutation in Chlorella PDS gene results in an enzyme
with lower affinity to norflurazon but higher efficiency in converting phytoene to
ζ-carotene. As a consequence, E17 is resistant against norflurazon and synthesizes
higher amounts of carotenoids including astaxanthin.
90
Chpater 4. Molecular characterization of the Chlorella zofingiensis mutant E17
4.2 Introduction
A number of norflurazon-resistant mutants of C. zofingiensis by chemical
mutagenesis were generated in chapter 3. One of them, named as E17, is a stable
mutant that had a higher astaxanthin content than the WT. Norflurazon inhibits the
activity of plant-type phytoene desaturase (PDS), a key enzyme involved in
catalyzing the rate-limiting step of carotenoid biosynthesis (Chamovitz et al.,
1993). Mutants resistant to norflurazon may result from the increased amount of
PDS gene product or from structural change of PDS that lowers its affinity to the
herbicide (Chamovitz et al., 1991, 1993; Michel et al., 2004). In this chapter, the
characterization of E17 was carried out, with respect to its growth,
carotenogenesis and PDS activity. This mutated PDS is the first example for a
point mutation in PDS which not only makes the alga resistant to the herbicide
norflurazon, but also enhances the biosynthesis of total carotenoids (TC)
including astaxanthin.
4.3 Methods and materials
4.3.1 Algal strain and culture conditions
The maintenance and inoculation of C. zofingiensis were described in
2.3.1. For herbicide treatment, the cell cultures were grown in Kuhl medium for 4
days and then inoculated into the same medium containing various concentrations
of norflurazon (Sigma), at an inoculum of 10% (by volume, average cell
concentration of 0.5 g L-1). For induction of astaxanthin biosynthesis, the 4-day
cultures mentioned above were exposed to continuous illumination of 250 µmol
photon m-2 s-1 or inoculated into fresh Kuhl medium containing 30 g L-1 glucose
and maintained in the dark.
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Chpater 4. Molecular characterization of the Chlorella zofingiensis mutant E17
4.3.2 Cell dry weight determination
Cell dry weight determination was described in 3.3.5.
4.3.3 Extraction and analysis of pigments
The extraction and pigment analysis were described in 3.3.7.
4.3.4 Chlorophyll fluorescence measurement
Cells were harvested by centrifugation and re-suspended in fresh Kuhl
medium. Chlorophyll fluorescence was measured using a Multi-Mode
Chlorophyll Fluorometer (Opti-Sciences, Hudson, USA) at room temperature.
Non-photochemical quenching (NPQ) was calculated using (Fm-Fm′)/Fm′, where
Fm is the maximum fluorescence in the dark-adapted state and Fm′ is the
maximum fluorescence in the light-adapted state.
4.3.5 RNA isolation and RT-PCR assay
RNA isolation and RT-PCR assay were described in 3.3.8.
4.3.6 PDS expression in E. coli
The PDS cDNA from E17 was amplified, sequenced and inserted into
pUC18 (Stratagene) as an in-frame fusion to the lacZ gene, resulting in plasmid
pUC-czPDS-L516F (a leucine codon at position 516 changed to a Phenylalanine
codon). This plasmid and pUC-czPDS constructed in chapter 2 were introduced
into E. coli JM109 for PDS enzyme expression. The E. coli cells were grown in
92
Chpater 4. Molecular characterization of the Chlorella zofingiensis mutant E17
SOB medium containing 50 μg mL-1 ampicillin at 37 °C with vigorous shaking,
and 1 mM IPTG was added when the optical density at 600 nm reached 0.5. After
a 5-h induction period the cells were harvested for preparation of crude PDS
proteins.
4.3.7 Enzyme and substrate preparation and in vitro PDS assay
The preparation of enzyme and substrate and the in vitro PDS assay were
described in 2.3.6 and 2.3.7, respectively.
4.4 Results
4.4.1 The growth and carotenogensis of E17
Chapter 3 described the treatment of C. zofingiensis cells with chemical
mutagen EMS followed by selection with the bleaching herbicide norflurazon. A
number of mutants resistant to norflurazon were obtained and analyzed for their
astaxanthin production. E17, a stable mutant, was found to produce much more
astaxanthin. E17 showed no loss of resistance to norflurazon after over 100 times
of subculture without norflurazon selection (data not shown). E17 could grow
well and produce nearly normal levels of total carotenoids in the presence of 0.25
µM norflurazon, in which the growth of wild type (WT) cells was greatly limited
due to total blocking of carotenoid formation. Norflurazon suppressed
carotenogenesis leading to a decreased formatio of TC including astaxanthin in
both WT and E17 cells (Figure 4.1). WT cells showed high sensitivity to
norflurazon as indicated by a decreased TC content (from 1.69 mg g-1 to 0.83 mg
g-1) in the present of 0.05 μM norflurazon. Higher concentrations of norflurazon
resulted in much lower amounts of TC (Figure 4.1). In addition, the formation of
astaxanthin in WT almost ceased when the cultures were treated with up to 0.25
93
Chpater 4. Molecular characterization of the Chlorella zofingiensis mutant E17
μM norflurazon. In contrast, E17 synthesized significant amounts of TC and
astaxanthin even at 0.5 μM norflurazon, a lethal concentration for the WT. Values
of half-maximum inhibitory concentration (I50) of norflurazon on cell biomass, TC
and astaxanthin formation in WT and E17 was shown in Table 4.1. E17 exhibited
7.1, 22.7 and 25.9 -fold higher resistance to norflurazon than WT, with respect to
cell biomass, TC accumulation and astaxanthin formation respectively.
Figure 4.1 Effect of norflurazon concentration on TC and astaxanthin contents of
WT and E17. (□) WT TC; (■) WT astaxanthin; (○) E17 TC; (●) E17 astaxanthin.
Cells were harvested from 4-day culture under standard growth conditions.
Table 4.1 Half-maximum inhibitory concentrations (I50) of norflurazon on cell
biomass, TC and astaxanthin formation of WT and E17. I50 was calibrated by
using Dixon plot as [norflurazon (μM)] versus [biomass (g L-1)], [TC content (mg
g-1)] or [astaxanthin content (mg g-1)].
I50 (μM)
Strain
Cell biomass
TC
WT
0.395
0.034
0.021
E17
2.802
0.702
0.544
94
Astaxanthin
Chpater 4. Molecular characterization of the Chlorella zofingiensis mutant E17
4.4.2 Enhanced production of TC and astaxanthin by E17 under
high light stress or glucose induction
High light irradiation was reported to trigger ketocarotenogensis in C.
zofingiensis cells (Orosa et al., 2000, 2001; Del Campo et al., 2004; Li et al.,
2009). Here a comparative study of WT and E17, in terms of TC and astaxanthin
contents under high light irradiation, was conducted. Consistent with previous
reports, C. zofingiensis WT and E17 synthesized much higher amounts of TC and
astaxanthin under high light than normal light (Figure 4.2A). E17 produced
apparently greater amounts of both TC and astaxanthin than WT during the whole
illumination period (Figure 4.2A). The carotenoid contents of WT and E17 cells
exposed to high light for 24 h are shown in Table 4.2. Lutein was the major
carotenoid, followed by zeaxanthin and astaxanthin. Significant higher amounts of
β-carotene, lutein, zeaxanthin, as well as the ketocarotenoids adonixanthin,
canthaxanthin and astaxanthin were found to accumulate in E17 cells. The TC and
astaxanthin contents (2.96 mg g-1 and 0.38 mg g-1) of E17 were 27.9 % and 43.7%
higher than that of WT respectively. The zeaxanthin content and the
zeaxanthin/(zeaxanthin + antheraxanthin + violaxanthin) ratio, two factors related
to the degree of NPQ, were both increased in E17 when compared with WT
(Table 4.2). In accordance with the higher zeaxanthin content, E17 had a greater
NPQ value (Figure 4.3). Both WT and E17 reached their maxima at 105 s and the
NPQ value of E17 was 33.3% higher than that of WT.
In addition to light, glucose is another inducer of astaxanthin biosynthesis
in C. zofingiensis cells (Ip et al., 2004). With glucose as the sole carbon and
energy source, C. zofingiensis grew fast and synthesized astaxanthin (Ip & Chen,
2005; Sun et al., 2008). The time course of TC and astaxanthin accumulation in
heterotrophic WT and E17 cells cultured in Kuhl medium with 30 g L-1 is shown
in Figure 4.2B. Compared with the non-induced cells on day 0, the TC and
astaxanthin contents of WT were slightly decreased on day 2. Thereafter both TC
95
Chpater 4. Molecular characterization of the Chlorella zofingiensis mutant E17
and astaxanthin increased and the astaxanthin content reached its maximum in 14
days. E17 exhibited the same induction pattern of TC and astaxanthin as WT.
However, E17 produced apparently greater amounts of carotenoids, for example
on day 6 the TC and astaxanthin contents were 2.50 mg g-1and 0.85 mg g-1, 18%
and 36% higher than that of WT respectively. Even on day 10 the astaxanthin
content of WT was slightly lower than that of E17 on day 6. It is worth noting that
after day 6 the difference in TC and astaxanthin contents between WT and E17
became smaller. In correlation with the earlier enhanced astaxanthin accumulation,
E17 exhibited an earlier change from green to orange (data not shown).
Figure 4.2 Accumulation of TC and astaxanthin in WT and E17 cells under
continuous illumination of high light (A) or under induction of 30 g L-1 glucose in
the dark (B). (□) WT TC; (■) WT astaxanthin; (○) E17 TC; (●) E17 astaxanthin.
96
Chpater 4. Molecular characterization of the Chlorella zofingiensis mutant E17
Table 4.2 Pigment contents of green cells of WT and E17 exposed to high light for 24 h a
Carotenoid composition (μg g-1)
TC (mg
g-1)
Chlorophyll
(mg g-1)
Zea / (vio +
Neo
Vio
Ant
Ast
Ado
Lut
Zea
Can
β-Car
Vio + ant +
ant + zea)
zea
WT
2.31 ± 0.09
6.23 ± 0.21
64 ± 2.1
38 ± 1.3
67 ± 3.1
261 ± 11
219 ± 8
1132 ± 48 300 ± 11
128 ± 4
101 ± 5
405
0.74
E17
2.96 ± 0.12
6.05 ± 0.27
75 ± 3.4
32 ± 1.5
64 ± 1.8
375 ± 15
297 ± 13 1397 ± 55 414 ± 17
160 ± 6
121 ± 3
510
0.81
a
Data were expressed as the mean of three independent experiments. Neo, neoxanthin; Vio, violaxanthin; Ant, antheraxanthin; Ast, astaxanthin;
Ado, adonixanthin; Lut, lutein; Zea, zeaxanthin; Can, canthaxanthin; β-Car, β-carotene
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Chpater 4. Molecular characterization of the Chlorella zofingiensis mutant E17
Figure 4.3 Kinetics of NPQ of dark-adapted WT (■) and E17 (●) measured in
24-h high light treated cells.
4.4.3 Characterization of E17 PDS
Norflurazon can be coped with by increased amounts of the target
enzyme or by a mutation that affects the binding properties. The sequencing of the
PDS gene from E17 revealed a single point mutation (C to T), which resulted in
an amino acid change from leucine (L) to phenylalanine (F) at codon 516 (from
CTC to TTC). This L residue is located in a conserved motif of the PDS
polypeptides. Another substitution of this L by arginine (R) was earlier revealed
by us to confer herbicide resistance on Chlorella PDS (Huang et al., 2008). To
verify whether PDS-L516F is related to norflurazon resistance, the in vitro assays
of the Chlorella PDS and PDS-L516F were carried out. In vitro desaturation
activities of the unaltered PDS and PDS-L516F were measured by determining
the conversion of phytoene to ζ-carotene in the presence of various concentrations
of norflurazon. The results were expressed as units of activity relative to that of
the control without the herbicide. A Dixon plot of the reciprocal of PDS
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Chpater 4. Molecular characterization of the Chlorella zofingiensis mutant E17
desaturation activity versus the concentration of norflurazon is shown in Figure
4.4. The concentrations for 50% inhibition (I50) of PDS desaturation activity were
calculated to be 0.072 μM for WT PDS and 2.242 μM for PDS-L516F,
respectively. Compared with the WT PDS, the mutated one showed 31-fold
greater resistance to the inhibitor norflurazon. Surprisingly, PDS-L516F exhibited
higher activity in converting phytoene to ζ-carotene than its unaltered counterpart
as well as the PDS-L516R that was shown to have attenuated desaturation activity
(Chamovitz et al., 1993; Steinbrenner & Sandmann, 2006; Huang et al., 2008). As
shown in Table 4.3, the specific activity of PDS-L516F was 0.0315 μg ζ-carotene
formed (mg protein)-1 h-1, about 29% and 50% higher than that of the unaltered
PDS and PDS-L516R, respectively.
Figure 4.5 Plot of the reciprocal of in vitro phytoene desaturase activities versus
concentrations of norflurazon for the E. coli-based recombinant unaltered PDS (●)
and PDS-L516F (■).
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Chpater 4. Molecular characterization of the Chlorella zofingiensis mutant E17
Table 4.3 In vitro phytoene desaturation activity of E. coli-epxressing PDS
proteins.
Crude enzymes
Specific activity
[μg ζ-carotene formed (mg protein)-1 h-1]
PDS
0.0245±0.0015
PDS-L516R
0.0193±0.0012
PDS-L516F
0.0315±0.0021
4.4.4 Transcription analysis of PDS, BKT and CHYb genes
It was earlier reported that the enhanced biosynthesis of carotenoids
including astaxanthin in C. zofingiensis by high light or glucose induction was
correlated to the increased transcript levels of carotenogenic genes (Li et al.,
2008b; Li et al., 2009). To further investigate whether the enhanced production of
TC and astaxanthin by E17 was attributed to the up-regulation of PDS and/or
other carotenogenic genes, the quantification of the transcript levels of PDS
together with BKT and CHYb, two enzymes directly involved in astaxanthin
formation from β-carotene, was performed. RT-PCR result showed that no
apparent difference in PDS transcripts was perceived between WT and E17 either
under standard growth conditions or induced conditions (Figure 4.5). In contrast,
higher transcript levels of BKT and CHYb were found in E17 cells in response to
1-day high light treatment or induced by high concentration of glucose (30 g L-1)
for 4 days. This result correlated well with the higher amounts of astaxanthin
accumulated in E17 than in WT cells under high light stress (Figure 4.2A) or
glucose induction conditions (Figure 4.2B).
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Chpater 4. Molecular characterization of the Chlorella zofingiensis mutant E17
Figure 4.5 Transcript levels of PDS, BKT and CHYb in WT and E17 cells
cultured under standard growth conditions (Ctr), high light induction for 1d (HL
1d) or 30 g L-1 glucose induction for 4d (30G 4d). Actin gene (ACT) was used as
control.
4.5 Discussion
C. zofingiensis synthesizes astaxanthin as secondary carotenoid that is
stored in lipid vessicles outside the chloroplasts under stress conditions such as
high light irradiation or nitrogen deficiency (Rise et al., 1994; Bar et al., 1995). A
mutation in PDS revealed that this enzyme is one of the limiting steps of its
biosynthetic pathway. An amino acid substitution (L516F) in PDS can not only
make C. zofingiensis resistant to norflurazon (Figure 4.1), but also enhance the
biosynthesis of TC and astaxanthin (Figure 4.2). Norflurazon-resistant phenotypes
had been found before in Synechococcus PCC 7942 and Synechocystis sp. PCC
6803 mutants resulting from either a point mutation or a deletion in the upstream
region of PDS gene (Chamovitz et al., 1991, 1993; Martinez-Ferez & Vioque,
1992). Point mutations in PDS give rise to a modified structure for the
herbicide-binding site; whereas the deletion in regulating region causes
up-regulation of PDS transcription and consequently a great increase of PDS
protein (Chamovitz et al., 1993). Point mutations in PDS also resulted in plant
mutants resistant to fluridone, another bleaching herbicide that also specifically
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Chpater 4. Molecular characterization of the Chlorella zofingiensis mutant E17
inhibits the desaturation activity of PDS (Michel et al., 2004; Arias et al., 2005).
Some mutations in PDS conferred cross-resistance to fluridone and norflurazon
(Arias et al., 2006) which is not the case for E17 cells (data not shown).
Norflurazon was reported to inhibit PDS by competition with the cofactor
plastoquinone (Breitenbach et al., 2001). Therefore, bleaching herbicides
including norflurazon can be regarded as plastoquinone analogs targetting the
same binding region (Sandmann, 2002).
E17 harbors a point mutation (C to T) in PDS, causing an amino acid
substitution (L to F) at codon position 516. Surrounding the L residue is a
conserved motif in PDS polypeptides from plants (Pecker et al., 1992; Scolnik &
Bartley, 1993), algae (Harker & Hirschberg, 1997; McCarthy et al., 2004; Huang
et al., 2008) and cyanobacteria (Chamovitaz et al., 1993). Previous mutations
including the substitution of the L residue to arginine (R) in PDS had been found
to confer norflurazon-resistant on Synechococcus sp. PCC 7942 but meanwhile
lower the catalytic activity in phytoene desaturation and consequently reduce the
production of downstream carotenoids (Sandmann et al., 1993). Similar results
were obtained when the same mutation occurred in the PDS from H. pluvialis and
C. zofingiensis (Steinbrenner & Sandmann, 2006; Huang et al., 2008). Thus the L
residue may play a decisive role in the binding of herbicide and the binding of
plastoquinone. In contrast to the substitution of L to R, the L to F exchange may
favor the kinetics for plastoquinone binding resulting in higher PDS specific
activity (Table 4.3). This leads to a higher TC accumulation in E17 (Figure 2).
Transgenic H. pluvialis overexpressing an additional PDS gene also enhanced TC
content (Steinbrenner & Sandmann, 2006). Determination of transcript levels of
carotenogenic genes showed no changes for PDS in E17 (Figure 4.5). However,
E17 accumulated higher levels not only of TC but also of astaxanthin than WT
(Figure 4.2; Table 4.2). This is well-explained by the increased transcripts of BKT
and CHYb in E17 (Figure 4.5). However, the regulatory mechanism leading to an
up-regulation of both transcripts in E17 as a consequence of higher PDS activity
remains open.
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Chpater 4. Molecular characterization of the Chlorella zofingiensis mutant E17
In conclusion, the PDS gene of C. zofingienesis E17 harbors a point
mutation which leads to one amino acid substitution and makes the mutated cells
resistant to norflurazon. In contrast to all other mutations conferring norflurazon
resistance, PDS in E17 shows increased specific activity leading to enhanced
biosynthesis of carotenoids including the high-value astaxanthin. The mutated
PDS gene may serve as a dominant selectable marker for genetic engineering of C.
zofingiensis and even other green algae to enhance the biosynthesis of astaxanthin.
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Chpater 5. Transformation of Chlorella zofingiensis with a phytoene desaturase gene for
enhanced astaxanthin accumulation
Chapter 5
Transformation of Chlorella zofingiensis with a phytoene
desaturase gene for enhanced astaxanthin accumulation
5.1 Abstract
PDS-L516F from the C. zofingiensis mutant E17 was revealed to show
31-fold greater resistance to norflurazon and have 29% higher desaturation
activity compared with the unaltered PDS (Chapter 4). In this chapter, the
transformation of C. zofingiensis with PDS-L516F gene was conducted, with an
attempt to enhance the production of carotenoids. PDS-L516F gene was
introduced into C. zofingiensis via biolistic approach. Transformants harboring the
mutated PDS gene showed strong resistance to the herbicide norflurazon. The
transformant P6 could accumulate 46.3% more astaxanthin than the WT. The
enhanced accumulation of astaxanthin in the transformant was revealed to be
related to the increase of PDS transcript. These results clearly show that the
mutated PDS gene is a useful selectable marker that can be used for genetic
engineering of carotenoid biosynthesis in C. zofingiensis.
5.2 Introduction
Over the past decades, the carotenoid biosynthetic pathway and
carotenogenic genes have been thoroughly investigated. It has greatly facilitated
the genetic manipulation of carotenoid biosynthesis for special purposes, such as
the enhancement of pre-existing carotenoids (Shewmaker et al., 1999; Romer et
al., 2000; Steinbrenner & Sandmann, 2006) and the production of new pigments
(Mann et al., 2000; Ralley et al., 2004; Morris et al., 2006; Jayaraj et al., 2008;
104
Chpater 5. Transformation of Chlorella zofingiensis with a phytoene desaturase gene for
enhanced astaxanthin accumulation
Zhong et al., 2008). While the engineering of carotenoid biosynthesis in higher
plants has achieved great progress, it still remains at the very preliminary stage in
many commercially important algae due to the shortage of suitable selectable
genes for development of functional transformation systems. Modified
endogenous genes as selectable markers proved effective in several algae
(Dawson et al., 1997; Randolph-Anderson et al., 1998; Kovar et al., 2002;
Steinbrenner & Sandmann, 2006).
Phytoene desaturase is considered to catalyze a rate-limiting step in
carotenoid biosynthesis (Chamovitz et al., 1993). It is also a target of various
bleaching herbicides such as norflurazon and fluridone. Overexpression of PDSor crtI-type phytoene desaturase gene could result in the elevated production of
carotenoids in transgenic plants and algae (Misawa et al., 1993; Romer et al.,
2000; Steinbrenner & Sandmann, 2006). In chapter 4, the PDS gene of C.
zofingiensis mutant E17 was isolated and characterized. This PDS gene, with a
codon change from L to F at position 516 (Figure 5.1A), was revealed to be
resistant to norflurazon and have an improved desaturation activity. Since C.
zofingiensis is very sensitive to this herbicide, PDS-L516F gene may be adopted
as a dominant selectable marker for the transformation of C. zofingiensis. It is
expected that transformants harboring the PDS-L516F gene would synthesize
more carotenoids than WT.
This chapter described the nuclear transformation of C. zofingiensis with
PDS-L516F gene as a selectable marker. Putative transformants showed
noflurazon resistance, increased PDS expression and elevated accumulation of
carotenoids including astaxanthin.
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Chpater 5. Transformation of Chlorella zofingiensis with a phytoene desaturase gene for
enhanced astaxanthin accumulation
Figure 5-1 (A) PDS from the norflurazon-resistant C. zofingiensis mutant E17
showing the codon change from Leu to Phe at position 516 and the chloroplast
transit peptide (TP) identified by the ChloroP program. (B) Map of C. zofingiensis
transformation vector pBlue-PDS-L516F with restriction sites indicated. The E17
PDS cDNA with its promoter and terminater (PDS-L516F) is inserted into the
vector pBluescript SKII (+). The construct also contains the ampicillin resistance
gene, the E. coli origin of replication (ColE1).
5.3 Materials and methods
5.3.1 Algal strain and culture conditions
The maintenance and inoculation of C. zofingiensis were described in
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Chpater 5. Transformation of Chlorella zofingiensis with a phytoene desaturase gene for
enhanced astaxanthin accumulation
2.3.1. For herbicide treatment, the cell cultures of WT and transformants were
grown in Kuhl medium for 4 days and then inoculated into the same medium
containing 0.5 μM norflurazon for growth under continuous illumination of 25
µmol photon m-2 s-1. For induction of astaxanthin accumulation, the 4-day cultures
mentioned above were inoculated into fresh Kuhl medium containing 30 g L-1
glucose and maintained in the dark.
5.3.2 Construction of the transformation vector pBlue-PDS-L516F
and transformation protocol
A 2.4-kb promoter and a 0.6-kb terminator of PDS were amplified,
digested (HindIII/EcoRI and BamHI/XbaI, respectively) and inserted sequentially
into the corresponding restriction sites of the pBluescript SKII(+) vector
(Stratagene),
followed
by
the
insertion
of
PDS-L516F
excised
from
pUC-czPDS-L516F (Chapter 4), giving rise to the transformation vector
pBlue-PDS-L516F (Figure 5.1B).
C. zofingienesis cells grown in Kuhl medium for 3 days under standard
growth conditions were collected and re-suspended to a density of around 1 × 108
cells mL-1 in liquid medium. One milliliter of the concentrated cells was used for
each bombardment and plated on filters on Kuhl plates. For transformation, the
Biolistic PDS-1000/He system was employed (Bio-Rad, Hercules, CA, USA).
Fifty microliters of a Gold particle solution (0.6 μm, 60 mg mL-1) was mixed with
5 μL of plasmid solution (1 μg μL-1), 50 μL of 2.5 M CaCl2, and 20 μL of 0.1 M
spermidine (Sigma). The mixture was incubated at room temperature for 10 min
and centrifuged for 10 s. The pellet was washed once with 70% ethanol and twice
with 100% ethanol, and re-suspended in 50 μL ethanol. Each 10 μL of the
DNA-coated gold particles was layered on a macrocarrier. Plates were bombarded
from a distance of 6 cm under vacuum of 28 mm Hg using 1, 350-lb in-2 rupture
disks. After a 24-h recovery, the bombarded cells were spread on Kuhl plates
107
Chpater 5. Transformation of Chlorella zofingiensis with a phytoene desaturase gene for
enhanced astaxanthin accumulation
containing 0.5 μM norflurazon. Colonies appearing after 3 to 4 weeks were picked
up and re-streaked three times on selective Kuhl plates.
5.3.3 Genomic DNA and RNA isolation
Genomic DNA and RNA isolation were described in 2.3.2
5.3.4 PCR determination of transformants
To distinguish the introduced PDS-L516F from genomic PDS gene, a
pair of PCR primers that locates at two individual exons interrupted by a 0.27-kb
intron were designed, which amplified 0.62- and 0.35-kb products for genomic
PDS and PDS-L516F genes respectively (Table 5.1). Primers for amplification of
a 0.22-kb fragment of ampicillin resistance gene were also employed (Table 5.1).
Cell samples harvested from 4-day cultures under illumination of 25 µmol photon
m-2 s-1 were used for determination of PDS expression.
Table 5.1 Primers for amplification of PDS and ampicillin resistant genes
Gene
Primers (5′-3′)
Product size (kb)
PDS
Forward: GATTGGGCGGAGTGATGAGG
0.62 (0.35)
(PDS-L516F)
Reverse: CTGTCGCTAATGCGGGTTTC
Ampicillin
Forward: AAGCCATACCAAACGACGAG
resistant gene
Reverse: GTCGTGTAGATAACTACGATA
5.3.5 Cell dry weight determination
Cell dry weight determination was described in 3.3.5.
108
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Chpater 5. Transformation of Chlorella zofingiensis with a phytoene desaturase gene for
enhanced astaxanthin accumulation
5.3.6 Extraction and analysis of pigments
The extraction and pigment analysis were described in 3.3.7.
5.3.7 RT-PCR assay
RT-PCR assay was described in 3.3.8 except that the amplification cycles
for PDS were lowered to 22.
5.4 Results
5.4.1 Analysis of C. zofingiensis transformants
After
transformation
of
C.
zofingiensis
with
the
vectors
pBlue-PDS-L516F and pBlue-PDS by particle bombardment and a recovering
period, the cells were spread on selective Kuhl plates containing 0.5 μM
norflurazon. After 3 to 4 weeks of growth, colonies appeared in samples
transformed with pBlue-PDS-L516F, but not in samples transformed with
pBlue-PDS that harbors the WT PDS gene. The colonies were re-streaked three
times on selective plates and then inoculated into liquid media. To determine the
stable nuclear integration of PDS-L516F gene, PCR analysis of the transformants
was performed with primers specific to the PDS gene and the ampicillin resistance
cassette of the transformation vector. The endogenous PDS gene was detected in
the WT and all transformnts (Figure 5.2A, upper band with a size of 0.62 kb). The
transformants examined had an additional shorter band amplified from the
PDS-L516F gene (Figure 5.2A, lower band with a size of 0.35 kb). PCR
determination of the ampicillin resistance cassette was also conducted and shown
in Figure 5.2B. All transformants except P3 showed a 0.22-kb band. This finding
suggested that some parts of the transformation vector may lose during its
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Chpater 5. Transformation of Chlorella zofingiensis with a phytoene desaturase gene for
enhanced astaxanthin accumulation
integration into the nuclear genome of C. zofingiensis.
To survey the expression of transgene in transformants, RT-PCR analysis
was performed. Cell samples were harvested from 4-day cultures under
illumination of 25 µmol photon m-2 s-1. Compared with WT cells that exhibited a
basal expression of the PDS gene, transformed cells contained increased levels of
PDS transcripts (Figure 5.2C).
Figure 5-2 PCR determination of C. zofingiensis transformants. Primers for
amplification of PDS and ampicillin resistance genes were shown in Table 5.1. (A)
PCR amplification of PDS-L516F gene in transformants. (B) PCR amplification
of ampicillin resistant gene in transformants. (C) RT-PCR determination of PDS
gene expression in transformants. For comparison, total RNA was equally loaded
(lower panel).
To obtain further information about the expression of transgenic
PDS-L516F, and the functionality of the 2.4-kb promoter and 0.6-kb terminator of
PDS gene, cDNA synthesis and subsequent sequencing of the PDS transcript pool
were conducted. No additional mutations were observed in the PDS cDNA except
for a mixture of CTT and TTT at codon position 516 for transformants. These
findings indicate that the 2.4-kb promoter region of PDS gene is sufficient to drive
the expression of transgene in C. zofingiensis.
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Chpater 5. Transformation of Chlorella zofingiensis with a phytoene desaturase gene for
enhanced astaxanthin accumulation
5.4.2 Resistance of transformants to norflurazon
To survey the effect of norflurazon on cell growth and carotenoid
accumulation, time courses of biomass, TC and astaxanthin contents of the WT
and transformants were conducted. Cell samples treated with 0.5 μM norflurazon
or without norflurazon were cultured under the continuous light illumination of 25
µmol photon m-2 s-1. In the presence of 0.5 μM norflurazon, the growth of WT
was severely inhibited, as indicated by the much lower cell biomass (Figure 5.3A,
left); the biosyntehsis of TC and astaxanthin was also severely inhibited and got
completely blocked as the exposure time to norflurazon prolonged (Figure 5.3B
and C, left). In contrast, the growth of the transformant P6 was not sensitive to
norflurazon, as indicated by the unaffected cell biomass (Figure 5.3A, right); the
accumulation of TC and astaxanthin was only slightly alleviated (Figure 5.3B and
C, right). At the expense of colored carotenoids, phytoene was found to
accumulate in both norflurazon-treated WT and P6 cells, together with the
concurrent decrease of chlorophylls, yet P6 was less affected (Table 5.2). These
results indicate that the transformant P6 exhibited strong resistance to norflurazon.
Similar to P6, other transformants were also resistant to norflurazon (data not
shown). In addition, the transformation stability was analyzed. After more than 50
times of subculture under non-selective conditions, the transformants showed no
loss of norflurazon resistance and stable integration of transgene in the genome
(data not shown).
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Chpater 5. Transformation of Chlorella zofingiensis with a phytoene desaturase gene for
enhanced astaxanthin accumulation
Figure 5.3 The effect of norflurazon on biomass (A), TC (B) and astaxanthin (C)
contents of WT (left) and transformant P6 (right). Cells were cultured under
illumination of 25 µmol photon m-2 s-1.
Table 5.2 Content of colored carotenoids, phytoene and chlorophyll in algal cells
cultured for 4 days with (+) or without (-) 0.5 μM norflurazaon
Pigments
WT
P6 transformant
(mg g-1 dry weight)
-
+
-
+
Colored carotenoids
1.79 ± 0.07
0.12 ± 0.01
1.92 ± 0.08
1.51 ± 0.05
Nd a
0.89 ± 0.03
Nd
0.19 ± 0.01
5.84 ± 0.23
0.94 ± 0.04
5.98 ± 0.31
3.39 ± 0.14
Phytoene
Chlorophylls
a
Nd, not detected
112
Chpater 5. Transformation of Chlorella zofingiensis with a phytoene desaturase gene for
enhanced astaxanthin accumulation
5.4.3 Enhanced biosynthesis of astaxanthin in P6 induced by
glucose
Since glucose can boost C. zofingiensis to synthesize carotenoids
including astaxanthin (Ip & Chen, 2005; Sun et al., 2008), it is interesting to
compare the time courses of TC and astaxanthin accumulation in heterotrophic
WT and P6 cells cultured with glucose, which is shown in Figure 5.4A. Upon
induction by glucose, the TC and astaxanthin contents in WT cells remained
almost unchanged during the first 2 days. Thereafter, a significant increase in both
TC and astaxanthin contents was observed. The transformant P6 showed the same
induction pattern of TC and astaxanthin as that of WT. However, P6 produced
apparently greater amounts of carotenoids than the WT during the whole
induction period. The carotenoid contents of WT and P6 induced with glucose for
5 days are shown in Table 5.3. Lutein, astaxanthin and adonixanthin were the
major carotenoids. Significant higher amounts of β-carotene, lutein, zeaxanthin,
canthaxanthin, adonixanthin and astaxanthin were observed to accumulate in P6
cells. The TC and astaxanthin contents of P6 were 2.45 mg g-1 and 0.64 mg g-1,
20.7% and 46.3% higher than that of WT respectively. Correlated with the higher
content of astaxanthin, P6 cells exhibited a deeper yellow-orange color than WT
(Figure 5.4B).
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Chpater 5. Transformation of Chlorella zofingiensis with a phytoene desaturase gene for
enhanced astaxanthin accumulation
Figure 5.4 Comparison of WT and P6 cultured with 30 g L-1 glucose in the dark.
(A) Time course of accumulation of TC (square) and astaxanthin (circle) in WT
(open) and P6 (filled). (B) Algal cells of WT and P6 from 5-day cultures.
Table 5.3 Pigment profiles in WT and P6 cells cultured with 30 g L-1 glucose in
the dark for 5 days a
TC
Chl
(mg g-1)
(mg g-1)
WT
2.03 ± 0.11
P6
2.45 ± 0.13
a
Carotenoid composition (μg g-1)
Neo
Vio
Ant
Ast
Ado
Lut
Zea
Can
β-Car
3.23 ± 0.15
69 ± 2.5
32 ± 1.4
73 ± 2.8
434 ± 24
366 ± 16
542 ± 24
224 ± 14
205 ± 13
86 ± 4.9
2.94 ± 0.13
61 ± 3.3
35 ± 1.8
61 ± 3.3
635 ± 29
438 ± 25
623 ± 38
256 ± 17
231 ± 11
109 ± 5.7
Data were expressed as mean values of three independent experiments. TC, total
carotenoids; Chl, Chlorophylls; Neo, neoxanthin; Vio, violaxanthin; Ant,
antheraxanthin; Ast, astaxanthin; Ado, adonixanthin; Lut, lutein; Zea, zeaxanthin;
Can, canthaxanthin; β-Car, β-carotene
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Chpater 5. Transformation of Chlorella zofingiensis with a phytoene desaturase gene for
enhanced astaxanthin accumulation
5.4.4 Transcription analysis of carotenogenic genes
It has been reported that the elevated biosynthesis of carotenoids
including astaxanthin in C. zofingiensis by glucose induction was correlated to the
increased transcript levels of carotenogenic genes (Li et al., 2009). To further
examine if the elevated production of TC and astaxanthin by P6 as compared to
the WT was attributed to the up-regulation of PDS and/or other carotenogenic
genes, the transcript levels of PDS, BKT and CHYb were semi-quantified by using
the RT-PCR approach. Cells samples induced for 0 day, 2 days and 4 days were
employed. At all stages examined, P6 contained a much higher amount of PDS
transcripts than WT (Figure 5.5). Although no apparent differences in transcripts
of BKT and CHYb between WT and P6 were observed for 0-day cell cultures,
significant higher transcript levels of these two genes were found in P6 after
induction of 2 days and 4 days (Figure 5.5). This result correlated well to the
higher amounts of TC including astaxanthin accumulated in P6 cells in response
to glucose induction (Figure 5.4A).
Figure 5.5 Transcript levels of PDS, BKT and CHYb in WT and P6 cells cultured
in the dark with 30 g L-1 glucose for 0 day (0 D), 2 days (2 D) and 4 days (4 D).
Actin gene (ACT) was used as control.
115
Chpater 5. Transformation of Chlorella zofingiensis with a phytoene desaturase gene for
enhanced astaxanthin accumulation
5.5 Discussion
C. zofingiensis represents a typical fast growing alga that can be
heterotrophically cultivated for astaxanthin production (Ip & Chen, 2005). The
enhanced cellular astaxanthin content, which may be achieved through genetic
engineering of the carotenoid biosynthesis, will make C. zofingiensis more
attractive for commercial use. At present, no appropriate transformation system is
available for the alga. The availability of suitable promoters, terminators and
promising reporter or resistance genes proved crucial for the development of a
transformation system of algae (Cerutti et al., 1997; Schroda et al., 2000; Poulsen
& Kroger, 2005; Steinbrenner & Sandmann, 2006). It was reported in chapter 4
that the PDS-L516F gene isolated from the C. zofingiensis mutant E17 encoded a
desaturase having 31-fold greater resistance to norflurazon than the WT one. The
PDS-L516F gene can therefore serve as a selectable marker for the transformation
of C. zofingiensis.
The integration of the transformation vector pBlue-PDS-L516F was
investigated by PCR analysis (Figure 5.2A and B). Using the primers specific to
two individual exons of PDS gene revealed that all transformants except the WT
had an additional band indicating the presence of PDS-L516F in the nuclear
genome. The integrated transgene conferred enhanced PDS expression on
transformants, as indicated by the increased amounts PDS transcripts (Figure
5.2C). This was further confirmed by the analysis of the total PDS gene transcripts,
which revealed the presence of a mixture of WT and mutated PDS transcripts.
These results also suggested that the expression cassette of PDS gene drove
efficiently the expression of PDS-L516F cDNA and that the endogenous PDS
gene was intact.
The expression of PDS-L516F gene not only conferred norflurazon
resistance on transformants (Figure 5.3), but also influenced their carotenoid
metabolism. Carotenoid analysis showed that the transformant P6 could
accumulate higher amount of TC than WT when cultured with glucose in the dark
116
Chpater 5. Transformation of Chlorella zofingiensis with a phytoene desaturase gene for
enhanced astaxanthin accumulation
(Figure 5.4A). This is in accordance with the previous findings that the
introduction of a mutated PDS gene resulted in elevated production of carotenoids
in transgenic H. pluvialis (Steinbrenner & Sandmann, 2006). In addition to the
primary carotenoids, secondary carotenoids such as canthaxanthin, adonixanthin
and astaxanthin were also found in higher amounts in P6 (Table 5.3). After 5 days
of glucose induction, P6 produced 46.3% more astaxanthin than WT did. It is
possible that the increased expression of PDS gene results in an increase in the
flux of colored carotenoids, which in turn up-regulate the transcription of such
carotenogenic genes as BKT and CHYb, leading to the enhanced synthesis of
astaxanthin (Figure 5.4 and 5.5). Similar phenomenon was observed for
transgenic tomato plants in which overexpression of a crtI-type phytoene
desaturase caused up-regulation of ζ-carotene desaturase and lycopene β-cyclase
genes and elevation of β-carotene synthesis (Romer et al., 2000). These results
suggest that phytoene desaturation is a rate-limiting step for carotenoid
biosynthesis in C. zofingiensis.
The transformant P6 harbored at least one copy of the PDS-L516F gene
from the C. zofingiensis mutant E17. Compared with E17, P6 contained much
higher amount of PDS transcripts, as indicated by the RT-PCR analysis (Figure
5.6A). However, no significant difference in both TC and astaxanthin contents
was observed between E17 and P6 (Figure 5.6B). This might be due to the
involvement of complex mechanisms in carotenoid biosynthesis, which restricts
carotenoid flux within a certain range when genetic engineering of the phytoene
desaturation step only is implemented. Considering the presence of substantial
amounts of the end product canthaxanthin and the intermediate product
adonixanthin (Table 5.3), CHYb may not accept canthaxanthin as substrate to
produce astaxanthin and BKT might not efficiently convert adonixanthin to
astaxanthin in C. zofingiensis. Therefore, the manipulation of specific astaxanthin
biosynthetic steps by introducing a carotenoid hydroxylase that can convert
canthaxanthin to astaxanthin and a carotenoid ketolase that can catalyze the
efficient conversion of adonixanthin to astaxanthin, should represent a feasible
117
Chpater 5. Transformation of Chlorella zofingiensis with a phytoene desaturase gene for
enhanced astaxanthin accumulation
strategy for enhanced accumulation of astaxanthin (Figure 1.16).
In conclusion, the results reported here clearly demonstrate that the
PDS-L516F gene can be adopted as a dominant selectable marker for stable
nuclear transformation of C. zofingiensis. The transformants overexpressing the
PDS gene could enhance the flux of colored carotenoids, providing a proof of
concept for genetic engineering of the carotenoid biosynthesis in C. zofingiensis
toward elevated astaxanthin production.
Figure 5.6 Comparison of E17 and P6 cultured with 30 g L-1 glucose in the dark.
(A) Transcript levels of PDS in E17 and P6 cells cultured for 2 days (2 D) and 4
days (4 D). Actin gene (ACT) was used as control. (B) Carotenoid accumulation in
E17 and P6 cells cultured for 4 days (4 D) and 6 days (6 D).
118
PART III
POTENTIAL ASSESSMENT OF CHLORELLA
ZOFINGIENSIS AS A BIODIESEL FEEDSTOCK
119
Chapter 6. Differential lipid and fatty acid profiles of photoautotrophic and heterotrophic
Chlorella zofingiensis: assessment of algal oils for biodiesel prodoction
Chapter 6
Differential lipid and fatty acid profiles of
photoautotrophic and heterotrophic Chlorella zofingiensis:
assessment of algal oils for biodiesel production
6.1 Abstract
Profiles of algal lipids and fatty acids, which depend on species and
culture conditions, are important data to evaluate the oils for biodiesel production.
The objective of this study was to document and compare the lipid class and fatty
acid composition of the green microalga Chlorella zofingiensis cultivated under
photoautotrophic and heterotrophic conditions. Compared with photoautotrophic
cells, a 900% increase in lipid yield was achieved in heterotrophic cells fed with
30 g L-1 glucose. Furthermore heterotrophic cells accumulated predominantly
neutral lipids (NL) that accounted for 79.5% of total lipids, with 88.7% of NL
being triacylglycerol (TAG); whereas photoautotrophic cells contained mainly the
membrane lipids glycolipids (GL) and phospholipids (PL). Together with the
much higher content of oleic acid (C18:1, 35.2% of total fatty acids), oils from
heterotrophic C. zofingiensis appear to be more feasible for biodiesel production.
The study highlights the possibility of using heterotrophic algae for producing
high quality biodiesel.
120
Chapter 6. Differential lipid and fatty acid profiles of photoautotrophic and heterotrophic
Chlorella zofingiensis: assessment of algal oils for biodiesel prodoction
6.2 Introduction
Energy is an indispensable factor to sustain our economic growth and
living standard. Up to date, fossil-derived fuels such as coal, petroleum and
natural gas have still served as the main global energy sources. The growing
consumption of fossil fuels, however, has caused many environmental problems
that threaten our ecosystem (Jain & Sharma, 2010; Lim & Teong, 2010).
Furthermore, fossil fuels are recognized as unsustainable due to their depleting
supplies (Chisti, 2008; Shafiee & Topal, 2009). Thus, clean and renewable energy
has to be explored. Biofuels, especially biodiesel, are carbon neutral, contributing
less emission of gaseous pollutants, and are therefore environmentally beneficial
(Ma & Hanna, 1999; Fukuda et al., 2001; Knothe, 2005b; Gerpen, 2005; Meher et
al., 2006; Hu et al., 2008). Biodiesel is chemically composed of fatty acid methyl
ester (FAME) that is traditionally derived from transesterification of vegetable oils
or animal fats (Lang et al., 2001; Al-Widyan & Al-Shyoukh, 2002; Zhang et al.,
2003; Demirbas, 2005).
Microalgae have been cited as promising feedstocks for biodiesel
production because of their rapid growth rate and high intracellular content of
lipids (Chisti, 2007; Hu et al., 2008). Two systems are commonly used to cultivate
algae: the open system (open raceway ponds) and the closed system
(photobioreactors). As high oil strains generally grow slower than low oil strains,
open systems can only be used for culturing “extremophile” that can tolerate
extreme conditions (e.g., high salinity or alkalinity) in which other strains cannot
survive (Vasudevan & Briggs, 2008). Unfortunately no such strains with high
content of oils have been isolated. Enclosed photobioreactors are desirable for
maintaining pure culture and fast growth of oil-rich algae. However, as algae only
accumulate large amounts of oil under stresses (mostly nutrient restrictions) that
also limit cell growth, it is difficult to balance the growth and oil production of an
alga with current photobioreactors. These challenges need to be addressed before
profitable biodiesel can be produced from algae.
121
Chapter 6. Differential lipid and fatty acid profiles of photoautotrophic and heterotrophic
Chlorella zofingiensis: assessment of algal oils for biodiesel prodoction
C. zofingiensis is a particular green alga in that it can grow well
photoautotrophically as well as heterotrophically (Orosa et al., 2000; Ip & Chen,
2005; Sun et al., 2008). High cell biomass (up to 53 g L-1) of the alga could be
achieved by using a glucose fed-batch fermentation strategy (Sun et al., 2008).
Furthermore, C. zofingiensis also accumulated high amounts of the secondary
ketocarotenoid astaxanthin when grown with glucose as the sole carbon and
energy source (Ip & Chen, 2005; Sun et al., 2008). Phylogenetic relatives of C.
zofingiensis, such as Chlorella vulgaris and Chlorella protothecoides were
reported to accumulate high levels of lipids when cultivated under heterotrophic
conditions (Miao & Wu, 2004; Liu et al., 2008; Hsieh & Wu, 2009). The fact that
carotenoid biosynthesis increases with oil synthesis further raises the possibility of
using C. zofingiensis as a cell factory for algal oil. However, knowledge on
Chlorella species to accumulate lipids and fatty acids under different growth
modes remains largely unknown. In addition, the feasibility of various algal oils
for producing high quality biodiesel need to be assessed.
The objective of this study was to document and compare the lipid class
and fatty acid composition of C. zofingiensis cultivated under photoautotrophic
and heterotrophic conditions so as to assess its feasibility for biodiesel production.
6.3 Methods and materials
6.3.1 Algal strain and culture conditions
The maintenance and inoculation of C. zofingiensis were described in
2.3.1. For photoautotrophic growth, the seed cells were inoculated into the fresh
medium and allowed to grow under continuous illumination of 30 µmol photon
m-2 s-1 with orbital shaking at 150 rpm. For heterotrophic growth, the seed cells
were inoculated into the medium containing 30 g L-1 glucose and cultured in the
dark with orbital shaking at 150 rpm.
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Chapter 6. Differential lipid and fatty acid profiles of photoautotrophic and heterotrophic
Chlorella zofingiensis: assessment of algal oils for biodiesel prodoction
6.3.2
Determination
of
glucose
concentration,
nitrate
concentration, cell dry weight and specific growth rate
The cells were centrifuged at 3,800 g for 5 min. Glucose concentration
and nitrate concentration in the supernatant were determined according to Miller
(1959) and Elton-Bott (1979), respectively. Cell dry weight determination was
described in 3.3.5. The specific growth rate (µ) at the exponential phase was
calculated according to the equation µ = (ln X2 − ln X1) / (t2 − t1), where X2 and
X1 are the cell dry weight concentration (g L−1) at time t2 and t1, respectively.
6.3.3 Lipid extraction and analysis
Cells were harvested and lyophilized for lipid extraction and analysis.
Total lipids were extracted from 200 mg of lyophilized cells with a solvent
mixture of chloroform, methanol and water (2:1:0.75, by vol.) according to the
modified Folch procedure (Christie, 2003). The extract was dried in a rotary
evaporator, and then weighed, re-suspended in chloroform, and finally stored at
-20 °C under nitrogen gas to prevent lipid oxidation.
Lipids were separated into neutral lipids (NL), glycolipids (GL) and
phospholipids (PL) using solid-phase extraction (Christie, 2003). A 500 mg
Sep-PakTM cartridge of silica gel (Waters) was first conditioned by elution with 5
ml of chloroform, and about 50 mg of lipids were then applied to the column.
Elution with 10 ml of solvent in the order of chloroform, acetone and methanol
yielded NL, GL and PL, respectively. Each fraction was dried under a stream of
nitrogen gas, weighted and then re-suspended in 0.1 mL of chloroform.
The NL fraction was
subjected to
one-dimensional thin-layer
chromatography (TLC) for lipid class separation and identification, using TLC
plates (20 × 20 cm) coated with silica gel 60 (Merck, Whitehouse Station, NJ,
USA) (Fan et al., 2007). Plates were activated in an oven at 100 °C for 2 h prior to
123
Chapter 6. Differential lipid and fatty acid profiles of photoautotrophic and heterotrophic
Chlorella zofingiensis: assessment of algal oils for biodiesel prodoction
use. The solvent mixture hexane/diethyl ether/acetic acid (70:30:1, by vol.) was
used. After co-chromatography with pure standards (Sigma), bands of lipid classes
were stained with 2,7-dichlorofluorescein (Sigma) and visualized under UV light.
6.3.4 Fatty acid analysis
Fatty
acid
methyl
esters
(FAMEs)
were
prepared
by
direct
transmethylation with sulphuric acid in methanol (Christie, 2003). The FAMEs
were analyzed
by using
an HP 6890 capillary gas chromatography
(Hewlett-Packard, Palo Alto, CA) equipped with a flame ionization detector (FID)
and a HP-INNOwax capillary column (30 m × 0.32 mm) (Agilent Technologies,
Inc., Wilmington, DE). Nitrogen was used as carrier gas. Initial column
temperature was set at 170 °C, which was progressively raised to 230 °C at
1 °C/min. The injector was kept as 250 °C with an injection volume of 2 µL under
splitless mode. The FID temperature was set at 270 °C. FAMEs were identified by
chromatographic comparison with authentic standards (Sigma). The quantities of
individual FAME were estimated from the peak areas on the chromatogram using
heptadecanoic acid (Sigma) as the internal standard.
6.4.5 Calculation of iodine value
The iodine value of algal oils was calculated according to AOCS
recommended practice Cd 1c-85, which estimates the grams of halogen absorbed
by 100 g of oil.
124
Chapter 6. Differential lipid and fatty acid profiles of photoautotrophic and heterotrophic
Chlorella zofingiensis: assessment of algal oils for biodiesel prodoction
6.5 Results
6.5.1 Growth characteristics and fatty acid accumulation
C. zofingiensis can grow photoautotrophically and heterotrophically
utilizing sugars as the carbon and energy sources (Del Campo et al., 2004; Ip &
Chen, 2005; Sun et al., 2008). The growth parameters of the algal cells grown
photoautotrophically or heterotrophically in batch cultures were characterized
here. As shown in Table 6.1, photoautotrophic cells grew slowly indicated by the
low specific growth rate (0.235 d-1) and cell biomass (1.9 g L-1). In contrast, the
alga grew fast under heterotrophic conditions supplemented with 30 g L-1 glucose,
with a specific growth rate of 0.769 d-1 and a cell biomass concentration of 9.7 g
L-1, which are 227% and 411% higher than those of the photoautotrophic cells
respectively. The glucose utilization and growth yield coefficient based on glucose
by heterotrophic cells were revealed to be 77.8% and 0.402 g/g, respectively. In
addition, heterotrophic cells consumed nitrated from the medium rapidly (Figure
6.1A).
Table 6.1 Growth kinetic parameters of C. zofingiensis in photoautotrophic and
heterotrophic batch cultures a
Parameters b
Photoautotrophic
Heterotrophic
0.235 ± 0.014
0.769 ± 0.046
Xmax (g L-1)
1.9 ± 0.11
9.7 ± 0.33
Yx/gu (g g-1)
—
0.402 ± 0.012
Glucose utilization (%)
—
77.8 ± 4.2
μ (d-1)
a
C. zofingiensis
Data are expressed as mean ± standard deviation of triplicates. b μ, specific
growth rate; Xmax, maximum biomass concentration; Yx/glu, growth yield
coefficient based on glucose.
125
Chapter 6. Differential lipid and fatty acid profiles of photoautotrophic and heterotrophic
Chlorella zofingiensis: assessment of algal oils for biodiesel prodoction
Figure 6.1 Cell growth (down triangle) and nitrate consumption (up triangle) by
heterotrophic C. zofingiensis (A) and accumulation of TFA (square) and oleic acid
(circle) in autotrophic (open) and heterotrophic (solid) algal cells (B).
To investigate the effect of growth modes on fatty acid accumulation, the
time courses of total fatty acids (TFA) and oleic acid (C18:1) accumulated in C.
zofingiensis grown photoautotrophically or heterotrophically were surveyed here.
Figure 6.1B shows that photoautotrophic cells maintained basal amounts of TFA
and oleic acid (15.8% and 3.23% of dry weight on day 14, respectively) during
the time surveyed. In contrast, a steady increase in both TFA and oleic acid was
126
Chapter 6. Differential lipid and fatty acid profiles of photoautotrophic and heterotrophic
Chlorella zofingiensis: assessment of algal oils for biodiesel prodoction
observed in heterotrophic cells during the culture time, which reached up to
42.1% and 15.2% of dry weight respectively on the 14th day. Similar to plants,
microalgae synthesize fatty acids in the chloroplast using a single set of enzymes,
of which acetyl-CoA carboxylase (ACCase) is the key one in regulating fatty acid
synthesis rates while stearoyl ACP desaturase adds the first double bond to acyl
chain and plays an important role in determining the ratio of unsaturated fatty
acids to saturated ones (Ohlrogge & Jaworski, 1997). Preliminary results showed
that glucose could triggered the strong up-regulation of ACCase and stearoyl ACP
desaturase genes in C. zofingiensis (data not shown), which may enhance the flux
of both TFA and unsaturated fatty acids and thus the accumulation of TFA and
oleic acid.
6.5.2 Lipid class composition
To further survey the lipid profiles of the alga, extraction and
characterization of the lipids from cells grown photoautotrophically or
heterotrophically for 14 days were conducted. The results were shown in Figure
6.2. Heterotrophic cells could accumulate lipids up to 51.1% of dry weight, which
is nearly 100% higher than that from photoautotrophic cells (25.8% of dry weight)
(Figure 6.2A). Total lipids were mainly separated into NL, GL and PL using
solid-phase extraction (Figure 6.2B). In photoautotrophic cells, membrane lipids,
namely GL sand PL, were the major lipid classes which altogether accounted for
70.6% of total lipids, while the storage lipids NL accounted for only 29.4% of the
total lipids (Figure 6.2B). In contrast, heterotrophic cells produced predominantly
NL which represented 80.9% of total lipids. Thus, photoautotrophic and
heterotrophic cells exhibited a great discrepancy of NL proportions (Figure 6.2B).
Unlike photoautotrophic cells, heterotrophic cells can channel excessive carbon
for the biosynthesis of storage lipids, for example NL, instead of converting
carbon into membrane lipids for building photosynthetic apparatus.
127
Chapter 6. Differential lipid and fatty acid profiles of photoautotrophic and heterotrophic
Chlorella zofingiensis: assessment of algal oils for biodiesel prodoction
Figure 6.2 Lipid content and fractionation of C. zofingiensis. (A) Lipid content of
photoautotrophic and heterotrophic cells. (B) Distribution of NL, GL, and PL in
lipids of photoautotrophic (white column) and heterotrophic (gray column) cells.
(C) Distribution of NL subclasses of photoautotrophic (white column) and
heterotrophic (gray column) cells. NL, neutral lipids; GL, glycolipids; PL,
phospholipids; SE, steroid ester; TAG, triacylglycerol; FFA, free fatty acids; DAG,
diacylglycerol; MAG, monoacylglycerol.
The NL fraction was separated by TLC into individual subclasses for
further analysis. For heterotrophic cells, TAG was the most abundant component
which accounted for 88.7% and 70.5% of NL and total lipids respectively (Figure
6.2C). Diacylglycerol (DAG, 25.2 mg g-1 dry weight), monoacylglycerol (MAG,
128
Chapter 6. Differential lipid and fatty acid profiles of photoautotrophic and heterotrophic
Chlorella zofingiensis: assessment of algal oils for biodiesel prodoction
9.1 mg g-1 dry weight), sterol and free fatty acids were found in small proportions
in heterotrophic cells (Figure 6.2C). As the key metabolites in TAG biosynthesis,
DAG and MAG were also found in low amount in other heterotrophically grown
algae (Alonso et al., 2000; Chen et al., 2007; Fan et al., 2008). Compared with
heterotrophic cells, photoautotrophic cells had a lower TAG content (65.9% of NL)
but higher contents of DAG and MAG (Figure 6.2C). Although TAG was the
major component of NL in photoautotrophic cells, its cellular content is merely
49.3 mg g-1 dry weight, which was much lower than that obtained from
heterotrophic cells (360.3 mg g-1 dry weight).
6.5.3 Fatty acid composition of individual lipid classes
Fatty acid composition is an important parameter to assess the feasibility
of algal oils for biodiesel production. Therefore here the fatty acid profiles of
individual lipid classes from both photoautotrophic and heterotrophic cells were
investigated and are presented in Table 6.2. C16:0, oleic acid and C18:2 were
found to be the major fatty acids, which together accounted for 65.3% or 77.6% of
TFA in photoautotrophic or heterotrophic cells respectively (Table 6.2). However,
saturated fatty acids (SFA), monounsaturated fatty acids (MUFA), and
polyunsaturated fatty acids (PUFA) displayed a significant interactive effect of
growth modes. For example, photoautotrophic cells contained a higher percentage
of PUFA (49.6%) but a lower MUFA (20.1%) as compared with that from
heterotrophic cells (39.1% and 37.4% respectively). The higher MUFA content in
heterotrophic cells is attributed predominantly to the higher proportion of oleic
acid (Table 6.2). Based on the fatty acid profile, the calculated iodine value of oils
from photoautotrophic and heterotrophic cells were 133.5 g I2/100 g and 117.9 g
I2/100 g, respectively.
129
Chapter 6. Differential lipid and fatty acid profiles of photoautotrophic and heterotrophic Chlorella zofingiensis: assessment of algal oils for biodiesel prodoction
Table 6.2 Fatty acid composition of individual lipid class of C. zofingiensis (% of TFA) a
Photoautotrophic
cells
Heterotrophic cells
Fatty acids
TL
TAG
DAG
MAG
SE
FFA
GL
PL
TL
TAG
DAG
MAG
SE
FFA
GL
PL
16:0
26.6 ± 1.1
25.1 ± 1.4
20.3 ± 0.8
25.9 ± 1.4
29.0 ±1.2
25.6 ±1.1
20.6 ± 0.9
37.4 ±2.3
22.2 ± 1.0
21.8 ± 1.2
20.2 ± 0.8
21.7 ± 0.7
50.5 ± 2.2
18.9 ± 0.7
27.4 ± 1.5
43.1 ± 1.7
16:1
2.2 ± 0.1
2.0 ± 0.1
9.8 ± 1.1
1.7 ± 0.1
3.8 ± 0.2
2.8 ± 0.2
1.0 ± 0.0
1.0 ± 0.1
1.7 ± 0.1
1.6 ± 0.1
4.5 ± 0.3
2.3 ± 0.1
6.6 ± 0.3
2.2 ± 0.1
2.3 ± 0.1
1.6 ± 0.1
16:2
7.7 ± 0.2
7.8 ± 0.3
11.1 ± 0.4
5.1 ± 0.3
4.6 ± 0.3
5.2 ± 0.2
8.0 ± 0.5
7.2 ± 0.4
8.3 ± 0.4
8.2 ± 0.3
9.5 ± 0.4
5.8 ± 0.2
4.7 ± 0.3
13.0 ± 0.5
19.1 ± 0.7
7.0 ± 0.2
16:3
6.6 ± 0.3
5.5 ± 0.3
10.3 ± 0.6
4.9 ± 0.3
31.5 ± 2.1
10.0 ± 0.7
7.2 ± 0.3
5.2 ± 0.3
2.1 ± 0.1
2.3 ± 0.1
3.2 ± 0.1
2.9 ± 0.1
6.2 ± 0.2
13.1 ± 0.5
3.4 ± 0.2
1.0 ± 0.1
16:4
1.1 ± 0.1
1.1 ± 0.1
2.6 ± 0.2
1.6 ± 0.1
1.4 ± 0.1
5.0 ± 0.3
3.4 ± 0.1
0.6 ± 0.0
0.2 ± 0.0
0.2 ± 0.0
0.8 ± 0.1
1.0 ± 0.1
0.9 ± 0.0
3.0 ± 0.2
0.8 ± 0.0
0.4 ± 0.0
18:0
3.7 ± 0.1
2.7 ± 0.2
4.1 ± 0.2
10.0 ± 0.3
6.2 ± 0.3
18.8 ± 0.8
6.6 ± 0.2
2.5 ± 0.1
1.2 ± 0.1
1.0 ± 0.1
5.8 ± 0.3
13.0 ± 0.5
10.1 ± 0.4
9.1 ± 0.4
8.3 ± 0.3
1.8 ± 0.1
18:1
17.9 ± 0.9
23.1 ± 1.2
15.3 ± 0.6
24.0 ± 1.4
13.6 ± 0.8
11.1 ± 0.5
18.1 ± 1.0
15.0 ± 0.4
35.2 ± 1.2
38.2 ± 1.9
25.7 ± 1.1
25.2 ± 0.9
12.8 ± 0.7
12.3 ± 0.7
14.8 ± 0.8
16.8 ± 0.7
18:2
20.8 ± 1.2
21.6 ± 0.9
13.3 ± 0.5
15.9 ± 0.6
5.3 ± 0.3
6.7 ± 0.2
22.2 ± 0.9
18.6 ± 0.9
20.2 ± 1.1
18.6 ± 0.5
19.0 ± 0.6
17.8 ± 0.8
5.8 ± 0.2
8.7 ± 0.3
5.3 ± 0.2
17.8 ± 0.5
18:3 n-6
1.4 ± 0.1
1.2 ± 0.1
1.2 ± 0.1
1.4 ± 0.1
1.2 ± 0.0
1.6 ± 0.1
0.4 ± 0.0
1.6 ±0.1
0.5 ± 0.0
0.5 ± 0.0
1.6 ± 0.1
1.6 ± 0.1
0.7 ± 0.0
2.0 ± 0.1
1.3 ± 0.1
1.0 ± 0.1
18:3 n-3
10.8 ± 0.4
8.9 ± 0.6
10.7 ± 0.7
8.4 ± 0.3
2.6 ± 0.1
9.8 ± 0.6
11.6 ± 0.4
10.2 ± 0.5
7.8 ± 0.5
7.4 ± 0.2
8.5 ± 0.3
7.1 ± 0.4
1.1 ± 0.1
14.1 ± 0.8
17.0 ± 0.6
9.1 ± 0.3
18:4
1.1 ± 0.1
1.0 ± 0.0
1.3 ± 0.1
1.2 ± 0.1
0.9 ± 0.0
3.5 ± 0.2
1.0 ± 0.1
0.9 ± 0.1
0.4 ± 0.0
0.4 ± 0.0
1.3 ± 0.1
1.7 ± 0.1
0.6 ± 0.0
3.6 ± 0.2
0.4 ± 0.0
0.5 ± 0.0
SFA (%)
30.3 ± 1.2
27.7 ± 1.0
24.3 ± 0.5
35.9 ± 1.6
35.3 ± 1.4
44.3 ± 2.5
27.2 ± 1.1
39.9 ± 2.2
23.6 ± 1.3
22.8 ± 1.0
26.0 ± 1.2
34.6 ± 1.9
60.5 ± 2.7
28.0 ± 0.8
35.7 ± 1.3
44.9 ± 2.3
MUFAc (%)
20.1 ± 0.7
25.1 ± 1.1
25.1 ± 1.6
25.7 ± 1.0
17.3 ± 0.5
14.0 ± 0.6
19.0 ± 1.0
16.0 ± 0.4
37.4 ± 1.3
41.2 ± 1.6
30.3 ± 1.5
27.5 ± 1.2
19.5 ± 1.1
14.5 ± 0.6
17.1 ± 0.5
18.3 ± 1.0
49.6 ± 2.3
47.2 ± 2.1
50.6 ± 2.7
38.4 ± 1.3
47.4 ± 1.6
41.7 ± 2.2
53.8 ± 2.7
44.1 ± 1.9
39.1 ± 2.1
36.0 ± 1.5
43.7 ± 1.7
37.9 ± 2.1
20.0 ± 1.1
57.5 ± 1.9
47.2 ± 2.4
36.8 ± 2.0
1.42 ± 0.1
1.39 ± 0.1
1.56 ± 0.1
1.23 ± 0.0
1.52 ± 0.1
1.36 ± 0.1
1.55 ± 0.1
1.24 ± 0.1
1.28 ± 0.1
1.26 ± 0.1
1.35 ± 0.1
1.20 ± 0.0
0.71 ± 0.0
1.72 ± 0.1
1.35 ± 0.1
1.05 ± 0.1
b
d
PUFA (%)
▽/mol
e
a
Data are expressed as mean ± standard deviation of triplicates, cells harvested after 14 days of cultivation. TL, total lipids; TAG, triacylglycerol; DAG, diacylglycerol;
MAG, monoacylglycerol; SE, steroid ester; FFA, free fatty acids; GL, glycolipids; PL, phospholipids. b SFA, saturated fatty acids; c MUFA, monounsaturated fatty
acids; d PUFA, polyunsaturated fatty acid; e ▽/mol, the degree of fatty acid unsaturation = [1.0 (% monoenes) + 2.0 (% dienes) + 3.0 (% trienes) + 4.0 (%
tetraenes) ]/100.
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Chapter 6. Differential lipid and fatty acid profiles of photoautotrophic and heterotrophic
Chlorella zofingiensis: assessment of algal oils for biodiesel prodoction
The overall distribution of TFA and oleic acid in the lipid classes of
heterotrophic cells is depicted in Figure 6.3. It was observed that heterotrophic C.
zofingiensis could produce a large amount of TAG (70.5% of Total lipids) as the
major storage lipid which contained 81.2% and 90.0% of TFA and total oleic acid
respectively. Since intracellular lipids can be enhanced by controlling the culture
conditions, such as limitation of nutrients (e.g., nitrogen, phosphorus, sulfur)
(Takagi et al., 2000; Khozin-Goldberg & Cohen, 2006) and salt stress (Takagi et
al., 2006), it can be expected that the TAG yields of heterotrophic C. zofingiensis
cells can further be increased by the same approaches.
Figure 6.3 Overall distribution of TFA (white column) and oleic acid (gray
column) in various lipid classes of heterotrophic C. zofingiensis.
6.6 Discussion
The cell biomass and cellular lipid content are two key factors for the
initial assessment of a microalga for biodiesel production. Under photoautotrophic
growth conditions, both cell biomass and lipid content of C. zofingiensis were low
131
Chapter 6. Differential lipid and fatty acid profiles of photoautotrophic and heterotrophic
Chlorella zofingiensis: assessment of algal oils for biodiesel prodoction
as compared with those from heterotrophic growth conditions (Table 6.1 and
Figure 6.2A). Accordingly, a much higher TFA content was observed in
heterotrophic cells (Figure 6.1B). The lipid yield of heterotrophic cells was 4.96 g
L-1, about 9 times higher than that of photoautotrophic cells (0.49 g L-1). NL
especially TAG has priority over PL or GL for biodiesel production due to their
higher content of fatty acids. In this regard, oils from heterotrophic cells are more
feasible for biodiesel production than that from the photoautotrophic cells since
heterotrophic cells achieved TAG up to 3.49 g L-1 while photoautotrophic ones
only 0.94 g L-1.
The composition and structure of fatty acid esters, such as unsaturation
degree and carbon chain length, determine the properties (e.g. cetane number,
viscosity, cold flow, oxidative stability, and iodine value) of biodiesel (Knothe,
2005). The fatty acids from C. zofingiensis were in medium length (16 to 18
carbons) with the maximum unsaturation degree being 3 (Table 6.2), which are
similar to that of plant oils currently used for biodiesel production (Singh & Singh,
2010). Compared with photoautotrophic cells, heterotrophic cells attained a much
higher amount of oleic acid which better balances oxidative stability and
low-temperature properties and therefore promotes the quality of biodiesel
(Knothe, 2008, 2009). With regarding to the oil unsaturation, oil from
photoautotrophic cells had an iodine value of 133.5 g I2/100 g that is over the
European standard of biodiesel (120 g I2/10g); whereas the iodine value of oil
from the heterotrophic cells (117.9 g I2/100 g) complied with the standard.
The general interest of using photoautotrophic algal cells for biodiesel
production is that sunlight and CO2 can be directly converted to oils. Due to the
low cell density and cellular lipid content by photoautotrophic cells, many
technical challenges remain such that algae have to be harvested from large,
shallow ponds and oils separated from the cells. To avoid those pitfalls, sugar was
employed to boost biomass and lipid yield by C. zofingiensis. The study clearly
showed that heterotrophic algal cells attained a much higher yield of oils than
photoautotrophic cells. Furthermore, oils from heterotrophic cells are more
132
Chapter 6. Differential lipid and fatty acid profiles of photoautotrophic and heterotrophic
Chlorella zofingiensis: assessment of algal oils for biodiesel prodoction
feasible for biodiesel production due to their high contents of NL and oleic acid.
The major drawback of using heterotrophic cultures for oils is the need of glucose
to feed the cells and in the meanwhile the emission of CO2. Glucose takes up
about 80% of total medium cost and depends on food-based agriculture (Li et al.,
2007). One strategy to overcome the drawback is to grow C. zofingiensis on the
sugars from industrial or agricultural waste and other “cellulosic” materials (Xu et
al., 2006; Jiang et al., 2009). It was found that cane molasses, a by-product of
sugar industry consisting of approximate 40-50% (w/w) of total sugars (Najafpour
& Poi Shan, 2003), could be ideally utilized by C. zofingiensis for lipid yield,
which will be described in next chapter.
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Chapter 7. Sugar-based growth and lipid accumulation for oil production in heterotrophic
Chlorella zofingiensis
Chapter 7
Sugar-based growth and lipid accumulation for oil
production in heterotrophic Chlorella zofingienesis
7.1 Abstract
The lipid production and fatty acid profile of Chlorella zofingiensis
cultured in the dark with various carbon sources were investigated. Of the sugars
surveyed, glucose was found to be the best one for the growth and lipid
production. When cultivated with 50 g L-1 of glucose, C. zofingiensis accumulated
lipids up to 52% of the dry weight, with triacylglycerol (TAG) accounting for
72.1% of the total lipids. Fatty acid profiles revealed that glucose contributed to
the highest yield of total fatty acids (TFA) and proportion of oleic acid (35.7% of
TFA), which corresponded to the strongest up-regulation of biotin carboxylase
(BC) and stearoyl ACP desaturase (SAD) genes. In fed-batch cultivation based on
glucose, the lipid yield and productivity of C. zofingiensis were further increased
to 20.7 g L-1 and 1.38 g d-1 L-1 respectively, representing around 3.9-fold of those
achieved in batch culture. Moreover, the low cost cane molasses was surveyed and
proved to be an ideal carbon source for boosting lipid production by C.
zofingensis. These results suggest that C. zofingiensis has great potential for
biodiesel production.
134
Chapter 7. Sugar-based growth and lipid accumulation for oil production in heterotrophic
Chlorella zofingiensis
7.2 Introduction
Chapter 6 described the superiority of heterotrophically grown C.
zofingiensis cells over photoautotrophically grown ones as biodiesel feedstocks. In
this chapter, to further explore the potential and economical feasibility of
heterotrophic C. zofingiensis as a biodiesel feedstock, various organic carbon
sources were surveyed. Firstly, two disaccharides (lactose and sucrose) and four
monosaccharides (galactose, fructose, mannose and glucose) were employed to
test the effect of sugars on heterotrophic growth and lipid production of C.
zofingiensis. Secondly, cane molasses was pretreated and adopted to develop a
cost-effective culture medium for lipid production by C. zofingiensis in
heterotrophic growth mode. Cane molasses is a byproduct of the sugar industry,
consisting of approximately 50% (w/w) total sugars (sucrose, glucose and
fructose), water, suspended colloids, heavy metals, vitamins and nitrogenous
compounds, etc. (Najafpour & Poi Shan, 2003). It is a low-cost raw material,
readily available and has been widely used to feed bacteria or yeast for the
production of various industrial important chemicals (Quesada-Chanto et al., 1994;
Sharma et al., 2008; Liu et al., 2009b; Jiang et al., 2009). This chapter, for the first
time, reported the economical production of lipids by C. zofingiensis from
pretreated cane molasses. The regulation of two key genes involved in fatty acid
biosynthesis was also surveyed in order to deepen our fundamental understanding
of biodiesel formation.
7.3 Methods and materials
7.3.1 Algal strain and culture conditions
The maintenance and inoculation of C. zofingiensis were described in
2.3.1.
135
Chapter 7. Sugar-based growth and lipid accumulation for oil production in heterotrophic
Chlorella zofingiensis
7.3.2 Pretreatment of molasses
Cane molasses was obtained from Jiang-men sugar-refinery (Guangdong,
PRC), and it contained 35% (w/w) sucrose, 10% (w/w) converted sugars (glucose
and fructose), 2.5% (w/w) other carbohydrates, 4.3% (w/w) crude protein, 0.06%
(w/w) crude fat, 9.6% (w/w) ash, 4.6% (w/w) salt, 8.9% (w/w) metal ions such as
calcium, potassium, sodium, iron, magnesium, copper, etc., and 25% (w/w) water.
For sulfuric acid treatment to remove cations, the molasses solution was adjusted
to pH 3.5 with 5 M H2SO4, and heated at 60 °C for 2 h; after centrifugation at
10,000 g for 15 min, the supernatant was adjusted to pH 6.5 with 10 M NaOH
(Jiang et al., 2009). For absorption treatment, the activated charcoal (Sigma) was
added to the molasses solution and the mixture was heated with continuous
stirring at 50 °C for 15 min; activated charcoal was removed by filtration and the
process was repeated until the solution was colorless (Kotzamanidis et al., 2002).
7.3.3 Batch and fed-batch culture
For batch culture, flasks, each containing 90 mL medium supplemented
with various carbon sources (lactose, galactose, sucrose, fructose, mannose,
glucose and molasses), were inoculated with 10% (v/v) of exponentially growing
inoculum and then incubated at 25 °C in an orbital shaker at 150 rpm in the dark.
For fed-batch culture, a two-stage fed-batch strategy was adopted using a 3.7-L
fermenter (Bioengineering Ag, Wald, Switzerland), as previously described (Sun
et al., 2008). The working volume of the fermentor was 3.0 L. The cultivation
conditions in the fermenter were controlled as follows: pH 6.5; temperature 25 °C;
agitation 450 rpm; and dissolved oxygen concentration at 50% saturation. During
fed-batch cultivation, the sterilized stock nutrient solution was fed into the
fermenter to maintain the sugar concentration at 5-20 g L-1.
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Chapter 7. Sugar-based growth and lipid accumulation for oil production in heterotrophic
Chlorella zofingiensis
7.3.4 Determination of glucose and nitrate concentration, cell dry
weight and specific growth rate
Determination of glucose and nitrate concentration, cell dry weight and
specific growth rate were described in 6.3.2.
7.3.5 Lipid extraction and analysis
Lipid extraction and analysis were described in 6.3.3.
7.3.6 Fatty acid analysis
Fatty acid analysis was described in 6.3.4.
7.3.7 RNA isolation and RT-PCR assay
RNA isolated and reverse transcription were described in 2.3.5. PCR
amplification was carried out using specific primers of biotin carboxylase (BC)
and stearoyl ACP-desaturase (SAD) genes (Table 7.1). C. zofingiensis actin (ACT)
primers were used to demonstrate equal amounts of templates and loading. The
GenBank accession numbers for BC and SAD were GQ996717 and GQ996719,
respectively. Amplification was done by conventional PCR [94 °C for 2 min
followed by 24 cycles (for ACT gene) or 26 cycles (for BC and SAD genes) of
94 °C for 15 s, 58 °C for 20 s, 72 °C for 30 s].
137
Chapter 7. Sugar-based growth and lipid accumulation for oil production in heterotrophic
Chlorella zofingiensis
Table 7.1 Primer sets for gene expression by RT-PCR
Gene
BC
Forward
Reverse
SAD
Forward
Reverse
ACT
Forward
Reverse
Primer (5'-3')
GTGCGATTGGGTATGTGGGGGTG
CGACCAGGACCAGGGCGGAAAT
TCCAGGAACGTGCCACCAAG
GCGCCCTGTCTTGCCCTCATG
TGCCGAGCGTGAAATTGTGAG
CGTGAATGCCAGCAGCCTCCA
7.4 Results
7.4.1 Heterotrophic growth and lipid production of C. zofingiensis
with various carbon sources
The lipid and fatty acid profiles of heterotrophic C. zofingiensis were
reported in chapter 6. To investigate the influences of different sugars on growth
and lipid production of C. zofingiensis, the algal cells were cultured in the dark
with lactose, galactose, sucrose, fructose, mannose or glucose as the carbon
source, respectively. Among the tested sugars, glucose gave the highest growth
rate (0.03 h-1), cell biomass (10.1 g L-1), lipid content (0.52 g g-1), and lipid yield
(5.27 g L-1). Algal cells cultured with mannose, fructose, or sucrose produced
slightly lower amounts of cell biomass and lipids. In contrast, lactose and
galactose were observed to be poor carbon sources for lipid production since low
biomass and lipid contents were obtained (Figure 7.1). These findings
demonstrated a close relationship between lipid biosynthesis and cell growth for
the heterotrophic alga.
138
Chapter 7. Sugar-based growth and lipid accumulation for oil production in heterotrophic
Chlorella zofingiensis
Glc
Figure 7.1 Cell biomass (black column), lipid content (gray column) and lipid
yield (white column) of C. zofingiensis cultured with 50 g L-1 various sugars. Lac,
lactose; Gal, galactose; Suc, sucrose; Fru, fructose; Man, mannose; Glc, glucose.
As shown in Figure 7.2, based on lipid class analysis of C. zofingiensis
cultured with glucose, neutral lipids (NL) were found to be the major constituent
that accounted for 80.9% of the total lipids, while phospholipids (PL) and
glycolipids (GL) together accounted for 19.1%. In NL, TAG were the
predominant component, accounting for 72.1% of the total lipids. In the batch
culture, up to 0.375 g g-1 TAG were accumulated in heterotrophic C. zofingiensis
cells. Unlike PL and GL that are membrane-bounded lipids, TAG serve primarily
as a storage form of carbon and energy (Hu et al., 2008). In addition, TAG are
superior to phospholipids (PL) or glycolipids (GL) for biodiesel due to their
higher content of fatty acids (Pruvost et al., 2009). Hence C. zofingiensis may act
as a promising host for biodiesel production due to its fast cell growth and high
content of TAG.
139
Chapter 7. Sugar-based growth and lipid accumulation for oil production in heterotrophic
Chlorella zofingiensis
Figure 7.2 Distribution of NL, GL and PL in total lipids extracted from
heterotrophic C. zofingiensis cells cultured with 50 g L-1 of glucose for 14 days.
The horizontal line inside the NL bar marks the portion of TAG in this fraction.
7.4.2 Fatty acid profiles of dark-grown C. zofingiensis cultures
Fatty acid composition considerably influences the properties of biodiesel
such as cetane number, heat of combustion, oxidative stability, cloud point,
lubricity, which finally influences the quality of biodiesel (Knothe, 2009). Here
the fatty acid compositions of the heterotrophically cultured C. zofingiensis with
various sugars were investigated (Table 7.2). It was found that C16:0, C16:2,
C18:1, C18:2 and C18:3 (n-3) were the major fatty acids, which together
accounted for more than 84% of the total fatty acids (TFA). The highest amounts
of TFA (45.4% of dry weight) and oleic acid (35.7% of TFA) were achieved in C.
zofingiensis cultured with glucose as the carbon source, which were about 5 and 3
times respectively of those given by lactose.
As an ideal biodiesel, the fatty acid esters should be oxidative and
low-temperature stable. Generally saturated fatty acid esters are oxidative stable,
while unsaturated fatty acid esters give low-temperature stability (Knothe, 2008).
The enhanced proportion of oleic acid ester in the total fatty acid esters has been
140
Chapter 7. Sugar-based growth and lipid accumulation for oil production in heterotrophic
Chlorella zofingiensis
considered as a feasible approach to balance the oxidative and low-temperature
stability with retaining the cetane number at an acceptable level (Knothe, 2009).
The high content of oleic acid in the heterotrophic cells further supports that C.
zofingiensis is a favorable host for producing high quality biodiesel.
Table 7.2 Fatty acid profiles of dark-grown C. zofingiensis cultured with 50 g L-1
various sugars for 14 days a
Fatty acids
Sugars
Lactose
Galactose
Sucrose
Fructose
Mannose
Glucose
C16:0
28.73 ± 1.22
27.81 ± 0.99
23.16 ± 0.83
23.62 ± 1.12
23.38 ± 1.03
22.62 ± 0.77
C16:1
3.42 ± 0.12
2.52 ± 0.13
1.49 ± 0.06
1.71 ± 0.08
1.49 ± 0.05
1.97 ± 0.08
C16:2
9.02 ± 0.35
11.08 ± 0.61
8.48 ± 0.39
8.31 ± 0.44
7.94 ± 0.29
7.38 ± 0.33
C16:3
5.28 ± 0.27
2.52 ± 0.11
1.96 ± 0.07
2.12 ± 0.09
1.83 ±0.06
1.94 ± 0.04
C16:4
0.92 ± 0.05
0.65 ± 0.03
0.23 ± 0.02
0.17 ± 0.01
0.19 ± 0.01
0.22 ± 0.02
C18:0
0.95 ± 0.06
1.61 ± 0.09
2.80 ± 0.09
2.34 ± 0.12
2.44 ± 0.10
2.09 ± 0.08
C18:1
12.59 ± 0.45
25.15 ± 0.89
31.99 ± 1.21
32.06 ± 0.93
33.30 ± 1.45
35.68 ± 1.23
C18:2
20.87 ± 0.87
16.48 ± 0.83
19.51 ± 0.75
19.27 ± 0.49
19.36 ± 0.98
18.46 ± 0.66
C18:3 (n-6)
1.39 ± 0.03
1.17 ± 0.05
0.55 ± 0.01
0.52 ± 0.03
0.52 ± 0.03
0.51 ± 0.02
C18:3 (n-3)
12.86 ± 0.39
7.74 ± 0.31
7.36 ± 0.35
7.48 ± 0.18
7.31 ± 0.23
7.24 ± 0.30
C18:4
1.24 ± 0.04
1.10 ± 0.06
0.46 ± 0.01
0.45 ± 0.01
0.45 ± 0.02
0.49 ± 0.02
Others
2.72 ± 0.09
2.17 ± 0.11
2.00 ± 0.07
1.94 ± 0.04
1.78 ± 0.08
1.40 ± 0.08
MUFAb (%)
16.01 ± 0.55
27.67 ± 0.82
33.48 ± 1.12
33.78 ± 1.03
34.79 ± 1.37
37.64 ± 1.26
PUFAc (%)
51.59 ± 2.11
40.74 ± 2.05
38.57 ± 1.28
38.33 ± 1.49
37.61 ± 1.44
36.24 ± 1.63
UFAd (%)
67.59 ± 2.95
68.42 ± 2.31
72.05 ± 3.72
72.10 ± 3.11
72.40 ± 3.13
73.89 ± 2.99
▽/mol e
1.43 ± 0.05
1.24 ± 0.06
1.22 ± 0.06
1.22 ± 0.03
1.21 ± 0.03
1.21 ± 0.05
TFA f
8.48 ± 0.33
20.30 ± 0.71
40.48 ± 1.29
43.01 ± 1.11
43.10 ± 1.79
45.38 ± 1.83
a
Data are expressed as mean ± standard deviation of triplicates. b MUFA,
monounsaturated fatty acids; c PUFA, polyunsaturated fatty acids; d UFA,
unsaturated fatty acids; e ▽/mol, degree of fatty acid unsaturation = [1.0 (%
monoenes) + 2.0 (% dienes) + 3.0 (% trienes) + 4.0 (% tetraenes) ]/100; f TFA,
total fatty acids (g) / cell dry weight (g) × 100%.
141
Chapter 7. Sugar-based growth and lipid accumulation for oil production in heterotrophic
Chlorella zofingiensis
7.4.3 Sugars up-regulate the transcription of BC and SAD genes of
C. zofingiensis
The de novo biosynthesis of fatty acid is initially catalyzed in chloroplast
by the key enzyme acetyl-CoA carboxylase (ACCase); while the introduction of
the first double bond to acyl chain is carried out by stearoyl ACP (SAD), the
enzyme playing an important role in determining the ratio of unsaturated and
saturated fatty acids (Hu et al., 2008). To reveal the relationship between the fatty
acid profiles and the regulation of ACCase and SAD, we investigated the
transcript levels of SAD and biotin carboxylase (BC, a subunit of ACCase) in
dark-grown C. zofingiensis cells cultured with various sugars using the RT-PCR
approach. The transcription of the two genes was shown to be differentially
regulated by various carbon sources (Figure 7.3). In coincidence with the higher
contents of TFA in the algal cells cultured with sucrose, fructose, mannose or
glucose, the transcript levels of BC in the cells were strongly up-regulated. In
contrast, galactose moderately enhanced the transcription of BC whereas lactose,
the poor carbon source for the cell growth and fatty acid accumulation, caused a
slight up-regulation of BC. These results are consistent with the findings of
Ohlrogge and Jaworski (1997) that the first reaction step of fatty acid biosynthesis
catalyzed by ACCase is a major point of flux control for this pathway. A similar
pattern induced by sugars was also found for SAD at mRNA levels, although
much stronger up-regulation was observed when compared with the expression of
BC (Figure 7.3). Similarly, lactose induced low-level expression of SAD while
glucose triggered high amounts of SAD transcripts, which is consistent with the
unsaturation values of fatty acids in the algal cells cultured with the tested sugars
(Table 7.2).
142
Chapter 7. Sugar-based growth and lipid accumulation for oil production in heterotrophic
Chlorella zofingiensis
Figure 7.3 Transcript levels biotin carboxylase (BC) and stearoyl ACP desaturae
(SAD) genes in heterotrophic C. zofingiensis cultured with 50 g L-1 of various
sugars. The expression of actin (ACT) gene was used as control. Ctr, control
cultured without sugar. Abbreviations see Figure 7.1.
7.4.4 Fed-batch fermentation enhances lipid production by C.
zofingiensis
As shown by the above results, the productivity of lipid is closely related
to the growth rate, cell density and cellular lipid content. Compared with batch
culture, fed-batch cultivation mode can extend the exponential growth phase of
the cell and further increase the biomass concentration fed with key medium
components or fresh medium at time intervals. It was indicated that C.
zofingiensis could reach a high specific growth rate (ca. 0.03 h-1) and a high
cell-density (ca. 53 g L-1) in a fed-batch culture system (Sun et al., 2008). To
further evaluate the lipid production of C. zofingiensis, a 3.7-L fermenter was used
to investigate the effect of glucose on cell growth and lipid accumulation in a
fed-batch culture system. The fed-batch procedure contained two stages of feeding:
three times of feeding with glucose-containing medium and four times of feeding
with glucose. The feeding time, time course of cell growth and lipid production
are shown in Figure 7.4. In the fed-batch fermentation a high lipid yield of 20.7 g
L-1 was obtained, with a lipid productivity of 1.38 g d-1 L-1. At the stage-fed with
glucose-containing medium, the algal cells grew fast but the cellular lipid content
143
Chapter 7. Sugar-based growth and lipid accumulation for oil production in heterotrophic
Chlorella zofingiensis
was low (data not shown), possibly due to the relatively low carbon/nitrogen (C/N)
ratio as indicated by the remaining concentrations of glucose and nitrate in the
medium (Figure 7.4). When stage-fed with glucose, due to the consumption of
nitrogen and the new addition of glucose, a high C/N ratio formed in the culture
medium, resulting in a favorable condition for lipid accumulation within cells
(Figure 7.4). These results further support that C. zofingiensis has the potential to
be cultivated by using fermentation biotechnology for large-scale production of
lipids.
Figure 7.4 Cell biomass, glucose and nitrate consumption and lipid production in
a two-stage fed-batch fermentation of C. zofingiensis in a 3.7-L fermenter. (■)
glucose concentration; (○) cell biomass; (column) lipid content; (□) lipid yield;
(◇) NO3- concentration; (↓) glucose-containing medium feeding; (↓↓) glucose
feeding.
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Chapter 7. Sugar-based growth and lipid accumulation for oil production in heterotrophic
Chlorella zofingiensis
7.4.5 Assessment of cane molasses as the carbon source for lipid
production by C. zofingiensis
C. zofingiensis could efficiently utilize glucose, fructose and sucrose for
growth and lipid production (Figure 7.1). Thus, it is reasonable to expect that the
low-cost cane molasses can be used to feed C. zofingiensis for economical
production of algal oils. However, the untreated molasses gave relatively low cell
biomass (5.5 g L-1) and lipid content (0.32 g g-1) and thus the low lipid yield (1.76
g L-1) (Table 7.3). Molasses normally contains most of the essential nutrients for
the growth of microorganisms, but it also contains lots of metal ions and
suspended colloids, which could inhibit the growth of microorganisms and
contribute to the inactivation of certain enzymes associated with product
biosynthesis (Jiang et al., 2009). The pretreatment of cane molasses is therefore
necessary. Sulfuric acid treatment is regarded as an efficient way to remove
cations and has been widely used for molasses pretreatment (Roukas et al., 1998;
Kotzamanidis et al., 2002; Kalogiannis et al., 2003; Jiang et al., 2009). Compared
with the untreated molasses, the sulfuric acid treated one induced the significantly
higher cell biomass (10.8 g L-1) and lipid content (0.48 g g-1) and thus the much
greater lipid yield (5.18 g L-1) (Table 7.3). The absorption treatment using
charcoal to remove color was also tested, but proved to be less efficient than the
sulfuric acid treatment, as indicated by the lower lipid yield (Table 7.3). In
addition, the combination of the above two treatments was not as good as sulfuric
acid treatment alone for producing lipids by C. zofingiensis (Table 7.3).
The fed-batch fermentation of C zofingiensis using the sulfuric acid
treated molasses for lipid production was also examined. Compared with glucose,
the pretreated molasses gave a slightly higher cell biomass but lower cellular lipid
content (Table 7.4). This might be explained by that molasses contains many
essential nutrients and exerts positive effect on the growth of this green alga. The
lipid yield and sugar-based lipid yield coefficient obtained from C. zofingiensis
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Chapter 7. Sugar-based growth and lipid accumulation for oil production in heterotrophic
Chlorella zofingiensis
cultured with the pretreated molasses were 19.7 g L-1 and 0.239 respectively,
which are comparable to those obtained from the alga cultured with glucose
(Table 7.4). These results indicate that cane molasses, when pretreated with proper
methods, can substitute glucose to feed C. zofingiensis for economical production
of heterotrophic algal oils.
Table 7.3 Lipid production from pretreated molasses by C. zofingiensis in batch
culture systems a
Pretreatment of
Lipid yield
(g L-1)
Residual total Sugar
molasses
Cell biomass Lipid content
(g L-1)
(g g-1)
sugar (g L-1)
unitization (%)
Untreated
5.5 ± 0.29
0.32 ± 0.02
1.76 ± 0.10
15.8 ± 0.65
47.4 ± 2.5
Sulfuric acid
treatment
10.8 ± 0.43
0.48 ± 0.03
5.18 ± 0.22
4.6 ± 0.28
84.7 ± 3.3
Absorption
treatment
7.9 ± 0.28
0.39 ± 0.01
3.08 ± 0.17
11.7 ± 0.46
61.0 ± 3.5
Sulfuric acid +
absorption
9.1 ± 0.47
0.43 ± 0.02
3.91 ± 0.21
6.8 ± 0.39
77.3 ± 4.2
a
Data are expressed as mean ± standard deviation of triplicates.
Table 7.4 Lipid production from pretreated molasses by C. zofingiensis in
fed-batch fermenter systems a
Sugar
Cell biomass Lipid content
(g L-1)
(g g-1)
Lipid yield
(g L-1)
Consumed
sugar (g L-1)
Lipid yield
coefficient b
Glucose
42.5 ± 1.9
0.48 ± 0.02
20.7 ± 0.8
83.9 ± 4.5
0.243 ± 0.01
Pretreated
molasses
46.9 ± 2.3
0.42 ± 0.02
19.7 ± 1.2
82.3 ± 3.1
0.239 ± 0.02
a
Data are expressed as mean ± standard deviation of triplicates. b Lipid yield
coefficient = lipid yield (g L-1) / consumed sugar (g L-1).
7.5 Discussion
C. zofingiensis can grow heterotrophically using sugars as the carbon and
energy sources (Ip & Chen, 2005; Sun et al., 2008). In the current chapter, several
146
Chapter 7. Sugar-based growth and lipid accumulation for oil production in heterotrophic
Chlorella zofingiensis
sugars were tested for lipid production by heterotrophic C. zofingiensis cells.
Glucose, mannose, fructose and sucrose were shown to be suitable organic carbon
sources for boosting lipid yield (Figure 7.1). Glucose also induced a high
proportion of NL including TAG (Figure 7.2), which are regarded to be superior to
GL and PL when considered as biodiesel sources. Fatty acid profile is also the
important information for the assessment of biodiesel feedstocks. As shown in
Table 7.2, glucose-fed algal cells were lower in saturated fatty acids and higher in
oleic acid, which are considered as good for the balance between oxidative
stability and cold flow of oils and thus be able to promote the quality of biodiesel
(Knothe, 2009). What’s more, the high lipid yield (20.7 g L-1) and lipid
productivity (1.38 g d-1 L-1) were obtained by using a well designed fed-batch
fermentation strategy (Figure 7.4). Compared with other microalgae, C.
zofingiensis showed obvious advantages in terms of lipid content and lipid
productivity (Table 5). All these results together implicate that the heterotrophic C.
zofingiensis might have the potential to be a biodiesel feedstock.
Table 7.5 Lipid production by C. zofingiensis and other microalgae
Microalgal species
Lipid content
(% dry weight)
Lipid yield
(mg L-1 day-1)
References
C. zofingiensis
48.0
1380
This study
Chlorella protothecoides
14.6-57.8
1214
Chlorella emersonii
25.0-63.0
10.3-50.0
Chlorella sorokiniana
19.0-22.0
44.7
Chlorella vulgaris
5.0-58.0
11.2-40.0
Mata et al.,
Chlorococcum sp.
19.3
53.7
2010
Dunaliella salina
6.0-25.0
116.0
Nannochloropsis oculata
22.7-29.7
84.0-142.0
Neochloris oleoabundans
29.0-65.0
90.0-134.0
147
Chapter 7. Sugar-based growth and lipid accumulation for oil production in heterotrophic
Chlorella zofingiensis
Other microorganisms that grow only heterotrophically such as bacteria
and yeasts have also been explored for biodiesel sources. Bacteria are less used
for biodiesel production since they have low contents of oils and produce mostly
complex lipid (Kalscheuer et al., 2006; Gouda et al., 2008). Yeasts have the ability
of accumulating high levels of cellular lipids, ranging from 5.32 to 37.1 g L-1 (Li
et al., 2008c). Generally speaking, fermentation-based biodiesel production
contributes the emission of CO2, which might limit the application of the
microorganisms for biodiesel production to some extent. Unlike yeasts, C.
zofingiensis can grow well both photoautotrophically, mixotrophically and
heterotrophically. Thus, by switching its growth modes, C. zofingiensis may
release much less CO2 during the accumulation of lipids, as has been
demonstrated by Xiong et al. (2010).
The major drawback of using heterotrophic cultures for oils is the need of
glucose to feed the cells because glucose, which takes up about 60% of total cost,
depends on food-based agriculture. The current price of industrial glucose is
around 340 USD per ton. According to Table 7.4, the conversion ration of sugar to
lipid is about 24.3%. Thus the roughly estimated cost of one kilogram oil obtained
from C. zofingiensis fed with glucose is around $2.33, 3-4 times higher than plant
oil. One strategy to overcome the drawback is to grow C. zofingiensis on the
sugars from industrial or agricultural waste and other “cellulosic” materials (Xu et
al., 2006; Jiang et al., 2009). Cane molasses, a low-cost by-product of sugar
industry containing approximate 40-50% (w/w) of total sugars, were proved to be
an ideal carbon source utilized by C. zofingiensis for lipid production (Table 7.3
and 7.4). Currently, the price of cane molasses is around one fifth of glucose,
which means the cost of microalgal oil can be reduced to $1.52 per kilogram
when molasses replaces glucose as the sugar source. Moreover, C. zofingiensis
could yield high amount of the high-value astaxanthin under heterotrophic
conditions (Ip & Chen, 2005). Thus, by feeding algae with “waste” sugars and
tying algal oil production to other products (e.g., astaxanthin), profitable biodiesel
from microalgae won’t remain a research project for long.
148
Chapter 8. Production of fatty acids by heterotrophic Chlorella zofingiensis: influences of
nutritional and environmental factors
Chapter 8
Production of fatty acids by heterotrophic Chlorella
zofingiensis: influences of nutritional and environmental
factors
8.1 Abstract
Biodiesel is chemically referred to as fatty acid methyl esters. This
chapter surveyed the effect of several nutrients, namely, nitrate, carbon/nitrogen
(C/N) ratio, phosphate and ferrous ion as well as the environmental conditions
including temperature and the initial pH of culture medium on the growth and
fatty acid profile of heterotrophic C. zofingiensis in batch culture. Results showed
that cell growth was promoted at sufficient nitrogen while the accumulation of
total fatty acids (TFA) was enhanced under nitrogen starvation/deprivation. A high
C/N ratio (e.g., 200) was found to be beneficial for the accumulation of TFA
including oleic acid (C18:1). Similar to nitrogen, phosphorus favored cell growth
at sufficient concentrations but induced TFA accumulation at starved/deprived
concentrations. Ferrous ion was required in a low concentration (25 μM) to
maximize cell growth and TFA accumulation. The alga grew well at a wide range
of temperatures (25-30 °C) and initial pH (4.5-8.5) of the culture medium for TFA
accumulation. The optimal nutritional and environmental conditions for TFA
production by C. zofingiensis were 5 mM nitrate, 5 mM phosphate, 25-100 μM
ferrous ion, cultivation temperature of 25 °C and the initial pH of 6.5. The results
of this study may serve as a guide to a large scale cultivation of C. zofingiensis
and possible other algae for biodiesel.
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Chapter 8. Production of fatty acids by heterotrophic Chlorella zofingiensis: influences of
nutritional and environmental factors
8.2 Introduction
Biodiesel, also known as fatty acid methyl esters, is derived from the
transesterification of feedstocks. Thus the yield and quality of biodiesel from a
feedstock are largely determined by the TFA content and fatty acid properties of
the feedstock. Microalgae have been considered as the most potential feedstocks
for biodiesel production (Chisti, 2007; Huang et al., 2010; Mata et al., 2010). The
fatty acid profiles of microalgae are species/strain-specific and can be greatly
affected by the culture conditions, e.g., nutritional factors and environmental
conditions, as reviewed in 1.3.5.2. Chapter 6 and 7 described the potential of
heterotrophic C. zofingiensis as a biodiesel feedstock. To further explore this
potential, in this study, the effect of several important factors including nitrate,
phosphate, ferrous ion, cultivation temperature and the initial pH of medium on
the growth and fatty acid profile of heterotrophic C. zofingiensis in batch culture
were surveyed.
8.3 Methods and materials
8.3.1 Algal strain and culture conditions
The maintenance of C. zofingiensis was described in 2.3.1. In the
experiments, the alga was grown in the medium supplemented with 30 g L-1
glucose unless otherwise stated. To investigate the effect of nutrient concentration,
nitrate and phosphate were tested at 0-10 mM; ferrous ion was used at 0-100 μM.
To examine the effect of C/N ratio, glucose was fixed at three different
concentrations of 10 g L-1, 30 g L-1 and 50 g L-1, with four different C/N ratios of
50, 100, 200 and 400 respectively. To survey the effect of environments,
cultivation temperatures (22-35 °C) and the initial pHs (4.5-8.5) were employed.
Unless otherwise stated, the media were adjusted to pH 6.5 prior to autoclaving at
150
Chapter 8. Production of fatty acids by heterotrophic Chlorella zofingiensis: influences of
nutritional and environmental factors
121 °C for 20 min. An inoculum of 10% (by volume, average cell concentration of
0.5 g L-1 dry weight) was inoculated into 250-ml flasks each containing 100 ml
medium under dim light. The cultures were then incubated with orbital shaking at
150 rpm in the dark for 14 days.
8.3.2 Determination of cell dry weight
The determination of cell dry weight was described in 6.3.2.
8.3.3 Fatty acid analysis
Fatty acid analysis was described in 6.3.4.
8.3.4 Statistical analysis
Statistical analyses of TFA yield were performed using the SPSS
statistical package. One way ANOVA test was applied. The statistical
significances were achieved when P < 0.05.
8.4 Results and discussion
8.4.1 Nitrate
Nitrogen is one of the most critical nutrients that affect cell growth and
lipid/fatty acid metabolism of microalgae. The heterotrophic growth and fatty acid
profile of C. zofingiensis at various nitrate concentrations were analyzed and
shown in Figure 8.1. The algal cell growth was retarded at low nitrate
concentrations (0-1.25 mM) and got promoted at high nitrate concentrations (5-10
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Chapter 8. Production of fatty acids by heterotrophic Chlorella zofingiensis: influences of
nutritional and environmental factors
mM) (Figure 8.1A). In contrast, the cellular TFA content was enhanced at low
nitrate concentrations (Figure 8.1A), which is consistent with previous studies
showing that nitrogen starvation/deficiency favored the accumulation of lipids and
TFA (Takagi et al., 2000; Li et al., 2005; Li et al., 2008a; Solovchenko et al., 2008;
Pruvost et al., 2009). One way ANOVA test showed that the effect of nitration
concentration on TFA yield was significant. The highest TFA yield was obtained
at 5 mM nitrate (Figure 8.1A). Nitrate concentration also affected the algal fatty
acid composition. As shown in Figure 8.1B, low nitrate concentrations enhanced
the proportion of monounsaturated fatty acids (MUFA) including oleic acid
(C18:1) but reduced the proportion of polyunsaturated fatty acids (PUFA).
Figure 8.1 Effect of nitrate concentration on growth and fatty acid profile of
heterotrophic C. zofingiensis.
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Chapter 8. Production of fatty acids by heterotrophic Chlorella zofingiensis: influences of
nutritional and environmental factors
The effect of nitrate with different glucose concentrations (10, 30 and 50
g L-1) was also investigated. The C/N ratios were fixed at 50, 100, 200 and 400.
Under all glucose concentrations surveyed, high C/N ratios benefited the
accumulation of TFA (Figure 8.2A, B and C, left) and increased the proportion of
MUFA in particular oleic acid (Figure 8.2A, B and C, right). A C/N ratio of 200
was found to enhance TFA content (Figure 8.2) and maintain cell biomass (data
not shown)
Figure 8.2 Effect of initial C/N ratio on TFA content (right) and fatty acid
composition (left) of heterotrophic C. zofingiensis. Glucose concentrations were
fixed at 10 g L-1 (A), 30 g L-1 (B) and 50 g L-1 (C).
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Chapter 8. Production of fatty acids by heterotrophic Chlorella zofingiensis: influences of
nutritional and environmental factors
8.4.2 Phosphate
Figure 8.3 illustrates the effect of phosphate concentrations on the cell
growth and fatty acid profile of heterotrophic C. zofingiensis. Low concentrations
of phosphate restricted cell growth but promoted cellular TFA content (Figure
8.3A). Unlike nitrate, however, phosphate deprivation did not greatly inhibit the
cell growth (Figure 8.3B). One way ANOVA test showed that the effect of
phosphate concentration on TFA yield was not significant The TFA yield reached
its maximum at 5 mM phosphate. Consistent with previous studies (Reitan et al.,
1994), phosphate sufficiency gave rise to the decreased proportion of MUFA
including oleic acid but the increased PUFA proportion in C. zofingiensis (Figure
8.3B).
Figure 8.3 Effect of phosphate concentration on growth and fatty acid profile of
heterotrophic C. zofingiensis.
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Chapter 8. Production of fatty acids by heterotrophic Chlorella zofingiensis: influences of
nutritional and environmental factors
8.4.3 Ferrous ion
A significant limitation on the cell growth and TFA content of C.
zofingiensis was occurred in the absence of iron (Figure 8.4A). Supplementation
of ferrous ion at 25-100 μM markedly promoted the cell growth and cellular TFA
content (Figure 8.4A). Similar results were also observed for the green alga
Chlorella vulgaris (Liu et al., 2008). One way ANOVA test showed that the effect
of ferrious concentration on TFA yield was significant. The increased
concentration of ferrous ion resulted in the increased proportion of MUFA
including oleic acid and decreased PUFA proportion, but almost unchanged
proportion of saturated fatty acids (SFA) (Figure 4B).
Figure 8.4 Effect of ferrous ion concentration on growth and fatty acid profile of
heterotrophic C. zofingiensis.
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Chapter 8. Production of fatty acids by heterotrophic Chlorella zofingiensis: influences of
nutritional and environmental factors
8.4.4 Cultivation temperature
C. zofingiensis grew well at the temperature of 22-28 °C; higher
temperature (e.g., 32 and 35 °C) dramatically reduced the cell biomass and
cellular TFA content (Figure 8.5A). Of the temperature range surveyed, 25 °C was
the best temperature for the cell growth and TFA accumulation and gave rise to
the highest TFA yield (Figure 8.5A). One way ANOVA test showed that the effect
of cultivation temperature on TFA yield was significant. The effect of temperature
on fatty acid composition was also investigated and shown in Figure 8.5B. Higher
temperatures (28-35 °C) caused the greater proportion of SFA and the lower
proportion of MUFA including oleic acid. This is consistent with the previous
studies suggesting that increased temperature resulted in increased fatty acid
saturation and the concurrently decreased fatty acid unsaturation (Wada & Murata,
1990; Thompson et al., 1992; Renaud et al., 2002; Jang et al., 2005).
Figure 8.5 Effect of cultivation temperature on growth and fatty acid profile of
heterotrophic C. zofingiensis.
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Chapter 8. Production of fatty acids by heterotrophic Chlorella zofingiensis: influences of
nutritional and environmental factors
8.4.5 Initial pH of culture medium
Figure 8.6 shows the cell growth and fatty acid profile of C. zofingiensis
in the batch culture with various initial pHs. The alga grew well at the entire range
of pH surveyed. The highest cell biomass and cellular TFA content were obtained
at pH 6.5, so was the TFA yield (Figure 8.6A). One way ANOVA test showed that
the effect of nitration concentration on TFA yield was not significant. pH at the
range of 5.5-6.5 gave rise to a slight increase in the proportion of MUFA including
oleic acid and a slight decrease in PUFA proportion (Figure 8.6B).
Figure 8.6 Effect of culture medium pH on growth and fatty acid profile of
heterotrophic C. zofingiensis.
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Chapter 8. Production of fatty acids by heterotrophic Chlorella zofingiensis: influences of
nutritional and environmental factors
In conclusion, the effect of several key factors including nitrate, C/N ratio,
phosphate, ferrous ion, cultivation temperature and the culture medium pH on the
cell growth and fatty acid profile of heterotrophic C. zofingiensis in the batch
culture was investigated in this study. The optimal nutritional and environmental
conditions for heterotrophic C. zofingiensis producing TFA were 5 mM nitrate, 5
mM phosphate, 25-100 μM ferrous ion, cultivation temperature of 25 °C and the
initial pH of 6.5.
158
Chapter 9. Isolation and characterization of biotin carboxylase gene and stearoyl ACP
desaturase gene from Chlorella zofingiensis
Chapter 9
Isolation and characterization of biotin carboxylase gene
and stearoyl ACP desaturase gene from Chlorella
zofingiensis
9.1 Abstract
Chlorella zofingiensis could accumulate abundant oil within cells. In
order to elucidate the molecular regulation of fatty acid accumulation, the key
genes involved in fatty acid biosynthesis need to be characterized. In the present
study, two key genes, namely biotin carboxylase (BC) and stearoyl ACP
desaturase (SAD) were cloned and characterized from C. zofingiensis. Protein
sequence alignments showed that the deduced amino acid sequences of these two
genes shared high identity to their corresponding genes from other microalgae and
plants. High light illumination and glucose were revealed to up-regulate the
expression of both BC and SAD genes, and therefore enhanced the accumulation
of total fatty acids (TFA) including oleic acid (C18:1). Unlike high light or
glucose, salt stress, however, could only trigger the up-regulation of SAD gene;
accordingly, the TFA content was slightly affected while the biosynthesis of oleic
acid was promoted. In this context, the fatty acid biosynthesis of C. zofingiensis
might be regulated, at least partly, at the transcriptional level. These results deepen
our understanding of fatty acid biosynthesis in C. zofingiensis, which can facilitate
the manipulation of the alga for enhanced production of oils.
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Chapter 9. Isolation and characterization of biotin carboxylase gene and stearoyl ACP
desaturase gene from Chlorella zofingiensis
9.2 Introduction
Similar to plants, microalgae synthesize fatty acids in the chloroplast
using a single set of enzymes, of which acetyl-CoA carboxylase (ACCase) is the
key one in regulating fatty acid synthesis rate while stearoyl ACP desaturase
(SAD) adds the first double bond to acyl chain and plays an important role in
determining the ratio of unsaturated fatty acids to saturated ones (Ohlrogge &
Jaworski, 1997). Although the fatty acid composition has been shown to be
regulated by the expression of its biosynthesis genes in plants, little is known with
microalgae in this regard (Hu et al., 2008). To elucidate the regulation of fatty acid
accumulation in microalgae, key genes need to be firstly cloned.
The production potential of biodiesel using the green microalga C.
zofingiensis as a feedstock was investigated in chapter 6 and 7. C. zofingiensis
could produce fatty acids up to 45% of cell dry weight, with 36% of TFA being
oleic acid. In the present study, two important genes encoding biotin carboxylase
(BC, one subunit of ACCase) and SAD were cloned and characterized from C.
zofingiensis, with the aim of examining their expressions during fatty acid
biosynthesis in response to high light, salt stress or glucose. These results expand
our knowledge about the fatty acid biosynthesis in C. zofingiensis and benefit
enhanced fatty acid production or modified fatty acid composition through genetic
engineering for better biodiesel production.
9.3 Methods and materials
9.3.1 Algal strain and culture conditions
The maintenance of C. zofingiensis was described in 2.3.1. To investigate
the expression of biotin carboxylase (BC) and stearoyl ACP-desaturase (SAD)
genes and the biosynthesis of fatty acids under different conditions, the algal cell
160
Chapter 9. Isolation and characterization of biotin carboxylase gene and stearoyl ACP
desaturase gene from Chlorella zofingiensis
cultures were exposed to continuous illumination of high light (250 µmol photon
m-2 s-1), maintained in the dark treated with 100 mM NaCl or 50 mM glucose for
further periods.
9.3.2 Genomic DNA and RNA isolation
Genomic DNA and RNA isolation were described in 2.3.2.
9.3.3 Cloning of BC cDNA, SAD cDNA and their corresponding
genes
The primer sets used in this study are listed in Table 9.1. To amplify the
BC cDNA from C. zofingiensis, degenerate primers BC-dF and BC-dR were
designed according to the conserved motifs GGGGRGM and FYFMEMNT in BC
proteins from Chlamydomonas reinhardtii, Arabidopsis thaliana, Nicotiana
tabacum and Brassica napus. First strand cDNA synthesis was carried out with 1
μg of total RNA extracted from 24 h high light induced cells, by using a
SuperScript III First-Strand Synthesis System according to manufacturer’s
instruction (Invitrogen). PCR amplification was programmed with 50 ng of cDNA
as template (30 cycles of 94 °C for 30 s, 55 °C for 20 s, 72 °C for 1 min). PCR
product was gel purified and sequenced, based on which specific primers (BC-F1
and BC-R2 for first round PCR, BC-F2 and BC-R2 for second round PCR) were
designed for rapid amplification of 5' and 3' cDNA ends (RACE). RACE was
performed by using the method described by Huang and Chen (2006). The primer
pair BC-F3 and BC-R3 derived from the sequences of 5' and 3' RACE fragments
were employed to amplify a full-length BC cDNA and its corresponding gene.
Similar procedures were followed for SAD cDNA amplification. The
degenerated primers SAD-dF and SAD-dR were derived from the conserved
161
Chapter 9. Isolation and characterization of biotin carboxylase gene and stearoyl ACP
desaturase gene from Chlorella zofingiensis
amino acid sequences (GDM/LITEEA and HGNTARQ/HA, respectively) of the
SAD proteins from A. thaliana, B. napus, Helianthus annuus, C. reinhardtii and
Haematococcus pluvialis. The primers SAD-F1, SAD-R1, SAD-F2 and SAD-R2
were used for 5' and 3' SAD RACE, while SAD-F3 and SAD-R3 were employed
for amplification of the full-length SAD cDNA and its corresponding gene.
Table1 Primers for gene cloning and expression
Aim
Oligonucleotide sequence 5'-3'
Partial BC fragment
BC-dF
GGCGGCGGCGGNMGNGGNATG
BC-dR
GTRTTCATYTCCATRAARTARAA
5' and 3' BC RACE
BC-F1
CGCCCTCACCCGCCCTTACAC
BC-R1
CCACATTGCCATACTTGTCAGC
BC-F2
GTGCGATTGGGTATGTGGGGGTG
BC-R2
CCATACTTGTCAGCCAGCACC
BC gene
BC-F3
CCTCTGCCTACATGCCCAGCTG
BC-R3
CTCAGCTGGTGCCTCATGCC
BC expression
BC-F2 & BC-R4
see BC-F2, CGACCAGGACCAGGGCGGAAAT
Partial SAD fragment
SAD-dF
GGCGAYWTGATHACNGARGARGC
SAD-dR
GCNTGNCKNGCNGTRTTNCCRTG
5' and 3' SAD RACE
SAD-F1
GGGTTTCATCTACACCTCCTTC
SAD-R1
TGTTCTCCTCAGCCGTCCACTCAC
SAD-F2
TCCAGGAACGTGCCACCAAG
SAD-R2
CACGCGTCCACCTGCCCCAAG
SAD gene
SAD-F3
CTCCTACTTTGGCACACTCAGC
SAD-R3
GCTTTCAATCTAGCTACTGTCG
SAD expression
SAD-F2 & SAD-R4
see SAD-F2, GCGCCCTGTCTTGCCCTCATG
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Chapter 9. Isolation and characterization of biotin carboxylase gene and stearoyl ACP
desaturase gene from Chlorella zofingiensis
9.3.4 RT-PCR assay
RT-PCR assay of BC, SAD and actin genes was described in 7.3.7.
9.3.5 Fatty acid analysis
Fatty acid analysis was described in 6.3.4.
9.4 Results
9.4.1 Cloning and characterization of the BC and SAD gene from
C. zofingiensis
A 380 bp fragment of BC cDNA was amplified by RT-PCR with the
degenerate primers BC-dF and BC-dR. On the basis of this sequence information,
two pairs of specific primers (BC-F1 and BC-R1, BC-F2 and BC-R2) were
designed for 5' and 3' RACE, which generated a 1.7 kb fragment. The sequence of
the fragment was determined as a fusion of the 5' and 3' ends of a putative BC
cDNA. The coding region of BC gene was amplified using primers BC-F3 and
BC-R3, which contains 1,743 bp encoding a deduced BC protein of 490 amino
acid residues (GenBank accession No. GQ996717). Upstream of the translation
start codon is a 107-bp 5' untranslated region and between the stop codon and poly
(A) tail is a 3' untranslated region of 351 bp nucleotides. TGTAA, the sequence
considered as a potential polyadenylation signal in green algae (Schmitt et al,
1992), is also found in the 3' untranslated region of the BC gene. The GC content
of the BC coding region is 52.9%, which is much lower than GC content of BC
genes from C. reinhardtii (67.8%) and H. pluvialis (63.7%). Protein sequence
alignments showed that the BC of C. zofingiensis shared a high identity to that of
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Chapter 9. Isolation and characterization of biotin carboxylase gene and stearoyl ACP
desaturase gene from Chlorella zofingiensis
other algae and higher plants, especially to the BC of H. pluvialis (74%) (Figure
9.1). The phylogenetic analysis indicates that the algal BC proteins are more
related to those from plants (Figure 9.2).
Figure 9.1 Amino acid sequence alignment of C. zofingiensis BC with that from
other microalgae and plants. Amino acid residues which are either well or
perfectly conserved in all sequences are indicated by (.) or (*) above the
alignment, respectively. Primers are underlined with names. C. reinhardtii
sequence (e_gwW.77.4.1) comes from ChlamyCenter (www.chlamy.org). Others
from GeneBank: Brassica napus, AAK60339; Arabidopsis thaliana, CAA70282;
Nicotiana tabacum, AAC41659
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Chapter 9. Isolation and characterization of biotin carboxylase gene and stearoyl ACP
desaturase gene from Chlorella zofingiensis
Figure 9.2 Phylogenic tree of BC sequences from bacteria, algae and plants. A.
arabaticum, Acetohalobium arabaticum (YP_003828313); E. guineensis, Elaeis
guineensis (ABF74732); G. max, Glycine max (AAF80468); M. tuberculosis,
Mycobacterium tuberculosis (NP_216238); R. communis, Ricinus communis
(XP_002519811); S. coelicolor, Streptomyces coelicolor (NP_630967); S.
linguale, Spirosoma linguale (ADB41216); others see Figure 9.1. Numbers
associated with the branches are bootstrap values; Bootstrap was analyzed by
using 1000 replicates.
To characterize the corresponding gene of the BC cDNA, PCR using
genomic DNA as the template was performed. A ca. 4.6 kb fragment was
generated and sequenced. Analysis of the obtained nucleotide sequence revealed
that the product was the corresponding gene of BC cDNA (GenBank accession No.
GQ996718). The coding region of the BC gene is interrupted by eight introns of
346, 288, 377, 439, 339, 298, 358, 422 bp, respectively. Intron/exon splice sites of
the BC gene are highly conservative, and all introns start with GT and end with
AG.
The primers for the isolation and characterization of C. zofingiensis SAD
gene were shown in Table 9.1. The obtained SAD cDNA (GenBank accession No.
GQ996719) contains 1,251 bp ORF that encodes a deduced SAD of 416 amino
acid residues. The SAD protein shared a high homology with that from the near
relative algae C. reinhardtii and H. pluvialis (Figure 9.3). The phylogenetic
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Chapter 9. Isolation and characterization of biotin carboxylase gene and stearoyl ACP
desaturase gene from Chlorella zofingiensis
analysis indicates that the algal SAD proteins are more related to those from
plants (Figure 9.4). The GC content of the SAD coding region is 53.2%, which is
slightly higher than that of SAD from A. thaliana (47.9%) but lower than that from
C. reinhardtii (64.0%) and H. pluvialis (60.4%). The coding region of SAD gene
(GenBank accession No. GQ996720) is interrupted by six introns of 77, 224, 283,
264, 233, and 319 bp, respectively.
Figure 9.3 Amino acid sequence alignment of C. zofingiensis SAD with its
counterparts from C. reinhardtii (estExt_gwp_1W.C_130244), H. pluvialis
(ABP57425), A. thaliana (AAK85232), B. napus (AAT65205) and Helianthus
annuus (AAB65144). Amino acid residues which are either well or perfectly
conserved in all sequences are indicated by (.) or (*) above the alignment,
respectively. Primers are underlined with names. Conserved motifs are marked
with boxes
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desaturase gene from Chlorella zofingiensis
Figure 9.4 Phylogenic tree of BC sequences from bacteria, algae and plants. A.
dehalogenans, Anaeromyxobacter dehalogenans (ZP_02322836); E. guineensis
(AAB41041); G. max (ABM45912); M. tuberculosis (CAE55326); O. sativa,
Oryza sativa (BAA07631); R. communis (XP_002526163); S. coelicolor
(NP_630790); S. linguale (ADB40562); Solanum tuberosum, Solanum tuberosum
(AAA33839); others see Figure 9.3. Numbers associated with the branches are
bootstrap values; Bootstrap was analyzed by using 1000 replicates.
9.4.2 High light irradiation up-regulates the transcripts of BC and
SAD and enhances the biosynthesis of TFA and oleic acid
High light could induce the rapid accumulation of fatty acids in H.
pluvialis (Zhekisheva et al., 2002, 2005; Chen, 2007). In the current study the
effect of high light on the regulation of fatty acid biosynthesis in C. zofingiensis
was assayed. The transcripts of BC and SAD genes of C. zofingiensis under high
light condition were detected by RT-PCR analysis. The non-induced algal cells
had basal BC transcripts (Figure 9.5A, lane 1). Upon exposure of the cells to high
light illumination, a significant increase in the steady-state mRNA level was
observed and the mRNA level reached its maximum at 24 h (Figure 9.5A, lanes
2-5). High light also up-regulated SAD transcripts, yet to much stronger extent as
compared with BC transcripts (Figure 9.5A, lanes 2 and 3). A slight decrease of
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Chapter 9. Isolation and characterization of biotin carboxylase gene and stearoyl ACP
desaturase gene from Chlorella zofingiensis
the steady-state SAD mRNA level was observed after 24 h upon onset of high
light induction (Figure 9.5A, lanes 4 and 5).
To correlate the transcript levels of BC and SAD genes and the
biosynthesis of fatty acids, the accumulation of TFA and oleic acid was examined
over the period of induction (Figure 9.5B). The Chlorella cells without light stress
accumulated only a small amount of TFA (Figure 4B, 0 h). However, upon high
light illumination, a great increase in the TFA content was observed (Figure 5B).
Similarly, high light illumination strikingly stimulated the biosynthesis of oleic
acid (Figure 9.5B). After 72 h of induction, the contents of TFA and oleic acid
reached 19.3% and 3.1% of dry weight, 62.9% and 150.4% higher than that
measured at 0 h, respectively. These results indicate that the up-regulation of BC
leads to the rapid synthesis of TFA while the enhanced SAD expression increases
the accumulation of oleic acid in C. zofingiensis cells.
Figure 9.5 High light induced expression of BC and SAD genes (A) and
accumulation of TFA and oleic acid (B) in C. zofingiensis cells. Dark column,
TFA content; gray column, oleic acid content
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Chapter 9. Isolation and characterization of biotin carboxylase gene and stearoyl ACP
desaturase gene from Chlorella zofingiensis
9.4.3 Salt stress induces the up-regulation of SAD gene and the
accumulation of oleic acid
Salt was reported to be able to promote accumulation of TFA and affected
the fatty acid composition in microalga (Rao et al., 2007). To examine the effect
of salt on expression of BC and SAD genes and fatty acid biosynthesis, 100 mM
NaCl was employed to treat C. zofingiensis cells. During the whole induction
period, the mRNA level of BC stayed almost constant (Figure 9.6A, lanes 1-5).
Correlated with the unchanged BC expression, no significant difference in TFA
content was observed among the algal cell samples treated with salt for 0 to 72 h
(Figure 9.6B). On the contrary, the SAD transcript was transiently up-regulated by
salt and reached a maximum level at 24 h (Figure 9.6A). Accordingly, the oleic
acid content increased upon induction of salt, and after 72 h of induction it
reached 1.7% of dry weight, 39.0% higher than that measured at 0 h, yet much
lower than that induced by high light (Figure 9.6B).
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Chapter 9. Isolation and characterization of biotin carboxylase gene and stearoyl ACP
desaturase gene from Chlorella zofingiensis
Figure 9.6 Effect of salt stress on the expression of BC and SAD genes (A) and
accumulation of TFA and oleic acid (B) in C. zofingiensis cells. Dark column,
TFA content; gray column, oleic acid content
9.4.5 Glucose triggers the up-regulation of BC and SAD genes and
induces the enhanced biosynthesis of TFA and oleic acid
It was reported that glucose could stimulate the biosynthesis of fatty
acids in C. zofingiensis (Chapter 6, 7 and 8). To survey the effect of glucose on
fatty acid biosynthesis at transcriptional level, C. zofingiensis cells were induced
by 50 mM glucose in the absence of light. The mRNA level of BC remained
constant during the first 12 h of glucose induction and was then greatly
up-regulated, reaching its maximum at 36 h (Figure 9.5A, lanes 1-4). Thereafter, a
slight decrease of BC transcripts was observed (Figure 9.5A, lane 5). The mRNA
level of SAD was also transiently up-regulated by glucose and began to decrease
after 24 h of glucose induction (Figure 9.5A).
Consistent with the up-regulation of BC and SAD genes, glucose
enhanced the biosynthesis of TFA and oleic acid (Figure 9.5B). After 72 h of
induction, the TFA content reached 29.3% of dry weight, which was 133.5%
higher than that measured at 0 h and was 52.0% higher than that induced by high
light. In contrast, after 72 h of glucose induction, more than 6-fold amounts of
oleic acid accumulated, which was much higher than that induced by high light.
Thus glucose may be superior to high light for inducing accumulation of TFA,
especially of oleic acid in C. zofingiensis cells.
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Chapter 9. Isolation and characterization of biotin carboxylase gene and stearoyl ACP
desaturase gene from Chlorella zofingiensis
Figure 9.5 Glucose induced expression of BC and SAD genes (A) and
accumulation of TFA and oleic acid (B) in C. zofingiensis cells. Dark column,
TFA content; gray column, oleic acid content
9.5 Discussion
The biosynthesis of fatty acids has been thoroughly investigated at the
molecular level in plants, whereas it is rarely touched in microalgae with a few
exceptions (Roessler & Ohlrogge, 1993; Roessler et al., 1994; Sakamoto et al.,
1994; Chen, 2007). This study reported the isolation and characterization from C.
zofingiensis of two key genes (BC and SAD) involved in fatty acid biosynthesis.
The deduced amino acid sequence of the BC cDNA from C. zofingiensis showed
high identity to its counterparts from other algae and plants (Figure 9.1). BC is a
subunit of ACCase, together with biotin carboxyl carrier protein and
carboxyltransferase to constitute the multisubunit form of ACCase (Bao et al.,
1997). The expression of BC is autoregulated to that of other subunits of ACCase
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Chapter 9. Isolation and characterization of biotin carboxylase gene and stearoyl ACP
desaturase gene from Chlorella zofingiensis
and thus could represent the expression of ACCase (Ke et al., 2000; James &
Cronan, 2004). SAD catalyzes the O2- and NAD(P)H-dependent insertion of a cis
double bond between carbons 9 and 10 of 18:0-ACP to form 18:1-ACP (Sobrado
et al., 2006). Similar to SADs from other algae and plants, the C. zofingiensis
SAD contains two EXXH motifs that are considered conservative in the
diiron-oxo protein class (Figure 9.2).
In response to high light irradiation, BC and SAD genes were
up-regulated in C. zofingiensis (Figure 9.3A); as a result, the treated cells
accumulated a higher amount of TFA including oleic acid (Figure 9.3B). This
might result from the overproduction of intracellular reactive oxygen species
(ROS), because high light can stimulate a drastic increase ROS level in C.
zofingiensis (Li et al., 2009). Further studies are needed to support this hypothesis.
The enhanced de novo fatty acid synthesis consumes a large amount of oxygen,
which in turn might serve as the ROS sequestration process and protect cells from
damage by ROS (Chen, 2007). The increased intracellular ROS level was also
observed in salt treated C. zofingiensis cells (Li et al., 2009). However, Unlike
high light, salt stress only stimulated the up-regulation of SAD (Figure 9.4A);
accordingly, the salt-treated cells accumulated a higher level of oleic acid whereas
no significant difference in TFA content was observed as compared to non treated
cells (Figure 9.4B). It was suggested that high light and salt stress might trigger
the production of different ROS species that differentially stimulated the
expression of BC and SAD genes.
Glucose not only fuel cellular carbon and energy metabolism but also
plays pivotal roles as signaling molecules, for example, to regulate genes involved
in lipid biosynthesis (Rolland et al., 2006). This study showed that glucose could
greatly up-regulate the expression of both BC and SAD genes and enhance the
accumulation of TFA including oleic acid (Figure 9.5). It was suggested that
glucose effects on the expression of genes might be mediated by glucose sensing,
changes in electron flow of respiratory electron transport, or ROS (Moller, 2001;
Moore et al., 2003; Ryu et al., 2004). Although glucose sensing and the
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Chapter 9. Isolation and characterization of biotin carboxylase gene and stearoyl ACP
desaturase gene from Chlorella zofingiensis
mitochondrial alternative pathway were revealed to be involved in the regulation
of carotenogenic genes of C. zofigiensis (Li et al., 2008b), the underlying
mechanism of glucose effect on fatty acid biosynthesis genes remains open.
The well studied pathway of fatty acid biosynthesis and the availability
of genes involved in the fatty acid biosynthesis have greatly facilitated the genetic
modification of plants for oil and fatty acid production, for example, to increase
the total oil content (Roesler et al., 1997; Lardizabal et al., 2008), to change the
composition of fatty acids (Flores et al., 2008; Graef et al., 2009), or to produce
new fatty acids for nutritional improvement (Kinney et al., 2004; Cheng et al.,
2010). Attempts to genetically manipulate microalgae through the induction of
ACCase gene was carried out, with the hope that increasing the level of ACCase
activity in the cells would lead to higher oil production (Sheehan et al., 1998).
These early experiments did not, however, demonstrate increased oil production in
the cells. Better understanding of regulatory processes of fatty acid biosynthesis
and lipid metabolism and better genetic manipulation tools might be needed for
the successful manipulation of microalga for enhanced oil production. The current
study described for the first time the isolation and characterization of two
important genes involved in the fatty acid biosynthesis, namely BC and SAD from
C. zofingiensis. These results might help better understanding the fatty acid
biosynthesis in C. zofingiensis at molecular level and benefit future genetic
modification of this alga to enhance fatty acid production and/or alter fatty acid
composition as a better biodiesel feedstock.
173
PART IV
RESEARCH SUMMARY AND RECOMMENDATION
FOR FUTURE WORK
174
Chapter 10. Research summary and recommendation for future work
Chapter 10
Research summary and recommendation for future work
10.1 Introduction
This thesis addressed the genetic engineering of Chlorella zofingiensis
for promoted astaxanthin production and the assessment of heterotrophic C.
zofingiensis as a biodiesel feedstock. Two approaches, namely, mutagenesis and
manipulation of carotenoid biosynthesis, were employed to enhance the cellular
astaxanthin content of C. zofingiensis. Lipid and fatty acid analyses were
conducted to assess the biodiesel production potential of heterotrophic C.
zofingiensis.
10.2 Research summary
C. zofingiensis represents the fast growing green microalga with the
ability of producing asatxanthin. To further increase the cellular astaxanthin
content, mutagenesis of the alga was performed. Chemical mutagens MNNG (45
μg L-1) and EMS (0.36 M) were employed to treat the algal cells for generating
mutations. Target mutants were isolated by screening the treated cells with 0.5 μM
norflurazon. Of more than 200 mutants, 5 ones that exhibited deeper red intensity
were selected for downstream characterization. No significant differences in cell
growth and pigment profile were observed between the WT and mutants under
normal growth condition without norflurazon. In contrast to WT cells that got
bleached by 0.25 μM norflurazon, the mutants grew well and synthesized
astaxanthin even when the culture medium contained up to 1 μM norflurazon.
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Chapter 10. Research summary and recommendation for future work
Furthermore, these mutants accumulated significant greater amounts of
astaxanthin (up to 54% greater) than the WT when the cultures were induced with
30 g L-1 glucose in the dark for 5 days. Correlated with the higher levels of
astaxanthin, the mutants showed higher transcript levels of BKT and CHYb genes
that are directly involved in astaxanthin biosynthesis.
Norflurazon specifically targets to phytoene desaturase (PDS) and
inhibits its desaturation activity. The norflurazon resistance and enhanced
astaxanthin accumulation of mutants might be attributed to certain mutations in
PDS gene. To confirm this, the C. zofingiensis PDS gene should be first isolated
and characterized. The open reading frame of this PDS gene, interrupted by six
introns, encoded a polypeptide of 558 amino acid residues. The deduced protein
sequence showed high homology with PDSs from other algae, cyanobacteria and
higher plants. Expression of the PDS gene in E. coli demonstrated that the enzyme
was able to convert phytoene to ζ-carotene efficiently. The PDS gene in C.
zofingiensis was shown to be up-regulated by high light and glucose treatment.
E17, one of the stable mutants, produced higher levels of total
carotenoids (TC) and astaxanthin than the WT when induced by high light
irradiation or glucose. A point mutation (C to T) was revealed to occur in the PDS
gene of E17, leading to an amino acid change (L516F) in its coding region. The
mutated PDS exhibited 31-fold resistance to norflurazon when compared to the
WT one as determined by an in vitro assay. Surprisingly, the mutated PDS
exhibited higher efficiency in converting phytoene to ζ-carotene. No difference in
PDS transcripts was found between E17 and WT cells cultured either in normal or
induced conditions. In contrast, higher transcript levels of BKT and CHYb were
found in E17 cells. Therefore, it is concluded that the point mutation in the PDS
gene makes E17 resistant against norflurazon and synthesize higher amounts of
carotenoids including astaxanthin.
Since the PDS gene from E17 encoded a enzyme resistant to norflurazon
and having higher desaturation activity, it may be adopted as a dominant
selectable marker for the transformation of C. zofingiensis. PDS-L516F gene was
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Chapter 10. Research summary and recommendation for future work
introduced into C. zofingiensis via the biolistic approach. Transformants
expressing the mutated PDS gene showed strong resistance to the herbicide
norflurazon. The transformant P6 could accumulate more TC and astaxanthin than
the WT when cultured with glucose in the dark. The enhanced accumulation of
TC and astaxanthin in the transformant was revealed to be related to the increase
of PDS transcript. These results clearly show that the mutated PDS gene is a
useful selectable marker and can be used to genetic engineer carotenoid
biosynthesis for enhanced astaxanthin production in C. zofingiensis.
C. zofingiensis also represents the oil-rich green alga that can grow well
photoautotrophicaly, heterotrophically and mixotrophically. Profiles of algal lipids
and fatty acids, which depend on species and culture conditions, are important
data to evaluate the oils for biodiesel production. So, firstly, the lipid class and
fatty acid composition of C. zofingiensis cultivated under photoautotrophic and
heterotrophic conditions were documented and compared. Compared with
photoautotrophic cells, a 900% increase in lipid yield was achieved in
heterotrophic cells fed with 30 g L-1 of glucose. Furthermore heterotrophic cells
accumulated predominantly neutral lipids (NL) that accounted for 79.5% of total
lipids with 88.7% of NL being triacylglycerol (TAG); whereas photoautotrophic
cells contained mainly the membrane lipids glycolipids (GL) and phospholipids
(PL). Together with the much higher content of oleic acid (C18:1, 35.2% of total
fatty acids), oils from heterotrophic C. zofingiensis appear to be more feasible for
biodiesel production.
Secondly, the lipid production and fatty acid profile of C. zofingiensis
cultured in the dark with various carbon sources were investigated. Of the sugars
surveyed, glucose was found to be the best one for the growth and lipid
production. When cultivated with 50 g L-1 glucose, C. zofingiensis accumulated
lipids up to 52% of the dry biomass, with TAG accounting for 72.1% of the total
lipids. Fatty acid profiles revealed that glucose contributed to the highest yield of
total fatty acids (TFA) and proportion of oleic acid (35.7% of TFA). In fed-batch
cultivation based on glucose, the lipid yield and productivity of C. zofingiensis
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Chapter 10. Research summary and recommendation for future work
were further increased to 20.7 g L-1 and 1.38 g d-1 L-1 respectively, much higher
than those achieved in batch culture. Moreover, the low cost cane molasses was
surveyed and proved to be an ideal carbon source for boosting lipid production by
C. zofingensis. These results suggest that C. zofingiensis has great potential for
biodiesel production.
In addition, the effect of several nutrients, namely, nitrate, phosphate and
ferrous ion as well as the environmental conditions including temperature and the
initial pH of culture medium on the growth and fatty acid profile of heterotrophic
C. zofingiensis in batch culture were surveyed. The optimal nutritional and
environmental conditions for TFA production by C. zofingiensis were 5 mM
nitrate, 5 mM phosphate, 25-100 μM ferrous ion, cultivation temperature of 25 °C
and the initial pH of 6.5.
In order to elucidate the molecular regulation of fatty acid accumulation,
the key genes involved in fatty acid biosynthesis, namely, biotin carboxylase (BC)
and stearoyl ACP desaturase (SAD) genes, were isolated and characterized from C.
zofingiensis. Protein sequence alignments showed that the deduced amino acid
sequences of these two genes shared high identity to that of corresponding genes
from other microalgae and plants. High light illumination and glucose induction
were revealed able to up-regulate the expression of both BC and SAD genes, and
therefore enhanced the accumulation of TFA including oleic acid. Unlike high
light or glucose, salt stress, however, could only trigger the up-regulation of SAD
gene; accordingly, the TFA content was slight affected while the biosynthesis of
oleic acid was promoted. In this context, the fatty acid biosynthesis of C.
zofingiensis might be regulated, at least partly, at the transcriptional level.
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Chapter 10. Research summary and recommendation for future work
10.3 Recommendation for future work
10.3.1 Future work for astaxanthin production by C. zofingiensis
It was reported in chapter 3 that the mutagenesis of C. zofingiensis
coupled with norflurazon selection gave rise to the mutants with the ability of
producing higher amounts of TC including astaxanthin. Following this strategy,
the cells of mutant E17 can be further mutagenized but selected with other
herbicides
that
target
different
carotenogenic
enzymes.
For
example,
diphenylamine, a specific inhibitor to carotenoid ketolase (Zhekisheva et al., 2005;
Wang et al., 2008), can be adopted to select the mutants showing improved
oxygenation activity of converting zeaxanthin to astaxanthin.
Another feasible way to further increase the cellular astaxanthin content
is directly engineering the astaxanthin biosynthetic pathway in C. zofingiensis.
Under induced conditions for carotenoid biosynthesis, apart from astaxanthin, C.
zofingiensis also accumulated substantial amounts of other secondary carotenoids
such as adonixanthin and canthaxanthin (Rise et al., 1994; Bar et al., 1995; Del
Campo et al., 2004; Wang et al., 2008). It might be possible that CHYb cannot
accept canthaxanthin as a substrate to produce astaxanthin and BKT has a
relatively low activity of hydroxylating adonixanthin to astaxanthin in C.
zofingiensis. Therefore, the astaxanthin biosynthetic pathway in C. zofingiensis
was proposed (Li et al., 2008b; Wang et al., 2008) and is shown in Figure 10.1. By
introducing a carotenoid hydroxylase that can convert canthaxanthin to
astaxanthin and a carotenoid ketolase that can catalyze the efficient ketolation of
adonixanthin to astaxanthin, the secondary carotenoid flux in C. zofingiensis
would be changed and directed to the only end product astaxanthin, resulting in
further much enhanced astaxanthin production.
The major drawback of using heterotrophic C. zofingiensis for
astaxanthin production is the need of glucose to feed the cells because of the
relatively high cost of glucose. It was reported in chapter 7 that cane molasses, the
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Chapter 10. Research summary and recommendation for future work
low cost by-product from sugar industry, could be well used as the carbon source
for the growth of heterotrophic C. zofingiensis. Thus, it is reasonable to expect
that cane molasses can be adopted to feed C. zofingiensis for economic production
of astaxanthin.
Phytoene
PDS
β-carotene
BKT
CHYb
β-cryptoxanthin
Echinenone
CHYb
BKT
Zeaxanthin
Canthaxanthin
BKT
Adonixanthin
BKT
Astaxanthin
Figure 10.1 The proposed astaxanthin biosynthetic pathway in C. zofingiens.
Enzymes are named according to the designation of their genes. PDS, phytoene
desaturase; BKT, carotenoid ketolase; CHYb, carotenoid hydroxylase.
10.3.2 Future work for biodiesel production by C. zofingiensis
C. zofingiensis can accumulate oils up to 52% of dry weight in
heterotrophic conditions (chapter 7). The oils are composed almost exclusively of
TAG, but they do contain small amounts of other lipidic compounds such as PL
and GL. TAG consist of three fatty acid chains bound to a glycerol backbone. As
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Chapter 10. Research summary and recommendation for future work
such, theoretically, genetic engineering for cellular oil enhancement can be
achieved through either increasing the synthesis of fatty acids or increasing their
incorporation onto the glycerol backbone. In the former case, this involves
attempts to enhance the partitioning of carbon toward fatty acid synthesis and to
increase the pool sizes of substrates for fatty acid synthesis, through the
overexpression of key metabolic enzymes, such as acetyl-CoA carboxylase
(Roesler et al., 1997). As for the later, diacylglycerol acyltransferase, the enzyme
catalyzes the last step of TAG biosynthesis - the incorporation of a fatty acyl-CoA
onto diacylglycerol, is centred and manipulated (Lardizabal et al., 2008). While
the oil enhancement in a feedstock severs a guide for better yield of biodiesel, the
genetic modification of fatty acid composition points to the better quality of
biodiesel. It has been suggested that the enhanced proportion of oleic acid gives
biodiesel a compromise between oxidative stability and low-temperature
properties and thus promotes biodiesel quality (Knothe, 2009). Oleic acid is
converted to linoleic acid in a single desaturation step carried out by △12 fatty acid
desaturase encoded by the FAD2 gene (Graef et al., 2009). Down-regulation of
FAD2 through posttranscriptional gene-silencing methods results in much higher
levels of oleic acid in soybean seeds (Buhr et al., 2002; Graef et al., 2009), which
may provide a guide for enhanced proportion of oleic acid in C. zofingiensis.
In this thesis, the potential assessment of C. zofingiensis as a biodiesel
feedstock focused on the lipid and fatty acid analyses. Further explorations can be
carried out, in terms of transesterification of C. zofingiensis derived oils for
producing biodiesel and analysis of the biodiesel properties including density,
viscosity, flash point, cloud point, cold-filter plugging point, heating value,
oxidative stability, etc.
Preliminary results suggested that the accumulation of lipids/fatty acids
was accompanied by the accumulation of astaxanthin in heterotrophic C.
zofingiensis. The underlying mechanism remains largely unknown and needs to be
further investigated. This also stimulates the idea of simultaneous production of
oils and astaxanthin by C. zofingiensis in the heterotrophic conditions, which,
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Chapter 10. Research summary and recommendation for future work
once implemented, will cut down the production cost and make this alga more
commercially economical.
182
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