LIPID PRODUCTION, PHOTOSYNTHESIS ADJUSTMENT AND CARBON
PARTITIONING OF MICROALGA CHLORELLA SOROKINIANA
CULTURED UNDER MIXOTROPHIC AND
HETEROTROPHIC CONDITIONS
By
TINGTING LI
A dissertation submitted in partial fulfillment of
the requirements for the degree of
DOCTOR OF PHILOSOPHY
WASHINGTON STATE UNIVERSITY
College of Engineering and Architecture
AUGUST 2014
To the Faculty of Washington State University:
The members of the Committee appointed to examine the dissertation of TINGTING LI
find it satisfactory and recommend that it be accepted.
___________________________________
Shulin Chen, Ph.D., Chair
___________________________________
Helmut Kirchhoff, Ph.D.
___________________________________
Philip T. Pienkos, Ph.D.
___________________________________
Yinjie Tang, Ph.D.
___________________________________
Manuel Garcia-Perez, Ph.D.
ii
ACKNOWLEDGEMENTS
I am very grateful to my advisor Dr. Shulin Chen who gave me valuable opportunity to study in
US, and provided guidance, support and encouragement during my Ph.D. program. Besides my
advisor, I would like to thank the rest of my committee members: Dr. Helmut Kirchhoff, Dr.
Philip T. Pienkos, Dr. Yinjie Tang, and Dr. Manuel Garcia-Perez, for their encouragement,
insightful comments, and hard questions. Help provided by Dr. Mahmoud Gargouri and Dr.
Jeong-Jin Park in analysis of RT-PCR and metabolites is greatly appreciated.
I want to thank my fellow labmates in BBEL: Yubin Zheng, Jijiao Zeng, Tao Dong, Pierre
Wensel, Liang Yu, Chao Miao, Difeng Gao, Xi Wang, Xiaochao Xiong, Jingwei Ma, Xiaochen
Yu, Jieni Lian, Yuxiao Xie, Xin Gao and Samantha, for their kind help and suggestion on my
research, and for all the funs we have created in Pullman during the past five years.
I would also like to thank my parents for their emotional support and understanding for not
being able to take care of them, and not being able to show up in family gatherings. My special
gratitude goes to my husband, for his support, encouragement, and magnanimity.
iii
LIPID PRODUCTION, PHOTOSYNTHESIS ADJUSTMENT AND CARBON
PARTITIONING OF MICROALGA CHLORELLA SOROKINIANA
CULTURED UNDER MIXOTROPHIC AND
HETEROTROPHIC CONDITIONS
ABSTRACT
by Tingting Li, Ph.D.
Washington State University
August 2014
Chair: Shulin Chen
Microalgae have attracted growing attention for renewable biofuel production. In this work,
microalga Chlorella sorokiniana was explored for its potential for lipid production. The results
showed that C. sorokiniana could tolerate high temperature (up to 42 oC) and had high
productivity. Under batch fermentation condition, 37.6 g L‒1 of biomass could be achieved with
80 g L‒1 of initial glucose supplementation.
Rapid growth rate and high lipid content were obtained through mixotrophic cultivation under
the experimental conditions. The specific growth rate (3.40 d─1) and the biomass yield (0.82 g
g─1) were 1.8- and 2.4-folds of those in heterotrophic culture. Most significantly, lipid content in
mixotrophic cells reached 45%, whereas it was only 13% in the control heterotrophically
cultured cells. Linear electron flow and CO2 refixation were demonstrated to be critical factors
for boosting the cell growth and lipid accumulation in mixotrophic culture.
iv
To gain further insights into the mixotrophic growth, regulation of photosynthesis was
examined. Photosynthetic activities and fluorescence parameters were measured and
characterized, followed by analysis of 77K fluorescence spectra, enzyme activities and enzyme
expression levels. In the exponential phase, the mixotrophic cells exhibited enhanced
photosynthetic activity; in the stationary phase, down-regulation of photosynthesis was observed,
which was proved to be closely correlated with nitrogen limitation.
Starch is another major carbon and energy storage compound in many microalgal cells. This
study revealed the pattern of starch and lipid accumulation in C. sorokiniana controlled by
nitrogen levels: when nitrogen was replete, starch was the predominant form of carbon storage;
after nitrogen was depleted, large amounts of lipid accumulation occurred accompanied by starch
degradation. Finally, lipid became the only carbon sink in the biomass partially contributed from
starch degradation and the turnover of some primary metabolites.
These results provide insight for using microalga C. sorokiniana for biofuel production.
v
Dedication
This dissertation is dedicated to my family.
vi
TABLE OF CONTENTS
Page
ACKNOWLEDGEMENTS ........................................................................................................... iii
ABSTRACT ................................................................................................................................... iv
LIST OF TABLES ........................................................................................................................ xii
LIST OF FIGURES ..................................................................................................................... xiv
CHAPTER ONE ..............................................................................................................................1
1.1 MICROALGAE BIOFUELS .................................................................................................1
1.2 MICROALGAE CULTIVATION .........................................................................................3
1.2.1 Photoautotrophic culture..................................................................................................3
1.2.2 Heterotrophic culture .......................................................................................................4
1.2.3 Mixotrophic culture .........................................................................................................4
1.3 GENERAL OVERVIEW OF PHOTOSYNTHESIS .............................................................5
1.4 LIPID AND STARCH METABOLISM ................................................................................7
1.5 AIM OF THIS WORK .........................................................................................................11
CHAPTER TWO ...........................................................................................................................13
2.1 ABSTRACT .........................................................................................................................13
2.2 INTRODUCTION ................................................................................................................14
2.3 MATERIALS AND METHODS .........................................................................................17
2.3.1 Organism and medium...................................................................................................17
2.3.2 Shake-flask cultures .......................................................................................................17
2.3.3 Batch culture ..................................................................................................................18
2.3.4 Analytical procedure......................................................................................................18
vii
2.3.5 Model analysis ...............................................................................................................19
2.4 RESULTS AND DISCUSSION ..........................................................................................21
2.4.1 Effects of temperature ...................................................................................................21
2.4.2 Effects of carbon sources ...............................................................................................25
2.4.3 Effects of nitrogen sources ............................................................................................27
2.4.4 Effects of initial carbon and nitrogen concentrations ....................................................28
2.4.5 Validating the optimal conditions using BP ANN model and genetic algorithm .........30
2.4.6 Batch culture in 5-L bioreactor ......................................................................................32
2.5 CONCLUSION ....................................................................................................................33
CHAPTER THREE .......................................................................................................................36
3.1 ABSTRACT .........................................................................................................................36
3.2 INTRODUCTION ................................................................................................................38
3.3 MATERIALS AND METHODS .........................................................................................41
3.3.1 Organism and culture conditions ...................................................................................41
3.3.2 Growth analysis .............................................................................................................42
3.3.3 Chemical analysis ..........................................................................................................42
3.3.4 Calculation of iodine value ............................................................................................43
3.4 RESULTS.............................................................................................................................44
3.4.1 Growth and lipid accumulation of microalgae ..............................................................44
3.4.2 Effects of light intensity ................................................................................................49
3.4.3 Effects of DCMU...........................................................................................................52
3.4.4 Effects of CO2 ................................................................................................................54
3.4.5 Effects of temperature ...................................................................................................55
viii
3.5 DISCUSSION ......................................................................................................................56
3.6 CONCLUSION ....................................................................................................................63
CHAPTER FOUR ..........................................................................................................................64
4.1 ABSTRACT .........................................................................................................................64
4.2 INTRODUCTION ................................................................................................................65
4.3 MATERIALS AND METHODS .........................................................................................68
4.3.1 Organism and medium...................................................................................................68
4.3.2 Growth analysis .............................................................................................................68
4.3.3 Photosynthetic oxygen evolution...................................................................................68
4.3.4 Photosynthetic activity measurements...........................................................................69
4.3.5 77K fluorescence spectroscopy .....................................................................................70
4.3.6 Measurements of rubisco activity ..................................................................................70
4.3.7 Quantitative RT-PCR conditions and analysis ..............................................................71
4.4 RESULTS.............................................................................................................................72
4.4.1 Photoautotrophic, mixotrophic and heterotrophic cultivation of C. sorokiniana ..........72
4.4.2 Comparison of photosynthesis and respiration for algal cells cultured mixotrophically
and photoautotrophically ........................................................................................................72
4.4.3 Fluorescence characterization of mixotrophic and photoautotrophic cells ...................75
4.4.4 ΦPSII vs. qE and ΦPSII vs. qL ..........................................................................................78
4.4.5 77K fluorescence spectra ...............................................................................................80
4.4.6 Adjustment of Rubisco and PRK...................................................................................82
4.5 DISCUSSION ......................................................................................................................84
4.6 CONCLUSION ....................................................................................................................88
ix
CHAPTER FIVE ...........................................................................................................................90
5.1 ABSTRACT .........................................................................................................................90
5.2 INTRODUCTION ................................................................................................................91
5.3 MATERIALS AND METHODS .........................................................................................94
5.3.1 Organism and culture conditions ...................................................................................94
5.3.2 Analytical procedure......................................................................................................94
5.3.3 Quantitative RT-PCR conditions and analysis ..............................................................95
5.3.4 Metabolite analysis ........................................................................................................96
5.4 RESULTS.............................................................................................................................98
5.4.1 Sequential accumulation of starch and lipids ................................................................98
5.4.2 Accumulation of starch and lipids depends on nitrogen status ...................................100
5.4.3 The biosynthesis of starch and lipids are both benefitted from active photosynthetic
activity ..................................................................................................................................102
5.4.4 Lipid biosynthesis does not require high light intensity ..............................................103
5.4.5 Declining starch levels are mainly attributed to starch catabolism .............................104
5.4.6 Lipid synthesis is largely dependent on starch degradation ........................................105
5.4.7 Metabolites profiling in C. sorokiniana ......................................................................107
5.5 DISCUSSION ....................................................................................................................111
5.5.1 Accumulation patterns of starch and lipids .................................................................111
5.5.2 Influential factors for lipid synthesis ...........................................................................112
5.5.3 The relationship between starch and lipid contents .....................................................113
5.5.4 Comparison of starch and lipid accumulation from an energy perspective ................114
5.6 CONCLUSION ..................................................................................................................117
x
CHAPTER SIX ............................................................................................................................118
BIBLIOGRAPHY ........................................................................................................................122
xi
LIST OF TABLES
1. Table 1.1 Energetic (ATP and NAD(P)H) requirements for the synthesis of TAG and storage
carbohydrate per unit carbon .....................................................................................................10
2. Table 2.1 Cell growth parameters of C. sorokiniana at various temperatures ..........................23
3. Table 2.2 Fatty acid profiles of heterotrophically cultured algae C. sorokiniana at different
temperatures ..............................................................................................................................24
4. Table 2.3 Effects of initial glucose and KNO3 concentrations in heterotrophic culture of C.
sorokiniana ................................................................................................................................30
5. Table 2.4 Comparison of results of batch-mode culture using glucose as substrate in this work
and previous reports ..................................................................................................................35
6. Table 3.1 Growth performance of C. sorokiniana at 25 oC .......................................................46
7. Table 3.2 Fatty acid profiles of photoautotrophically, heterotrophically and mixotrophically
cultured C. sorokiniana .............................................................................................................47
8. Table 3.3 Growth performance of C. sorokiniana under three different culture modes at 25 oC
and 37 oC ...................................................................................................................................55
9. Table 3.4 Comparison of current research results with previous reports of microalgal cultures
under three different culture modes...........................................................................................60
10. Table 4.1 Primers for quantitative real-time PCR detection of expression genes in C.
sorokiniana ................................................................................................................................71
11. Table 4.2 Growth parameters of C. sorokiniana under different culture conditions ...............74
12. Table 4.3 Characteristic P-I curve parameters (Rd, Pmax, Ik and α-slope) of C. sorokiniana
cultured under mixotrophic and photoautotrophic conditions .................................................75
xii
13. Table 4.4 Effects of different growth conditions on total Rubisco carboxylase activity and
chlorophyll to protein ratio ......................................................................................................82
14. Table 4.5 Chlorophyll fluorescence characteristics of algal cells............................................89
15. Table 5.1 Primers for quantitative real-time PCR detection of expression genes in C.
sorokiniana ..............................................................................................................................96
xiii
LIST OF FIGURES
1. Figure 1.1 Simplified overview of the representative pathways in microalgal lipid and starch
biosynthesis in mixotrophic growth ............................................................................................9
2. Figure 2.1 Cell growth curves of C. sorokiniana for heterotrophic cultivation at various
temperatures ..............................................................................................................................22
3. Figure 2.2 DCW of C. sorokiniana under different carbon sources and nitrogen sources ........26
4. Figure 2.3 Cell growth curves of C. sorokiniana for heterotrophic cultivation at different
initial glucose concentrations ....................................................................................................29
5. Figure 2.4 Comparison of the BP neural network prediction and experimental data. ...............31
6. Figure 2.5 Cell growth and substrate consumption of heterotrophic C. sorokiniana in batch
culture ........................................................................................................................................32
7. Figure 3.1 Effects of glucose concentrations on the growth of C. sorokiniana in the light
condition and in the dark condition ...........................................................................................45
8. Figure 3.2 Effects of light intensities on the growth of C. sorokiniana ....................................50
9. Figure 3.3 Comparison of specific growth rate and lipid content of mixotrophically cultured C.
sorokiniana under various light intensities ...............................................................................51
10. Figure 3.4 Effects of DCMU supplementation on growth and lipid content of C. sorokiniana
under mixotrophic and heterotrophic conditions.......................................................................53
11. Figure 3.5 Influence of CO2 content on DW and lipid content of C. sorokiniana under
photoautotrophic and mixotrophic culture conditions, respectively .........................................54
12. Figure 4.1 Cultivation of C. sorokiniana under different conditions ......................................73
13. Figure 4.2 Light-saturation curves of photosynthesis in mixotrophically and
photoautotrophically cultured C. sorokiniana ...........................................................................76
xiv
14. Figure 4.3 Photosynthetic characteristics of C. sorokiniana grown mixotrophically and
photoautotrophically ................................................................................................................78
15. Figure 4.4 ΦPSII vs. qE and ΦPSII vs. qL of C. sorokiniana grown mixotrophically and
photoautotrophically ................................................................................................................79
16. Figure 4.5 77K fluorescence emission spectra of mixotrophic and photoautotrophic cells after
excitation of chlorophylls at 435 nm .......................................................................................81
17. Figure 4.6 RT-PCR results of Rubisco and phosphoribulokinase ...........................................83
18. Figure 5.1 Mixotrophic and heterotrophic cultivation of C. sorokiniana. ...............................99
19. Figure 5.2 Starch and lipid accumulation in C. sorokiniana in response to mixotrophic
growth is dependent on photosynthetic electron transport ....................................................101
20. Figure 5.3 Time course of lipid and starch accumulation in C. sorokiniana in response to N
starvation ................................................................................................................................103
21. Figure 5.4 RT-PCR results of critical enzymes involved in the synthesis of starch and lipid
and in the degradation of starch .............................................................................................105
22. Figure 5.5 Effect of cycloheximide on changes of starch and lipid content in mixotrophic
growth of C. sorokiniana .......................................................................................................107
23. Figure 5.6 Modeling of metabolite shifts during mixotrophic growth ..................................109
24. Figure 5.7 Primary metabolite profiles in C. sorokiniana during the mixotrophic growth with
time course .............................................................................................................................110
xv
CHAPTER ONE
INTRODUCTION
1.1 MICROALGAE BIOFUELS
More than 80% of global energy demand today is met from fossil fuel (Brennan & Owende,
2010). It has become obvious that continued reliance on fossil fuel energy resources is
unsustainable, owing to higher energy prices, finite fossil fuel reserves, and rising atmospheric
CO2 levels. As a result, researchers and policy-makers are exploring alternative energy
feedstocks in the hope of averting some of the most unfortunate scenarios. This includes the
production of biofuel, which is biodegradable, renewable, and non-toxic. Biofuel also contributes
no net carbon dioxide or sulfur to the air and emits less atmospheric pollutants than fossil fuel
(Sharif Hossain et al., 2008).
In recent years, the demand for liquid biofuels in the transport sector has shown rapid global
growth. First generation biofuels are mainly produced from food and oil crops including
rapeseed oil, sugarcane, sugar beet, and maize as well as vegetable oils and animal fats (Fukuda
et al., 2001; Kulkarni & Dalai, 2006). However, those sources can lead to competition with
edible food, and cannot realistically satisfy the potential market for biofuel even not designated
for other utilization (Chisti, 2008). The Energy Independence and Security Act (EISA) mandates
the minimum annual levels of renewable transportation fuel in the United States to be 36 billion
in 2020. Starting in 2016, all of the increase in the renewable fuel standard target must be met
with advanced biofuels, defined as cellulosic ethanol and other biofuels derived from feedstock
other than corn starch (Sissine, 2007). Thus, technically and economically viable biofuel
resources are urgently needed.
Microalgae have shown great potential to produce a wide spectrum of fuel products in pilot
studies: (1) hydrogen (H2) via direct and indirect biophotolysis, (2) biodiesel through
1
transesterification, (3) biomethane via anaerobic digestion, (4) bioethanol by genetic engineering,
(5) bio-oil via thermochemical conversion, and (6) green diesel and gasoline through direct
catalytic hydrothermal liquefaction (Damartzis & Zabaniotou, 2011; Fatih Demirbas, 2009;
Nigam & Singh, 2011). “Drop-in” liquid fuels converted from microalgal lipids are compatible
with the existing liquid transportation infrastructure.
Many advantages exist for using microalgae-derived biofuels: (1) microalgae cells have much
higher solar energy conversion efficiencies in photosynthesis (Melis, 2009). The maximum
conversion efficiency of solar energy to biomass is 8-10% for microalgae, but only 4.6% for C3
plants and 6% for C4 plants; (2) microalgae are capable of all year round production in some
areas, and some microalgae are capable of accumulating large amounts of lipids (up to 70% w/w)
(Scott et al., 2010), therefore, oil productivity of microalgae cultures exceeds the yield of the best
oilseed crops, e.g. biodiesel yield of 12,000 L ha‒1 for microalgae (open pond production)
compared with 5950 L ha‒1 for oil palm (Schenk et al., 2008); (3) the cultivation of microalgae
for biofuel production can potentially be carried out on non-arable land, and thus do not directly
compete with food production (Chisti, 2008); (4) considering minimal evaporation of closed
photobioreactor systems and the capability of some marine and halophilic algal strains,
microalgal cultivation can save large amounts of fresh water compared with traditional biofuel
crops; in addition, some microalgae can recycle water and nutrients from effluent streams,
therefore, it is beneficial for treatment of waste water (Cantrell et al., 2008); they can also use
flue gases from power plants, offering additional environmental benefits; (5) a production failure
in an algal pond can generally be brought back on line in a matter of days whereas a crop failure
in a terrestrial crop production system may take up to a year before another harvest can occur; (6)
Some microalgae can also synthesize desirable compounds, like β-carotenoids, docosahexaenoic
2
acid (DHA), and astaxanthin, with commercial or pharmaceutical applications (Spolaore et al.,
2006).
However, production cost associated with microalgae biofuel is the major barrier in its
commercialization.
1.2 MICROALGAE CULTIVATION
In general, microalgae growth occurs in three modes, including photoautotrophy, heterotrophy,
and/or mixotrophy. As photosynthetic microorganisms, microalgae depend on sunlight and CO2
as the energy and carbon source for growth. Under heterotrophic growth conditions, instead of
harvesting light and assimilating CO2 from air, microalgae use organic carbon substrates such as
glucose, acetate, and glycerol as their energy and carbon source. Mixotrophic growth is a
combination of phototrophic and heterotrophic conditions, where some microalgae strains have
the capability to combine autotrophic photosynthesis and the heterotrophic assimilation of
organic compounds, either simultaneously or sequentially.
1.2.1 Photoautotrophic culture
Currently, the most common procedure for cultivation microalgae is photoautotrophic culture,
which can be carried out in either open systems, such as ponds, or in closed systems such as
photobioreactors (PBRs). The open culture option has been investigated by many researchers due
to its relative low cost. However, open ponds are subject to daily and seasonal changes in
temperature and humidity, susceptibility of contamination by invading species and insects
(Rusch & Malone, 1998; Theegala et al., 1999), and relatively low biomass productivity due to
the poor carbon dioxide utilization efficiency (Chisti, 2007). Compared with open ponds, PBRs
are designed to overcome the limitations of open pond systems. They provide with much higher
biomass productivity through better control on the cultural systems, such as high surface-to-
3
volume ratio, reduced contamination risks, reduced loss of water. However, PBRs have suffered
from technical problems of scalability and the high capital costs. Generally, the photoautotrophic
culture mode has a low biomass concentration accompanied by high biomass harvesting cost. To
date, photoautotrophic cultivation of microalgae, such as Arthrospira, Chlorella, Dunaliella,
Aphanizomenon,
Haematococcus,
Crypthecodinium,
and
Schizochytrium
are
only
commercialized for high value add products (Spolaore et al., 2006).
1.2.2 Heterotrophic culture
A feasible alternative to photoautotrophic culture is to use heterotrophic culture, which
eliminates the requirement for light, and can be performed in conventional microbial bioreactors.
It is much easier to alter conditions to improve the yield of biomass and reduce the cost of
microalgal biomass production. Moreover, heterotrophic culture of microalgal can provide high
biomass density and induce the accumulation of high amount of lipid (Miao & Wu, 2006; Xiong
et al., 2008). It has been successfully applied to commercial production of high-value molecules
such as docosahexaenoic acid (DHA) (Spolaore et al., 2006). The primary challenges with this
strategy are the cost and availability of suitable feedstocks such as lignocellulosic sugars.
Because these systems rely on primary productivity from other sources, they could compete for
feedstocks with food and other biofuel technologies. Besides, under heterotrophic conditions,
some photosynthetically derived products are not accumulated.
1.2.3 Mixotrophic culture
Mixotrophic growth offers several advantages. It’s well documented that algae have higher
growth rate and produce more cell biomass under mixotrophic conditions (Endo et al., 1977;
Jeon et al., 2006; Mandal & Mallick, 2009). Mixotrophically cultured algae can simultaneously
drive phototrophy and heterotrophy to utilize both inorganic (CO2) and organic carbon substrates,
4
thus leading to an additive or synergistic effect of the two processes that enhances the
productivity. Another interesting point is photoinhibitory effects is reduced or even stopped
while algae grow mixotrophically, which is explained either by the protective influence of the
nutrient or the shift in photoinhibitory-light intensity to a higher level (Chojnacka & Noworyta,
2004). Photooxidative damages caused by oxygen accumulation, especially in closed
photobioreactors, may also be alleviated because of the oxygen consumption in the heterotrophic
metabolic activities (Chojnacka & Marquez-Rocha, 2004). Compared with phototrophic and
heterotrophic cultivation, however, there is less information in the literature concerning
mixotrophic cultivation for microalgal oil production (Chen et al., 2011).
1.3 GENERAL OVERVIEW OF PHOTOSYNTHESIS
Photosynthesis is essentially the source of all the carbon compounds. It is the first step in the
conversion of light to chemical energy and ultimately responsible for driving the production of
the feedstocks. In plants and eukaryotic algae photosynthesis occurs in the chloroplast and can be
divided into the light reactions, in which light energy is stored as ATP and NADPH, and the dark
reactions, in which the products of the light reactions are used to reduce CO2.
In the light reactions, light is absorbed by light harvesting complex (LHC) proteins. Besides
their role as light energy capture proteins, LHC also plays a role in the dissipation of excess
energy as heat or fluorescence via a mechanism called ‘energy-dependent non-photochemical
quenching’ (NPQ) of chlorophyll fluorescence (Kramer et al., 2004), which would otherwise
damage the photosynthetic reaction centers, in particular photosystem II (PSII) (Adir et al., 2003).
Excitation energy then funneled to the photosynthetic reaction centers of photosystem I (PSI)
and PSII to drive the photosynthetic reactions. The majority of photosynthetic energy is stored in
a process termed linear electron flow (LEF) (Heilmann et al.). LEF involves light-stimulated
5
electron transfer in two separate reaction centers, from water through PS II and PS I and finally
to NADPH, coupled with H+ shuttle across the thylakoid membrane, together with establishing
proton motive force (pmf). The pmf can be used to drive ATP synthesis by the transport of H+
through the ATP synthase back into the stroma. The pmf also acts as a major regulator of
photosynthesis, triggering photoprotective qE ‘quenching’ of excitation energy (Baker et al.,
2007). The light reaction of photosynthesis must be finely regulated, and the output of ATP and
NADPH must be precisely balanced to match the needs of downstream metabolism requirement,
such as fatty acid synthesis, CO2 fixation, nitrogen assimilation, and other basic cell structural
maintenance.
The dark reaction, known as Calvin cycle, is an integral part of the photosynthetic process and
is the starting point of carbon metabolism in photosynthetic organisms. The Calvin cycle can be
divided into three main steps that involve carboxylation, reduction and substrate (ribulose-1,5bisphosphate (RuBP)) regeneration. In the first step, CO2 is fixed, and two 3-phosphoglycerate
molecules
from
RuBP,
CO2
and
H2O
are
catalyzed
by ribulose-1,5-bisphosphate
carboxylase/oxygenase (Rubisco). Rubisco is the most abundant protein on Earth, constituting
some 30% of total proteins in most leaves (Parry et al., 2003), and all carbon found in living
organisms on Earth was once fixed by this enzyme from atmospheric CO2. Apart from
carboxylase activity for carbon fixation, it also functions as an oxygenase to catalyzes the
condensation of O2 with ribulose-1,5-bisphosphate as part of photorespiration. O2 and CO2
compete for the same catalytic site, so that the efficiency of CO2 fixation can be impaired in
certain aerobic environments. In the second step, an ATP/NADPH-dependent reduction phase,
these carboxylic acids are reduced to form two molecules of glyceraldehyde-3-phosphate (G3P).
G3Ps are then subject to further use, e. g. synthesis of starch and fatty acids after conversion to
6
dihydroxyacetone phosphate. In a third step, consisting of a series of reactions a proportion of
G3P is converted back to RuBP required to allow the photosynthetic reduction cycle to continue
(Raines, 2003). The last reaction of the third step is irreversibly catalyzed by
phosphoribulokinase (PRK), a nuclear-encoded plastid-localized enzyme. There is no known
alternative pathway to synthesize RuBP in photosynthetic organisms except the Archaea, which
utilize a special Form III Rubisco (Rumpho et al., 2009). Hence, all other photosynthetic
organisms are dependent on PRK to sustain the cyclic activity.
1.4 LIPID AND STARCH METABOLISM
Most of microalgae are able to accumulate energy-rich reserve compounds suitable for
renewable energy applications, such as lipids and starch, which can range from 6 to 64% of the
total biomass (Subramanian et al., 2013). In eukaryotic microalgae and plants, the de novo fatty
acid biosynthesis begins in the chloroplast. The first step in fatty acid synthesis is the conversion
of acetyl-CoA to malonyl CoA, catalyzed by acetyl-CoA carboxylase (ACCase). Subsequently,
the fatty acids synthesized in the chloroplast are transferred to the endoplasmic reticulum (ER)
membranes for synthesis of membrane polar lipids and cytosolic storage neutral lipid (Hu et al.,
2008). Neutral lipids are mainly in the form of triacylglycerol (TAG), and can be converted to
fatty acid methyl esters (FAMEs), the main components of biodiesel, through transesterification,
or refined into other fuel constituents.
Synthesis and accumulation of large amounts of TAGs can occur in microalgae when placed
under stress conditions imposed by chemical or physical environmental stimuli, either acting
individually or in combination (Hu et al., 2008), such as the use of nutrients stress, including
nitrogen and/or phosphorus starvation, light irradiation, pH, temperature, heavy metals and other
chemicals. Nutrient starvation is one of the mostly widely used lipid induction techniques in
7
microalgal TAG production and has been reported for use in many species. Almost all the
microalgae species studied so far, could increase lipid production under nitrogen stress, although
the occurrence and the extent to which TAGs are produced is species/strain-specific, and are
ultimately controlled by the genetic make-up of individual organisms. However, high lipid
production due to nitrogen stress is at the cost of slow growth rates and low biomass density and
thus finally affecting the total biomass and lipid productivity (Sharma et al., 2012).
Starch is another major carbon and energy storage compound occurring in many plant and
microalgal cells, particularly under stress conditions. Starch synthesis and degradation pathways
have been better understood in algae and plants. The first committed step of starch synthesis in
the plastid is catalyzed by ADP-glucose pyrophosphorylase (AGPase), which converts glucose 1phosphate and ATP to ADP-glucose and PPi. Subsequently, ADP-glucose is used by starch
synthases and branching enzymes to elongate the glucan chains of the starch granule (Zeeman et
al., 2010c). Therefore, AGPase is demonstrated to be a key factor for control of starch synthesis.
Besides starch synthesis, starch degradation has direct effect on the level of starch in the cell.
There is good evidence that starch degradation is dependent on the reversible phosphorylation of
glucans at the surface of the starch granule, which serves to solubilize the granule surface, thus
allowing hydrolases access to the glucan chains. Currently it is considered that the hydrolysis of
the starch is catalyzed mainly by β-amylases and debranching enzymes (isoamylase). These
two types of enzymes hydrolyse the 1,4- and 1,6-glycosidic bonds, respectively (Zeeman et
al., 2010b).
However, regulation of starch and lipid synthesis and the possible interaction between the two
pathways is still not completely known. From viewpoint of carbon source, synthesis of starch
and lipids shares the same carbon precursor (triose-phosphate, Fig 1.1) and is expected to be two
8
Figure 1.1 Simplified overview of the representative pathways in microalgal lipid and starch
biosynthesis in mixotrophic growth.
competitive pathways. From aspect of energy, ATP and NADPH requirement of these two
processes are different (Table 1.1), considering TAG molecules containing fatty acids with the
following chain lengths and degree of saturation, C16:0, C18:1, and C18:3, and having the
9
Table 1.1 Energetic (ATP and NAD(P)H) requirements for the synthesis of TAG and storage
carbohydrate per unit carbon.
ATP/NAD(P)H
Synthesis reactions related with lipid and starch:
3 CO2 →3-phosphoglyceraldehyde →acetylCoA
AcetylCoA mediated fatty acid elongation (e.g., C16:0)
Fatty acid desaturation
3CO2→Glycerol
6CO2 →2 3-phosphoglyceraldehyde →glucose
Glucose →storage carbohydrate
Fatty acids (C16:0, C18:1, C18:3) synthesis summary:
26 acetyl CoA =
Fatty acid elongation C16:0 + 2 C18:0 =
4 desaturations
Total
Per carbon
TAG (C55H98O6) synthesis summary:
Total from fatty acids synthesis
1 glycerol
Total
Per carbon
Starch synthesis (55 units, 330 C) summary:
Total for 55 glucose
Per carbon
7 ATP, 4 NAD(P)H (CO2
release)
7 ATP, 14 NADPH
1 NADH equivalent/bond
desaturated
9 ATP, 7 NAD(P)H
18 ATP, 12 NAD(P)H
1 ATP/glucose
182 ATP, 104 NAD(P)H
23 ATP, 46 NAD(P)H
4 NAD(P)H equivalents
205 ATP, 154 NAD(P)H
3.94 ATP, 2.96 NAD(P)H
205 ATP, 154 NAD(P)H
9 ATP, 7 NAD(P)H
214 ATP, 161 NAD(P)H
3.89 ATP, 2.93 NAD(P)H
1045 ATP, 660 NAD(P)H
3.17 ATP, 2 NAD(P)H
molecular formula C55H98O6. As summarized in Table 1, the energy required per carbon for TAG
synthesis is 3.89 ATP and 2.93 NADPH, and for starch synthesis is 3.17 ATP and 2 NADPH.
For starch synthesis, the energy requirement is dominated by carbon reduction in the Calvin
cycle. Activation of glucose units for starch polymerization requires only one ATP for the
synthesis of ADP-glucose, and it has been reported that exogenous glucose could be directly
converted into starch without prior conversion to triose-phosphates (Fettke et al., 2010).
10
1.5 AIM OF THIS WORK
Though microalgae have certain advantages when compared to higher plants, their actual
potential is still far from being realized, and the microalgal oil yield is also proved to be straindependent. Economic analyses indicate that growth rate and lipid content are two predominant
cost drivers for economical algal biofuel production.
In this work, the possibility of microalga C. sorokiniana as candidate for biofuel production
with high productivities of biomass and lipid was firstly evaluated, which could also provide
support for the microalgae C. sorokiniana being used as a candidate strain for simultaneous
saccharification and fermentation techniques with cellulosic materials as feedstock. Secondly,
mixotrophic growth of C. sorokiniana was explored as the most efficient cultivation strategy for
biomass and lipid production, considering mixotrophy as a way of keeping the features of both
heterotrophy and photoautotrophy. It is expected that CO2 release from glucose metabolism
would be refixed in the Calvin Cycle (Fig 1.1), which would lead to higher growth rate and
higher biomass yield based on glucose. Since light was reported as important factor for lipid
accumulation within the cells, higher lipid content in mixotrophic cells of C. sorokiniana was
also expected. Thirdly, photosynthesis regulation in mixotrophic cells of C. sorokiniana was
examined, based on the possibility that glucose metabolism may exert influences on utilization of
inorganic carbon and thus further alter electron flow of photosynthesis in algal cells. This study
will help us gain deeper understanding about mixotrophic cultivation of microalgae, and would
provide a clue on how to keep high photosynthesis efficiency for carbon acquisition and finally
for biomass accumulation. Fourthly, how starch and lipids accumulation is regulated in algal
cells of C. sorokiniana was investigated. Understanding carbon partitioning in algal cells into
lipid and/or starch is critical for biofuels strain development and cultivation strategy design.
11
The following four manuscripts presented at four separate chapters in the dissertation are:
[1] Tingting Li, Yubin Zheng, Liang Yu, Shulin Chen. (2013) High productivity cultivation of a
heat-resistant microalgae Chlorella sorokiniana for biofuel production. Bioresource Technology,
131:60-67.
[2] Tingting Li, Yubin Zheng, Liang Yu, Shulin Chen. (2014) Mixotrophic cultivation of a
Chlorella sorokiniana strain for enhanced biomass and lipid production. Biomass and Bioenergy,
66:204-213.
[3] Tingting Li, Helmut Kirchhoff, Mahmoud Gargouri, Jie Feng, Asaph Cousins, Jeong-Jin Park,
Philip T. Pienkos, David R. Gang, Shulin Chen. Regulation of photosynthesis in mixotrophic
cells of a microalga strain Chlorella sorokiniana (ready to be submitted).
[4] Tingting Li, Mahmoud Gargouri, Jie Feng, Jeong-Jin Park, Difeng Gao, Chao Miao, Tao
Dong, David Gang, Shulin Chen. Regulatory mechanism of starch and lipid accumulation in a
microalga strain Chlorella sorokiniana (submitted).
An overall summary was developed as Chapter 6 that highlights the main conclusions of the
work.
12
CHAPTER TWO
HIGH PRODUCTIVITY OF A HEAT-RESISTANT MICROALGA CHLORELLA
SOROKINIANA FOR BIOFUEL PRODUCTION
2.1 ABSTRACT
To augment biomass and lipid productivities of heterotrophic cultured microalgae Chlorella
sorokiniana, the influence of environmental temperature and medium factors, such as carbon
source, nitrogen source, and their initial concentrations was investigated in this study. The
microalga C. sorokiniana could tolerate up to 42 oC and showed the highest growth rate of 1.60
d‒1 at 37 oC. The maximum dry cell weight (DCW) and corresponding lipid concentration was
obtained with 80 g L‒1 of initial glucose and 4 g L‒1 of initial KNO3 at 37 oC. In 5-litre batch
fermentation, the DCW increased dramatically from 0.9 g L‒1 to 37.6 g L‒1 in the first 72 hrs
cultivation, with the DCW productivity of 12.2 g L‒1 d‒1. The maximum lipid content of 31.5%
was achieved in 96 hrs and the lipid productivity was 2.9 g L‒1 d‒1. The results showed C.
sorokiniana could be a promising strain for biofuel production.
Keywords
Chlorella sorokiniana; algae; heterotrophic; heat-resistant; lipid
13
2.2 INTRODUCTION
Biofuel is an alternative to fossil fuel, which is biodegradable, renewable, and non-toxic. It also
contributes no net carbon dioxide or sulfur to the air and emits less atmospheric pollutants.
Biofuel is mainly produced from vegetable oils, such as soybean oil, sunflower oil, palm oil and
rapeseed oil (Kulkarni & Dalai, 2006). However, those sources can lead to competition with
edible food, and cannot realistically satisfy the potential market for biofuel even not designated
for other utilization. Microalgae have been reported as a promising feedstock for biofuel
production because of their ability to grow on non-arable land, direct CO2 mitigation, high
photosynthetic efficiency, rapid growth rate and high lipid content. Lipid content in microalgae
can exceed 80% by weight of dry biomass, and 20% – 50% is quite common. The average oil
yield per hectare could far exceed what can be generated from soybeans or other oil crops (Chisti,
2007). However, to date, phototrophic cultivation of microalgae, such as Arthrospira, Chlorella,
Dunaliella, Aphanizomenon, Haematococcus, Crypthecodinium, and Schizochytrium are only
commercialized for high value-added products due to some limiting factors, i.e., low growth rate
and light penetration (Spolaore et al., 2006). Alternatively, heterotrophic cultivation of
microalgae eliminates the requirement for light, allows rapid production of biomass with high
cell density and high lipid content, and facilitates large-scale production of microalgae biomass
(Xiong et al., 2008). Xiong et al. (2008) reported that the cell density of Chlorella protothecoides
could reach 51.2 g L‒1 after 167 hrs fed-batch cultures with 50.3% lipid content. In addition,
lipids extracted from heterotrophically cultured microalgal cells had similar properties to diesel
fuel in terms of oxygen content, heating value, density and viscosity (Miao & Wu, 2004).
In our previous research, we have successfully established a two-stage heterotrophic and
phototrophic microalgae culture system, which combines the benefits of low processing costs of
14
phototrophic growth and high-cell density cultivation of heterotrophic growth (Zheng et al.,
2012). To enhance the overall productivity, it’s necessary to optimize the heterotrophic cultural
process. In this study, we investigated the potential of C. sorokiniana for biofuel production,
mainly focusing on the effects of environmental temperature and media nutrient. Temperature is
an important and easily-controlled environmental factor affecting algal growth and the formation
of temperature-dependent components within the cells such as fatty acids. Many microalgae have
the ability to grow over a wide temperature range, which is particularly true for Chlorella sp. that
can adapt to 5 oC – 42 oC. Yet for the optimal growth temperature, it is species and/or strain
specific. For example, C. vulgaris growth was negatively affected at temperature above 30 oC,
and further temperature increase (38 oC) led to cell death (Converti et al., 2009). For C.
sorokiniana UTEX 2805, it can grow better under 40 oC – 42 oC than lower temperature (28 oC),
while for C. sorokiniana 211-32 (SAG) the maximum biomass was obtained under 28 oC
(Cordero et al., 2011; de-Bashan et al., 2008). As for lipid content there is no general correlation
with growth temperature as it varies from species to species. In the study of Seto et al. (1984), an
increase of the cultivation temperature from 20 oC to 25 oC led to 60% total lipid content
increase of C. minutissim, while for C. vulgaris increased temperature from 25 oC to 30 oC led to
a decrease of lipid content from 14.7% to 5.9% (Seto et al., 1984). Between the fatty acid
characteristics and growth temperature is also not a pattern.
Therefore, environmental temperature’s effect on microalgal growth is definitely one of the
major factors that need to be evaluated. Besides, nutrient media characteristics, such as carbon
source, nitrogen source, and their initial concentrations, are also extremely important factors
affecting cell growth and lipid accumulation. The effective and reliable mathematical methods –
15
back propagation artificial neural network (BP ANN) and genetic algorithm (GA) were used to
search and confirm the optimal conditions and maximum productivity.
16
2.3 MATERIALS AND METHODS
2.3.1 Organism and medium
The green microalga Chlorella sorokiniana (UTEX 1602) was obtained from the Culture
Collection of Alga at the University of Texas (Austin, TX, USA). This strain was maintained at 4
C on an agar slant of Kuhl medium (Kuhl & Lorenzen, 1964) supplemented with 10 g L‒1
o
glucose.
The Kuhl medium consisted of (per L) 1000 mg KNO3, 621 mg NaH2PO4·H2O, 89 mg
Na2HPO4·2H2O, 246.5 mg MgSO4·7H2O, 9.3 mg EDTA, 0.061 mg H3BO3, 14.7 mg
CaCl2·2H2O, 6.95 mg FeSO4·7H2O, 0.287 mg ZnSO4·7H2O, 0.01235 mg (NH4)6Mo7O24·4H2O,
0.169 mg MnSO4·H2O, and 0.00249 mg CuSO4·5H2O, respectively. Unless otherwise specified,
in heterotrophic cultivation the inocula were prepared in a 250-ml flask containing 50 ml Kuhl
medium supplemented with 20 g/L glucose in our experiments.
2.3.2 Shake-flask cultures
To find the optimal heterotrophic culture condition, the following factors were studied in 250-ml
flasks: 1) Temperature: the inoculated flasks were incubated at 21 oC, 30 oC, 37 oC, 40 oC, 42 oC,
and 45 oC, respectively. 2) Carbon source: the media containing 1 g L‒1 KNO3 included either 20
g L‒1 glucose, fructose, mannose, acetate, lactose, galactose, glycerol and sucrose. 3) Nitrogen
source: the media containing 20 g L‒1 glucose included either peptone, NH4Cl, KNO3, urea,
glycine, NH4NO3 and yeast extract, with the same nitrogen concentration of 0.14 g L‒1 in each. 4)
Initial carbon and nitrogen concentration: five different glucose concentrations (20, 40, 60, 80,
100 g L‒1) were combined with five different KNO3 concentrations (1, 2, 4, 6, 8 g L‒1) in Kuhl
media, respectively. After inoculation with 5% (v/v) exponentially growing inocula, except for
different temperature trial, all the following cultural experiments were conducted at 37 oC with
17
orbital shaking at 150 rpm in darkness. For each experimental condition, three replicates were
performed, and the standard deviation was calculated.
2.3.3 Batch culture
Batch culture of heterotrophic C. sorokiniana was carried out in a 5-L bioreactor containing 3-L
medium. The medium pH value was kept at 6.1 by feeding with 1 mol/L NaOH or H2SO4
solution; dissolved oxygen concentration was controlled by increasing agitation speed and
airflow to keep it around 50% saturation; temperature was maintained at 37 oC.
2.3.4 Analytical procedure
To determine the dry cell weight (DCW), a 5 mL cell suspension sample was centrifuged at the
speed of 1000 × g for 5 min. The cell pellet was washed twice with distilled water, and then
dried in a pre-weighed aluminum dish at 105 °C for 3 hrs, and were subsequently cooled down to
room temperature in a desiccator before weighing. To measure the cell growth, the OD600 of cell
suspensions was evaluated and correlated with the DCW (g L‒1) by a regression equation (1):
DCW = 0.4483OD600 – 0.5285, R2 = 0.991
(1)
The specific growth rates (µ) were calculated using equation (2):
µ
(
)
(
)
(2)
where t0 and t1 is the beginning and end of the logarithmic growth phase, and N0 and N1 is the
microalgal density at the time of t0 and t1, respectively. The fatty acid was determined by gas
chromatograph as described as in reference (Zheng et al., 2012).
The glucose concentrations were analyzed by a Dionex ICS-3000 ion chromatography system
(Dionex Corporation) equipped with a CarboPac TM PA 20 (4×50mm) analytical column and
18
CarboPac TM PA 20 (3×30mm) guard column. Samples were filtered with 0.2μm pore-size filter
before injection and eluted with 0.01 M NaOH at a flow rate of 0.500 ml min‒1. The analytes
were detected and quantified against standard curves by electrochemical detection in a pulsed
amperometric detector.
Nitrate-nitrogen was determined by the H2SO4-salicylic acid method as described by Cataldo
et al. (1975).
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.
2.3.5 Model analysis
BP is a commonly used learning algorithm in artificial neural network (ANN) applications which
consists of input layer, hidden layer and output layer. In this study, the relationship between
DCW and four variables including temperature, initial glucose concentration, initial KNO3
concentration, and time were dealt with. Therefore, there are four nods in the input layer, ten
nods in the hidden layer and one in the output layer. Considering that the growth of algae is
known to be governed by highly nonlinear process, the logsig function is adopted as the
activation function in the hidden layer and the purelin function in the output layer. In order to
improve the generalization performance of the network, the Levenberg-Marquart algorithm was
used to prevent the network from being over trained. The performance of the BP model was
assessed using the MSE (Mean Square Error) as a measure of goodness-of-fit.
Genetic algorithm (GA) is a stochastic optimization technique that searches for an optimal
value of a complex objective function and is used to solve complicated optimization problems by
simulation or mimicking a natural evolution process. Based on the correlation and prediction of
BP ANN model, GA code was used to optimize the conditions of algal cultivation and search the
19
maximum productivity. The output of prediction from BP ANN model was set as fitness function.
The population size was 200; the cross fraction was 0.3; the mutation fraction was 0.1 and the
maximum generations were 1000. BP ANN model and GA code were written in M file and
implemented in Matlab R2010a.
20
2.4 RESULTS AND DISCUSSION
2.4.1 Effects of temperature
The growth curves under different cultivation temperatures are shown in Figure 2.1, and values
of the corresponding growth parameters are tabulated in Table 2.1. After 4 days of cultivation,
almost all C. sorokiniana cultures grew into the stable stage. It was observed that the highest
growth rate (1.60 d‒1), DCW (7.73 g L‒1) and DCW yield (0.37 g g‒1), lipid (2.21 g L‒1) and lipid
yield (0.11 g g‒1) were all obtained at 37 oC. At the temperature range between 21 oC and 37 oC,
all of those parameters had a positive correlation with temperature; yet further increase in
temperature from 37 oC to 42 oC led to certain inhibition of cell growth and no growth was
observed when the temperature reached 45 oC. However, even at 40 oC and 42 oC, DCW still
reached 4.79 g L‒1 and 4.18 g L‒1, respectively, which was much higher compared with 2.77 g L‒
1
of 21 oC. So, the microalga C. sorokiniana could tolerate temperatures as high as 42 oC, but its
optimal growth temperature was 37 oC.
The results indicate that heterotrophic C. sorokiniana has the ability to tolerate heat. It has
been well documented that of all Chlorella species, C. sorokiniana Shihira et Krauss was the
most resistant to heat and high light intensity (de-Bashan et al., 2008). Normally, 20 °C – 30 °C
is the optimal temperature for many commercial microalgae, such as Arthrospira (Sánchez-Luna
et al., 2007), Chlorella (Prescott, 1968), and Dunaliella (García et al., 2007). However, microbial
heat will be generated during fermentation and the heat production rate is positively associated
with culture volume and cell density, therefore, specifically designed cooling system is required
for the large scale high density fermentation of some algal strains (Starfelt et al., 2010). The
heat-resistant characteristics of C. sorokiniana can not only help the algae to reach a much higher
growth rate and biomass productivity, but potentially reduce the energy consumption and cost
21
DCW (g L-1)
8
6
4
2
0
0
1
2
3
4
Cultivation time (d)
Figure 2.1 Cell growth curves of C. sorokiniana for heterotrophic cultivation at various
temperatures. (21 oC, closed circle; 30
o
C, open circle; 37
o
C, closed triangle; 40 oC, open
triangle; 42 oC, closed square; 45 oC, open square)
that would be incurred when cooling fermentors in large scale production schemes. This ability
also provides this algal strain with the possibility of being used as a candidate strain for
simultaneous saccharification and fermentation techniques with cellulosic materials as feedstock.
For the effect of temperature on lipid content, as shown in Table 2.1, the highest lipid content
was achieved at 30 oC, which was different from the biomass production.
However, the
temperature with 37 oC still resulted in the highest lipid production of 2.21 g L‒1. As the growth
temperature increased from 21 oC to 30 oC, the total lipid content increased from 22.80% to
37.04%; as the growth temperature exceeded 37 oC, the total lipid content decreased dramatically
from 30.05% to 3.82%, so the high temperature exhibited negative effects on lipid accumulation.
22
Table 2.1 Cell growth parameters of C. sorokiniana at various temperatures a
Parameters
Specific growth rate µ (d-1)
DCW yield (g g-1)b
DCW (g L-1)
Lipid content (%)
Lipid (g L-1)
Lipid yield (g g-1) b
a
21 oC
1.02
0.17
2.77
22.80
0.67
0.04
30 oC
1.41
0.28
5.12
37.04
1.95
0.10
37 oC
1.60
0.37
7.73
30.05
2.21
0.11
40 oC
1.26
0.26
4.79
10.90
0.52
0.03
42 oC
1.35
0.21
4.18
3.81
0.15
0.01
45 oC
-c
-
The values are averages of three independent measurements with significance declared at
P < 0.05.
b
Biomass or lipid yield on sugar consumed
c
The algae did not grow at temperature 45 oC
In Seto et al. (1984), it was found that higher temperature (from 20 oC to 25 oC) was favorable
for lipid accumulation in C. minutissim. Other studies documented that low temperature would
induce the lipid accumulation and result in a high lipid content in C. vulgaris (from 25 oC to 30
o
C), Scenedesmus sp. LX1 (from 10 oC to 30 oC), Thraustochytrium aureum ATCC 34304 (from
4 oC to 32 oC), and Nannochloropsis oculata (from 15 oC to 20 oC) (Converti et al., 2009; Hur et
al., 2002; Xin et al., 2011). Chen et al. (2008) (Chen et al., 2008) reported that the cultivation
temperature (from 15 oC to 27 oC) had little effect on the lipid content of microalgal biomass,
which was agreement with Amaretti et al. (2010) on the oleaginous yeast (Amaretti et al., 2010),
where the temperature ranging from ‒3 oC to 20 oC did not influence the lipid production of
Rhodotorula glacialis. Our results showed the lipid content increased from 21 oC to 30 oC and
then decreased in the range of 30 oC to 42 oC. In summary, the temperature played a significant
role on the lipid accumulation of the alga C. sorokiniana.
23
Table 2.2 Fatty acid profiles of heterotrophically cultured algae C. sorokiniana at different
temperaturesa
Fatty acids
C14:0
C16:0
C16:1
C16:2
C18:0
C18:1
C18:2
C18:3
TFAb
SFAc
MUFAd
PUFAe
Iodine
value
a
21 oC
0.19
22.42
2.94
3.33
1.17
24.46
26.99
14.00
22.80
23.92
27.92
44.56
118.74
30 oC
0.11
21.38
3.88
1.54
1.19
32.78
30.28
5.73
37.04
23.45
37.62
37.94
106.91
37 oC
0.12
20.06
4.51
1.39
2.18
36.33
27.58
2.85
30.05
23.94
41.58
32.69
97.00
40 oC
0.28
22.07
2.73
5.34
1.78
21.63
34.47
3.82
10.90
25.62
25.92
43.73
105.75
42 oC
0.28
23.41
2.81
5.04
1.71
23.86
33.79
3.44
3.81
26.52
27.77
42.47
104.94
The values are averages of three independent measurements with significance declared at
P < 0.05.
b
total fatty acids (g)/dry cell weight (g) × 100%.
c
percentage of saturated fatty acids (% of total fatty acids).
d
percentage of monounsaturated fatty acids (% of total fatty acids).
e
percentage of polyunsaturated fatty acids (% of total fatty acids).
The fatty acid composition under different cultivation temperatures is shown in Table 2.2. The
major fatty acids components produced by microalgae C. sorokiniana were palmitic acid (C16:0),
oleic acid (C18:1), and linoleic acid (C18:2). Within the temperature range of 21 oC – 42 oC, no
significant change on the contents of total unsaturated fatty acids were detected, but the ratio of
24
monounsaturated fatty acids (MUFA) increased and the polyunsaturated fatty acids (PUFA)
decreased from 21 oC – 37 oC. The degree of unsaturation, which was reflected by the calculated
iodine values, decreased from 21 oC to 37 oC and increased from 37 oC to 42 oC. A similar trend
was observed in Patterson’s (1970) research (Patterson, 1970), where by increasing heterotrophic
growth temperature of C. sorokiniana (Shihira and Krauss) from 22 oC to 38 oC resulted in a
decrease in unsaturation. The decreased degree of unsaturation with the increase of growth
temperature was widely observed in many other organisms because the synthesis of unsaturated
fatty acids under lower temperature could help the cell maintain the proper membrane fluidity
and functions in cold environment (Gounot, 1991). The saturation and the composition of the
fatty acids in algal oil are critical for making biofuel. As shown in table 2.2, most parameters of
the oil from C. sorokiniana, such as the iodine value, the PUFA (at least four double bonds) and
the linolenic acid (C18:3) contents (except at 21oC) were complied with the European biodiesel
standard EN 14214:2008. On the other hand, the effects of temperature on the lipid synthesis of
C. sorokiniana were obvious, which led to a wide span in oil properties, i.e., the iodine value
(97–119) and linolenic acid (3–14). Therefore, the growth temperature has the potential to be
used as a tool to adjust biofuel characteristics to meet different requirements.
2.4.2 Effects of carbon sources
To figure out the influences of carbon sources on biomass and lipid production in C. sorokiniana,
four monosaccharides (glucose, fructose, mannose and galactose), two disaccharides (lactose and
sucrose), acetate and glycerol were investigated and the results are shown in Figure 2.2A. The
ability of C. sorokiniana to utilize carbon sources was obviously quite different, as reflected by
the different DCW. Glucose showed the best results for heterotrophic growth of C. sorokiniana
with 7.38 g L‒1 DCW. Microalgal cells cultured with fructose, acetate, galactose and sucrose
25
produced less DCW of 0.95 g L‒1, 2.00 g L‒1, 1.79 g L‒1, and 2.65 g L‒1, respectively. Mannose
and glycerol gave the lowest DCW of 0.36 g L‒1 and 0.34 g L‒1, respectively.
8
(A)
DCW (g L-1)
6
4
2
0
Glucose Fructose Mannose Galactose Lactose Sucrose Glycerol Acetate
Carbon Sources (20 g L-1)
8
(B)
DCW (g L-1)
6
4
2
0
Peptone Yeast extract KNO3
Urea
Glycine NH4NO3
NH4Cl
Nitrogen Sources (0.01 M N)
Figure 2.2 DCW of C. sorokiniana under different carbon sources and nitrogen sources. (A) the
optimization results of carbon sources with 1 g L-1 KNO3 as nitrogen; (B) the optimization
results of nitrogen sources with 20 g L-1 glucose as carbon.
It is generally accepted that glucose is the most commonly used carbon source for
heterotrophic cultures of microalgae, yet its uptake still depends on the light and the specific
26
algae species (Perez-Garcia et al., 2011). Some algae species, such as Agmenellum
quadruplicatum, Goniotrichium elegans, Navicula pelliculosa, and Nostoc sp. can only absorb
glycerol as carbon source, with the condition of light and no external CO2 supply. O’Grady and
Morgan (2011) reported glycerol had similar growth rate and lipid yield compared with glucose.
In our research, glucose was the most suitable carbon source for heterotrophic culture of C.
sorokiniana.
2.4.3 Effects of nitrogen sources
Besides carbon, nitrogen is another important component contributing to algal dry weight
accumulation. Among organic nitrogen sources (glycine, peptone and yeast extract) and
inorganic nitrogen sources (KNO3, urea, NH4Cl and NH4NO3), KNO3 gave the best result in
producing biomass (6.81 g L‒1) (Figure 2.2B). Glycine and urea could also serve as suitable
nitrogen sources, providing 5.77 g L‒1 and 5.29 g L‒1, respectively. Interestingly, C. sorokiniana
algal cells cultured with complex nitrogen source peptone and yeast extract have poor biomass
production capability (2.51 g L‒1 and 3.08 g L‒1, respectively). This was not in agreement with
the conclusion that the complex nitrogen source, such as yeast extract, tryptone and peptone,
might be superior to simple nitrogen source in heterotrophic culture of microalgae, since they
can provide amino acids, vitamins and other growth factors (O’Grady & Morgan, 2011).
Generally speaking, ammonium is the most preferred nitrogen source for microalgae Chlorella
sp. (Perez-Garcia et al., 2011). Yet in this study both ammonium nitrogen resource NH4Cl and
NH4NO3 was found to hardly support the growth of algae, with 0.74 g L‒1 and 0.82 g L‒1 DCW,
respectively. The reason for those observations might be related to the pH decrease due to the
assimilation of ammonium ion, since a final pH 4.0 with ammonium was observed in flask
27
cultures without automatic control of pH. Shi et al. (2000) also reported the similar problem
during algal growth with ammonium as nitrogen source.
2.4.4 Effects of initial carbon and nitrogen concentrations
To avoid glucose and/or nitrogen becoming limiting factor or having negative effects on algal
growth, and to guarantee carbon and nitrogen supply in the medium for a high cell-density
culture in a batch system, the tolerance of C. sorokininana to high concentrations of glucose and
nitrate was therefore studied. It was found that the highest DCW and total lipid content were
33.46 g L‒1 and 40.64%, respectively, in the flask culture at 80 g L‒1 glucose and 4 g L‒1 KNO3.
The maximum DCW increased with the initial glucose concentration ranging from 20 g L‒1 to 80
g L‒1, then leveled off at the initial glucose concentration of 100 g L-1 (Table 2.3). Yet, the lagphase increased, and the time to attain the maximum DCW was extended (Figure 2.3). For the
effects of varying initial nitrogen concentrations on DCW, the results showed there was no
inhibitory effect under 20 g L‒1, 40 g L‒1, and 60 g L‒1 initial glucose concentrations. On the
contrary, the increase in initial nitrate concentration leads to an increase in the maximum DCW.
When the initial glucose concentration increased up to 80 g L‒1 and 100 g L‒1, inhibitory effects
appeared at or above 6 g L‒1 initial nitrate concentrations (Figure 2.3). This pattern of growth on
nitrogen was very similar with C. protothecoides, for which the increased initial nitrogen
concentration leads to an increase in the resulting cell concentrations, except there was 25‒ 46%
shortened fermentation cycle for C. protothecoides (Shi et al., 2000).
28
25
12
20
DCW (g L-1)
DCW (g L-1)
(B)
(A)
10
8
6
4
15
10
5
2
0
0
0
1
2
3
0
4
1
2
Time (d)
30
(C)
4
(D)
40
25
20
DCW (g L-1)
DCW (g L-1)
3
Time (d)
15
10
30
20
10
5
0
0
0
1
2
3
4
0
5
1
2
3
4
5
6
7
Time (d)
Time (d)
30
(E)
DCW (g L-1)
25
20
15
10
5
0
0
1
2
3
4
5
6
7
Time (d)
Figure 2.3 Cell growth curves of C. sorokiniana for heterotrophic cultivation at initial glucose
concentration of 20 g L-1 (A), 40 g L-1 (B), 60 g L-1 (C), 80 g L-1 (D), and 100 g L-1 (E). Symbols
for initial KNO3 concentrations in (A), (B), (C), (D), and (E): closed circle, 1 g L-1; open circle, 2
g L-1; closed triangle, 4 g L-1; open triangle, 6 g L-1; closed square, 8 g L-1.
29
Table 2.3 Effects of initial glucose and KNO3 concentrations in heterotrophic culture of C.
sorokinianaa
Initial
Initial KNO3
glucose
concentration
concentration
(g L-1)
(g L-1)
1
2
20
4
6
8
1
2
40
4
6
8
1
2
60
4
6
8
1
2
80
4
6
8
1
2
100
4
6
8
a
DCW
(g L-1)
7.09
10.55
10.18
10.73
10.37
6.16
13.07
14.83
18.89
19.43
6.67
13.75
22.92
25.43
24.36
7.41
15.82
33.46
25.35
22.28
6.64
16.02
24.20
24.43
20.59
Lipid content
(%)
Lipid
( g L-1)
30.83
43.04
13.19
10.36
9.38
28.09
32.38
36.69
15.48
16.67
25.97
32.29
28.09
23.63
27.84
31.57
41.00
40.64
31.60
33.95
30.71
40.11
34.74
33.70
31.69
2.19
4.54
1.34
1.11
0.97
1.73
4.23
5.43
2.92
3.24
1.72
4.23
5.93
5.40
6.61
2.34
6.48
13.78
8.01
7.59
2.04
6.42
8.40
8.22
6.52
The values are averages of three independent measurements with significance declared at
P < 0.05.
2.4.5 Validating the optimal conditions using BP ANN model and genetic algorithm
To confirm the optimal heterotrophic culture condition we determined in this study, the
relationship between DCW and four variables including temperature, initial glucose
30
concentration, initial KNO3 concentration, and time was predicted using BP ANN model (Figure
2.4). To construct the BP ANN model, the modeling data were randomly divided into the data
(238 sets) for training and the data (20 sets) for test and validation. A good agreement is
obtained between experimental data and prediction. The coefficients of determination (R2) of
training, test and validation are close to or higher than 0.98. The output of prediction from BP
ANN model was put into GA code as a fitness function. After globe searching, the maximum
DCW productivity is 34.01g L‒1. The optimal conditions are determined as 36.34 oC, 83.99 g L‒1
of carbon source, 3.32 g L‒1 of nitrogen source, and 5.16 day. These prediction data were
extremely close to and therefore validated the experimental results.
Figure 2.4 Comparison of the BP neural network prediction and experimental data.
31
2.4.6 Batch culture in 5-L bioreactor
Based on the optimized parameter in shaking flasks, batch fermentation was carried out in a 5-L
bioreactor to enhance C. sorokiniana growth to higher cell densities. The consumption of
glucose and KNO3, and the accumulation of biomass and lipid of C. sorokiniana at different
culturing times are shown in Figure 2.5. The alga grew slowly during the first 12 hrs where the
5
1
0
Glucose (g L-1)
2
60
40
40
30
20
20
10
0
0
0
12
24
36
48
60
72
84
96
12
10
8
6
4
Total lipid (g L-1)
3
50
DCW (g L-1)
KNO3 (g L-1)
4
80
2
0
108
Time (h)
Figure 2.5 Cell growth and substrate consumption of heterotrophic C. sorokiniana in batch
culture (closed circle, DCW; open circle, total lipid; closed triangle, glucose; open triangle,
KNO3).
DCW increased from 0.9 g L‒1 to 2.0 g L‒1. In the next 60 hrs, the DCW increased exponentially
with a productivity of 0.6 g L‒1 h‒1. The consumption of glucose by C. sorokiniana responded
well to the cell growth, which showed little variations within the beginning 12 hrs and dropped
quickly through the log phase. At the end of cultivation, glucose was consumed completely. For
nitrate, it was consumed quickly and almost run out within 36 hrs, which was at least 48 hrs
32
ahead of glucose. As the nitrate ran out, the total lipid content began to increase linearly to 11.4 g
L‒1, which resulted in a lipid productivity of 0.15 g L‒1 h‒1 from 24 hrs to 96 hrs. The overall
yields of biomass and lipid on glucose were 0.50 g g‒1 and 0.15 g g‒1. In summary, DCW
achieved 37.6 g L‒1 in 72 hrs, and DCW productivity was 12.2 g L‒1 d‒1. Remarkably, DCW
productivity of C. sorokiniana in this study was much higher than that of other oleaginous
microalgae strains, even higher than reported microalgal strains such as Ulkenia SAM 2179,
Schizochytrium sp. and Crypthecodinium cohnii (Table 2.4), which have been commercialized
for DHA-rich oil production, although the inoculum quantity used in our research was far less
than that of the others. It has been reported that a small quantity of inoculum could obviously
postpone microbial growth, then extending the culture time, and finally decreasing biomass
productivity (Lachhab et al., 2001). However, in our study, the culture time needed to reach the
maximum DCW was just 72 hrs with a small inoculum size, which could be contributed to
higher growth rate under higher culture temperature (37 oC) (Table 2.1). Due to the short
cultivation time, C. sorokiniana exhibited the highest DCW productivity but the lipid content
was only 31.5%, which was far less than other strains and also lower than that cultured in flasks
at the same glucose and KNO3 levels (Table 2.3). This might be due to the pH difference in
culture medium, which was controlled at 6.1 in the fermentor, and was observed at over 7.0
without control in the flask cultures (Zheng et al., 2013).
2.5 CONCLUSION
After considering effects of temperature, carbon sources, nitrogen sources, and their initial
concentrations in heterotrophic cultivation process of C. sorokiniana, the maximum DCW and
the lipid content was obtained with 80 g L‒1 of initial glucose and 4 g L‒1 of initial KNO3 at 37
C. In 5-liter batch fermentation, much higher yield (37.6 g L‒1) and productivity (12.2 g L‒1 d‒1)
o
33
of DCW was obtained, with lipid productivity of 2.9 g L‒1 d‒1. Fed-batch culture will be
employed in future work to enhance DCW and lipid productivities. Chlorella sorokiniana has
high potential for biofuel production.
34
Table 2.4 Comparison of results of batch-mode culture using glucose as substrate in this work and previous reports
Strain
Temperature Inoculum Culture DCW
(oC)
(g L-1)
time (h) (g L-1)
Lipid
content
(%)
Lipid
(g L-1)
Lipid
productivity
(g L-1 d-1)
0.5
57.8
1.8
0.3
(Xiong et
al., 2008)
Reference
35
Chlorella
protothecoides
28
0.3
140
Crypthecodinium
cohnii
27
1.5
74
27.7
9.0
12.3
3.4
1.1
(de Swaaf
et al., 1999)
Ulkenia SAM
2179
28
-a
166
68.7
9.9
52.1
35.8
5.2
(Kiy et al.,
2007)
Schizochytrium
limacinum SR21
28
-a
125
59.2
11.4
70.3
41.6
8.0
(Yaguchi et
al., 1997)
Chlorella
sorokiniana
37
0.9
72
37.6
12.2
31.5
11.4
2.9
This work
a
means not detected or mentioned
3.2
DCW
productivity
(g L-1 d-1)
CHAPTER THREE
MIXOTROPHIC CULTIVATION OF A CHLORELLA SOROKINIANA STRAIN FOR
ENHANCED BIOMASS AND LIPID PRODUCTION
3.1 ABSTRACT
Because
some
algae
can
be
grown
photoautotrophically,
heterotrophically,
and/or
mixotrophically, they show promise for lipid production as biofuel feedstock with different
conditions. In this study, we analyzed microalga Chlorella sorokiniana for biomass and lipid
production under three culture modes. The best growth performance was obtained under
mixotrophic conditions compared with photoautotrophic and heterotrophic conditions. With the
addition of 4 g L─1 glucose, the specific growth rate (3.40 d─1) and maximum biomass dry
weight (DW, 3.55 g l─1) in the mixotrophic culture were 1.8- and 2.4-fold of those in the
heterotrophic culture, and 5.4- and 5.2-fold of those under the photoautotrophic culture. Most
significantly, the biomass yield based on consumed glucose reached values of up to 0.82 g g ─1,
which was only 0.34 g g
─1
in the corresponding heterotrophic condition. Moreover, the lipid
content in mixotrophic algae also substantially increased. At a light intensity of 50 µmol m─2 s─1,
a lipid content of 45% was achieved compared to only 13% in the heterotrophic culture. The
PSII inhibitor 3-(3,4-dichlorophenyl)-1,1-dimethylurea (DCMU) had little effect on the
maximum DW and lipid content in the heterotrophic culture, but caused a significant decline in
the mixotrophic culture. Mixotrophic cultures supplied with air grew as well as cultures supplied
with 1% CO2. Mixotrophy also changed the temperature profiles for growth, increasing the
optimum from 25 oC to 37 oC. These results indicate that C. sorokiniana may be an ideal
candidate for mixotrophic cultivation that offers great potential in the production of renewable
biomass for bioenergy applications.
36
Keywords: microalgae; Chlorella sorokiniana; mixotrophic; lipid; biofuel.
37
3.2 INTRODUCTION
Microalgae-based biofuel production has advantages over land-based crops in productivity due
to algae’s greater solar energy-conversion efficiency (Melis, 2009). Algae cultures can also
utilize non-arable land and non-potable water sources. The most common method for microalgae
cultivation is photoautotrophic culture. However, a series of challenges must be overcome in
order to grow algae as biofuel feedstock in open ponds at a large scale. For instance, higher
growth rates are often accompanied by lower lipid content (<20% of DW), and strategies to
increase lipid content (40-50% of DW) such as nutrient starvation typically reduce growth rates.
As a result, achieving higher lipid content usually comes at the expense of lower biomass
productivity (Chen et al., 2011). In addition, photoautotrophic cultures tend to result in low cell
densities due to cellular self-shading that hinders light availability in the lower layers of algae.
This leads to unsatisfactory algal biomass productivity and high harvesting costs.
To address these issues, the use of heterotrophic cultures offers an alternative that eliminates
the requirement for light. In addition, since they require only organic compounds as carbon and
energy sources, rapid production of biomass with high cell density and lipid content can occur
(Chen, 1996; Xiong et al., 2008). Cell density and lipid content has been achieved as high as
103.8 g L─1 and 38.7% of DW, respectively, in a 5-L bioreactor using a two-stage fed-batch
strategy (Zheng et al., 2013).
However, heterotrophic cultivation has drawbacks, such as the need to supplement organic
carbon as a substrate, as well as the need for extra energy to supply oxygen in the fermentation
process. Moreover, heterotrophic growth cannot directly take advantage of the photosynthesis
efficiency of photoautotrophic algae. Another drawback of the heterotrophic process is the fact
that part of carbon source is released to the environment in the form of CO2, without utilization.
38
To date, only a small number of microalgal species have been cultured heterotrophically in
conventional bioreactors. The few commercialized heterotrophic processes focus mainly on the
manufacture of polyunsaturated fatty acids, which are traditionally extracted from fish oil
(Spolaore et al., 2006).
Mixotrophic cultivation, in which organic carbon and light energy are simultaneously supplied
to the algae in the system, is another potential mode for biomass and lipid production. This
strategy of fixing CO2 and assimilating organic carbon supplied concomitantly promises to
greatly increase the cell concentration of microalgae. Studies show that some microalgae grow
rapidly and have higher growth rates and biomass yields under mixotrophic conditions than
under heterotrophic conditions. For example, Lee et al. (1996) (Lee et al., 1996) cultured
Chlorella sorokiniana (UTEX1230) mixotrophically in an outdoor, enclosed tubular
photobioreator supplemented with 0.1 M initial glucose. This provided the optimum biomass
productivity of 10.2 g L─1 d─1 and biomass yield of 0.56 g g─1 glucose during the day time, and
5.9 g L─1 d─1 and 0.35 g g─1 glucose during the night. The daily volumetric productivity of
photoautotrophic culture of C. sorokiniana in a similar photobioreactor was found to be about 3
times lower. Moreover, Marquez-Rocha (1999) demonstrated that at any given dilution rate, the
mixotrophic culture of Arthrospira platensis in continuous culture had a higher bioenergetics
yield than the photoautotrophic culture (Márquez-Rocha, 1999). Mixotrophic growth offers
additional potential advantages over photoautotrophic growth, including reduced or even halted
photoinhibition and prohibited photooxidative damages, especially in closed photobioreactors
where oxygen accumulation occurs (Chojnacka & Marquez-Rocha, 2004; Chojnacka &
Noworyta, 2004).
39
Mixotrophy involves complex processes in which light intensity, temperature, organic and
inorganic substrates are critical factors in growth and product accumulation. Previous research
reveals that growth, lipid content, and fatty acid fractions are linked to light intensity and
temperature. However, diverse trends are observed for different algal strains from the Chlorella
genus (Bhatnagar et al., 2011; Li et al., 2012). C. sorokiniana (UTEX 1602) is a thermotolerant
microalgal strain in heterotrophic cultivation, and shows optimal growth with high lipid content
at 37 oC (Li et al., 2013). However, the effect of temperature on mixtrophic culture is unkown.
Organic carbon concentration in mixotrophic growth is another critical factor that plateaus or
exhibits inhibitory effects at high doses of supplementation (Cheirsilp & Torpee, 2012; Liang et
al., 2009).
In mixotrophic growth, organic carbon is used for biomass and energy production via
heterotrophic metabolism, which results in CO2 evolution and may stimulate photoautotrophic
carbon assimilation. Based on this hypothesis, an external CO2 supply may be less necessary for
mixotrophic growth, thereby reducing production costs. To develop an effective system for algal
cultures, the effects of glucose concentration, light intensity, temperature and CO2
supplementation on algae growth performance must be determined.
To meet this call, this study explored the impact of initial glucose concentration, light intensity,
PSII inhibitor 3-(3,4-dichlorophenyl)-1,1-dimethylurea (DCMU), CO2 concentration, and
temperature on the growth of C. sorokiniana under photoautotrophic, heterotrophic and
mixotrophic culture modes. Results highlight the promise of mixotrophic cultivation of
microalgae C. sorokiniana as an efficient strategy to produce microalgal biofuels.
40
3.3 MATERIALS AND METHODS
In the first phase of this study, we characterized the growth of C. sorokiniana under three
different culture modes in a Kuhl basal medium supplemented with glucose at different
concentrations in heterotrophic and mixotrophic conditions. Next, we cultured the algae under
different light intensities to understand the role of light in algal growth and lipid accumulation
under mixotrophic conditions. We further tested the role of photosynthesis in the process of
biomass and lipid accumulation by applying the PSII inhibitor DCMU, which inhibits
photosynthesis by interrupting the flow of electrons from PSII to plastoquinone (Rippka, 1972).
We also tested the hypothesis that CO2 supply might be less important for mixotrophic growth.
Finally, we evaluated the influence of temperature on three different culture modes.
3.3.1 Organism and culture conditions
Green microalgae Chlorella sorokiniana (UTEX 1602) were obtained from UTEX, the Culture
Collection of Algae at The University of Texas at Austin. The strain was maintained at 4 oC on
an agar slant of Kuhl medium (Kuhl & Lorenzen, 1964), which consisted of (per L): 1000 mg
KNO3, 621 mg NaH2PO4·H2O, 89 mg Na2HPO4·2H2O, 246.5 mg MgSO4·7H2O, 9.3 mg EDTA,
0.061 mg H3BO3, 14.7 mg CaCl2·2H2O, 6.95 mg FeSO4·7H2O, 0.287 mg ZnSO4·7H2O, 0.01235
mg (NH4)6Mo7O24·4H2O, 0.169 mg MnSO4·H2O, and 0.00249 mg CuSO4·5H2O.
In routine experiments, inocula were cultured at 25 oC in a 250-ml flask containing 150 ml
Kuhl medium, agitated with filter-sterilized air that was enriched with CO2 to 1% (v/v).
Illumination was provided by cool-white fluorescent lamps to give a constant light intensity of
100 µmol m─2 s─1. For experimental cultures, 2% (by volume, average cell concentration of 50–
80 mg L─1 DW) of exponentially growing inocula were transferred into each flask and grown at
25 oC with illumination on an orbital shaker at 150 rpm. We supplemented the Kuhl medium
41
with a corresponding content of glucose in both the heterotrophic and mixotrophic cultures. For
heterotrophic cultures, flasks were covered with foil. For the DCMU treatment, we added the
compound (in methanol) directly to the growth media, and treated control cells with methanol
alone.
We performed three replicates for each experimental condition and calculated the standard
deviations.
3.3.2 Growth analysis
To determine DW, we centrifuged a 5 mL cell suspension sample at 1000 × g for 5 min. The cell
pellet was washed twice with distilled water, dried in a pre-weighed aluminum dish at 105 °C for
3 hrs, and subsequently cooled to room temperature in a desiccator before weighing. To measure
the cell growth, we evaluated the OD750 of cell suspensions and correlated this with the DW (g
L─1) using a regression equation:
DW = 0.5232OD750 – 0.0248, R2 = 0.999
(1)
The specific growth rate (µ) at the exponential phase was calculated by:

 ln X 2  ln X 1 
t2  t1
(2)
where X2 and X1 are DW (g L─1) at time of t2 and t1, respectively.
3.3.3 Chemical analysis
We analyzed glucose concentrations using a Dionex ICS-3000 ion chromatography system
(Dionex Corporation) equipped with a CarboPac TM PA 20 (4 × 50 mm) analytical column and
CarboPac TM PA 20 (3 × 30 mm) guard column. Samples were filtered with 0.2 μm pore-size
filter before injection, and eluted with 0.01 mol L─1 NaOH at a flow rate of 0.5 ml min─1. The
analytes were detected and quantified against standard curves through electrochemical detection
42
in a pulsed amperometric detector. We also analyzed the cellular lipid content and lipid profile
with gas chromatography, using tridecanoic acid (C13:0) as an internal standard. The
corresponding chromatographic conditions and the extraction/transesterification method are
described in O'Fallon et al. (2007).
3.3.4 Calculation of iodine value
We calculated the iodine value, an indicator of total unsaturation of oil or fat, according to
AOCS recommended practice Cd 1c-85, which estimates grams of halogen absorbed by 100 g of
oil.
43
3.4 RESULTS
3.4.1 Growth and lipid accumulation of microalgae
We measured significant differences in the specific growth rate, maximum DW, and lipid
accumulation for the three culture modes. As shown in Figure 3.1 (A and B) and Table 3.1, the
photoautotrophic cells grew slowly, with a low specific growth rate of 0.63 d─1 and maximum
DW of 0.68 g L─1 after 6 days. In contrast, the algae grew much faster under heterotrophic and
mixotrophic conditions. For heterotrophic growth, maximum DW increased with initial glucose
concentrations from 2 to 10 g L─1. In the culture with 20 g L─1 glucose, the glucose was not
completely utilized, due to the limitation of nitrogen in the medium (data not shown).
Compared with heterotrophic growth, all doses of glucose in the mixotrophic culture led
immediately to logarithmic growth without an obvious lag phase. The highest growth rate and
DW were obtained in the mixotrophic culture. Using 4 g L─1 glucose as an example (Table 3.1),
the growth rate and DW achieved under mixotrophic culture were 3.40 d
─1
and 3.55 g L─1
respectively. These figures were 1.9- and 2.4-fold greater than those in the corresponding
heterotrophic condition. Although the maximum DW dramatically increased when the initial
glucose concentration increased from 2 to 6 g L─1,
no further increase was observed at
concentrations higher than 8 g L─1. It is important to note that at glucose concentrations greater
than 8 g L─1, the contribution of glucose to growth reached a plateau. The highest biomass yield
in the mixotrophic culture was 0.82 g g─1, with supplementation of 4 g L─1 glucose. This was
2.4-fold of that under the corresponding heterotrophic condition.
44
6
A
DW (g L)
5
4
3
2
1
0
0
1
2
3
4
5
6
Time (day)
5
B
DW (g L)
4
3
2
1
0
0
1
2
3
4
5
6
Time (day)
Figure 3.1 Effects of glucose concentrations on the growth of C. sorokiniana under a light
intensity of 100 µmol m─2 s─1 (A) and in the dark condition (B). Symbols for initial glucose
concentrations in (A) and (B ):
, 0 g L─1; ●, 2 g L─1; ○, 4 g L─1; ▼, 6 g L─1;
g L─1; □, 20 g L─1.
45
, 8 g L─1; ■, 10
Table 3.1 Growth performance of C. sorokiniana under three different culture modes for 6 days
at 25 oC, with a light intensity of 100 µmol m─2 s─1 for the former two conditions a
Photoautotrophic
Mixotrophic
Heterotrophic
a
Initial
glucose
(g L─1)
0
2
4
6
8
10
20
2
4
6
8
10
20
Specific
growth
rate (d ─1)
0.63
3.20
3.40
3.10
3.31
3.29
3.25
1.90
1.81
2.07
1.90
1.83
1.94
Max.
Biomass
(g L─1)
0.68
1.66
3.55
4.57
5.01
5.08
4.91
0.73
1.46
2.78
3.62
4.23
4.12
Biomass
yield
(g g ─1) b
─c
0.73
0.82
0.74
0.61
0.57
0.50
0.32
0.34
0.45
0.44
0.43
0.38
The values are averages of three independent measurements with significance declared at
P < 0.05.
b
Biomass yield based on glucose consumed.
c
Cannot be determined.
As shown in Table 3.2, the algae accumulated lipid under all culture conditions. Surprisingly,
the lipid content was considerably high in the mixotrophic mode with increasing glucose
supplies until it reached 8 g L─1. After cultivation for 6 days, the photoautotrophically grown
cells had a total lipid content of 9.0%. The lipid content of heterotrophic cultures ranged from
6.2 to 17.6%, while 13.4 to 34.7% of lipid was achieved under mixotrophic conditions.
46
Table 3.2 Fatty acid profiles of photoautotrophically, heterotrophically and mixotrophically cultured C. sorokiniana.a
47
Fatty
acid
C16:0
C16:1
C16:2
C18:0
C18:1
C18:2
C18:3
TFAb
SFAc
MUFAd
PUFAe
Iodine
value
Photoautotrophic
20.99
5.56
4.82
0.33
2.95
13.79
33.31
8.98
22.32
22.60
52.02
134.84
Heterotrophic, initial glucose (g L─1)
2
4
6
8
10
20
18.05 20.62 25.38 25.45 25.93 25.82
2.72
2.41
2.49
4.31
3.95
3.94
6.54
6.69
6.07
4.11
3.21
3.36
0.75
0.77
0.95
1.45
1.37
1.39
6.12
5.30 13.08 19.53 23.50 22.73
24.57 26.05 28.89 28.31 28.99 28.73
26.56 24.27 14.01 10.24 7.90
8.55
6.25
7.37 12.62 17.63 22.37 21.93
19.70 21.92 26.68 27.19 27.56 27.47
18.04 17.99 23.49 29.44 31.80 31.31
57.67 57.02 48.98 42.66 40.10 40.64
Mixotrophic, initial glucose (g L─1)
2
4
6
8
10
20
24.62 26.15 25.16 23.92 24.20 23.97
3.05
2.46
1.53
1.85
1.78
1.69
6.31
3.00
4.27
3.92
3.97
4.08
1.04
1.18
1.26
1.70
1.76
1.85
7.62
9.48 15.33 20.14 20.06 20.16
24.50 30.30 33.01 32.75 32.43 32.24
20.54 17.53 12.21 9.74
9.83 10.00
13.35 21.39 31.58 34.67 31.86 32.18
26.11 27.67 26.69 25.87 26.20 26.07
19.47 20.10 22.94 27.02 26.92 26.99
51.35 50.83 49.49 46.42 46.23 46.32
138.8
123.1
134.4
117.1
109.4
105.6
106.5
119.8
117.0
a
The values are averages of three independent measurements with significance declared at P < 0.05.
b
Percentage of total fatty acids (% of DW).
c
Percentage of saturated fatty acids (% of total fatty acids).
d
Percentage of monounsaturated fatty acids (% of total fatty acids).
e
Percentage of polyunsaturated fatty acids (% of total fatty acids).
113.9
113.5
113.8
The composition and saturation level of the fatty acids in algal oil are critical in the production
of biofuels. Table 3.2 shows the fatty acid composition of C. sorokiniana. The major fatty acids
components are palmitic acid (C16:0), oleic acid (C18:1), linoleic acid (C18:2), and linolenic
acid (C18:3), which are similar to the fatty acid composition of soybean oil (Canakci & Gerpen,
2001). Generally, increased glucose supplementation in mixotrophic and heterotrophic cultures
had similar effects on both the profiles of fatty acids and the content of each fatty acid
component; therefore, linolenic acid content (C18:3) decreased, while oleic acid content (C18:1)
and linoleic acid content (C18:2) increased. As the initial glucose concentration increased, the
contents of saturated fatty acids (SFA) and monounsaturated fatty acids (MUFA) did not change
significantly in the mixotrophically cultured algae. However, they showed a significant increase
in the heterotrophically cultured algae. The content of polyunsaturated fatty acids (PUFA) in
both heterotrophic and mixotrophic algae decreased significantly with high glucose
concentrations. Since in microalga C. sorokiniana, the proportion of PUFA (largely contributed
by C18:3), decreases with the increase in glucose concentration, the rate of the desaturation
reaction may decrease and the saturated fatty acids are not converted to PUFA (Amaretti et al.,
2010).
The degree of unsaturation, reflected by the calculated iodine values, showed a wide span in
the two culture modes. With the exception of the 2 g L─1 of initial glucose, the iodine values
(less than 120) in the mixotrophic algae complied with the European biodiesel standard EN
14214:2008. The iodine value in mixotrophic algae was generally higher than that in
heterotrophic algae under the same initial glucose concentrations. Low levels of unsaturation in
heterotrophically grown cells may be partly attributed to a reduction in the chloroplast
48
photosynthetic membrane, which contains considerable levels of unsaturated fatty acids (Scheer
& Parthier, 1982).
3.4.2 Effects of light intensity
Although supplemented glucose can be completely consumed under all light conditions in
mixotrophic growth, our results show that light intensity has a crucial effect on biomass
accumulation (Figure 3.2, 3.3). In general, we observed a significantly higher specific growth
rate as light intensities increased from 0 to 200 µmol m─2 s─1. For biomass concentration, the
maximum DW under dark condition and under 50 µmol m─2 s─1 of light intensity were
approximately at the same level. Cell growth peaked at 2.5 days and then declined sharply in the
strictly dark condition. However, under light intensity of 50 µmol m─2 s─1, growth slowly
continued after 2.5 days. Mixotrophic cells grew more rapidly under light intensities between
100 and 200 µmol m─2 s─1 and reached a higher peak at 2.5 days. Specific growth rates increased
with increased light intensity for mixotrophic cultures but decreased for photoautotrophic
cultures (Figure 3.3).
We compared the mixotrophic growth rate and sum of the growth rates obtained in
heterotrophic and photoautotrophic cultures to determine whether mixotrophic growth was
simply due to the combination of heterotrophic and photoautotrophic growth (Figure 3.3). At a
light intensity of 50 µmol m─2 s─1, the mixotrophic growth rate and the sum of the heterotrophic
and photoautotrophic growth rates were approximately the same. However, at light intensities of
100 and 200 µmol m─2 s─1, the cell growth rates in mixotrophic cultures were higher than the
sum.
49
5
A
DW (g L)
4
3
2
1
0
0
1
2
3
4
5
Time (day)
DW (g L)
0.8
B
0.6
0.4
0.2
0.0
0
1
2
3
4
5
6
7
Time (day)
Figure 3.2 Effects of light intensities on the growth of C. sorokiniana with (A) or without (B) 6 g
L─1 of initial glucose. Symbols for light intensity in (A) and (B):
µmol m─2 s─1; ○, 100 µmol m─2 s─1; ●, 200 µmol m─2 s─1.
50
, 0 µmol m─2 s─1; ▼, 50
Specific growth rate (d 1)
5
Mixotrophic
Heterotrophic
Photoautotrophic
A
4
3
2
1
0
50
100
200
Light intensity (mol m2 s1)
50
Lipid content (%)
40
B
30
20
10
0
0
50
100
200
Light intensity (mol m2 s1)
Figure 3.3 Comparison of specific growth rate (A) and lipid content (B) of C. sorokiniana under
various light intensities supplemented with 6 g L─1 glucose.
51
These results indicate that higher light intensities in a certain range can improve the growth of
C. sorokiniana in mixotrophic cultures. For biomass production, a light intensity of 100 µmol
m─2 s─1 resulted in higher growth rate and biomass yields compared with a light intensity of 50
µmol m─2 s─1. However no further increase was observed when light intensity reached 200 µmol
m─2 s─1. Additionally, although increasing light intensity from 50 to 100 µmol m─2 s─1 produced
negative effects on lipid content (Figure 3.3), lipid productivity still improved with higher light
intensity (1.30 g L─1 vs. 1.42 g L─1 under 50 and 100 µmol m─2 s─1, respectively).
3.4.3 Effects of DCMU
When 10 µM DCMU was added, photosynthetic activity in the culture was completely inhibited
and cells could not grow photoautotrophically (data not shown). The effect of DCMU on the
growth of C. sorokiniana is shown in Figure 3.4. The heterotrophic culture was barely influenced
by DCMU, whereas severe inhibition was observed in the mixotrophic culture ‒ the maximum
DW decreased to the same level as the heterotrophically cultured algae, although the growth rate
of mixotrophically grown cells with DCMU was higher than that of the heterotrophic cells.
As shown in Figure 3.4, DCMU did not play a significant role in the lipid content of
heterotrophic cultures. For mixotrophically grown cells, the addition of DCMU led to lipid
content (13.4%) comparable to that of heterotrophic cells, but much lower than that of the
control without DCMU (34.5%).
52
5
A
DW (g L1)
4
3
2
1
0
0
1
2
3
4
5
Time (day)
Lipid content (%)
40
B
30
20
10
0
ic
U
ic
MU
oph
oph
CM
r
r
t
D
t
DC
o
o
+
+
r
x
te
Mi
hic
hic
He
rop
rop
t
t
o
o
x
ter
Mi
He
Figure 3.4 Effects of DCMU supplementation on growth (A) and lipid content (B) of C.
sorokiniana under mixotrophic and heterotrophic conditions with 6 g L─1 glucose. ●,
mixotrophic culture without DCUM; ○, mixotrophic culture with 10 µM DCMU; ▼,
heterotrophic culture without DCMU;
, heterotrophic culture with 10 µM DCMU.
53
3.4.4 Effects of CO2
Under a light intensity of 100 µmol m─2 s─1, C. sorokiniana was cultured with low CO2 (air) or
high CO2 (1% CO2) concentrations photoautotrophically and mixotrophically for 7 days (Figure
3.5). In photoautotrophic culture, enriched CO2 increased DW and lipid content significantly, to
6.3- and 2.2-fold of those in the low CO2 condition. Thus, the availability of CO2 is a limiting
factor for photoautotrophic growth under this light intensity. However, comparable figures for
DW (4.70 g L─1) and lipid content (33.48%) were obtained for the mixotrophic culture with 6 g
L─1 glucose, independent of CO2 concentration.
DW
Lipid content
40
DW (g L1)
4
30
3
20
2
10
1
0
Air
Air
O2
C
+
%
ose
1 %_
c
+1
u
e
1 Gl
s
co
_
L
1 Glu
g
6
L
6g
Lipid content (%)
5
0
CO 2
Figure 3.5 Influence of CO2 content on DW and lipid content of C. sorokiniana under
photoautotrophic and mixotrophic culture conditions, respectively.
54
3.4.5 Effects of temperature
Results show that specific growth rates of mixotrophic and heterotrophic cultures significantly
increased and the photoautotrophic culture decreased at 37 oC compared with 25 oC. Mixotrophic
cultures showed comparable or enhanced results for maximum DW, biomass yield, and lipid
content (Table 3.3). Similar results were also observed for the heterotrophic culture, in
accordance with our previous results.
Table 3.3 Growth performance of C. sorokiniana under three different culture modes at 25 oC for
6 days and at 37 oC for 3 days, with a light intensity of 100 µmol m─2 s─1 a
Culture mode
Temperature
(oC)
Photoautotrophic
25
37
25
Mixotrophic
37
25
Heterotrophic
37
Initial
glucose
(g L─1)
0
0
6
8
10
6
8
10
6
8
10
6
8
10
Specific
growth
rate (d ─1)
0.63
0.40
3.10
3.31
3.29
3.58
3.75
3.72
2.07
1.90
1.83
2.14
2.55
2.62
Max.
Biomass
(g L─1)
0.68
0.27
4.57
5.01
5.08
4.31
6.15
6.04
2.78
3.62
4.23
3.20
4.68
5.67
Biomass
yield
(g g ─1) b
─c
─
0.74
0.61
0.57
0.72
0.78
0.58
0.45
0.44
0.43
0.54
0.59
0.54
TFA
(%)
8.98
─
31.58
34.67
31.86
27.04
31.95
32.40
12.62
17.63
22.37
17.43
25.66
30.67
a
The values are averages of three independent measurements with significance declared at P < 0.05.
b
Biomass yield based on glucose consumed.
c
Cannot be determined.
55
3.5 DISCUSSION
This study compared the growth of C. sorokiniana under photoautotrophic, heterotrophic and
mixotrophic conditions. The mixotrophic growth showed the highest specific growth rate and
maximum DW, which were even higher than the sum of both in the heterotrophic and
photoautotrophic cultures. In addition, the biomass yield based on glucose in the mixotrophic
culture was much higher than that of the heterotrophic culture (Table 3.1). This suggests that
significant photosynthetic reutilization of the CO2 produced from glucose metabolic processes
took place. Traditional methods of measuring photosynthetic rates reveal evidence for the
photosynthetic reutilization of CO2 evolved by respiration. For example, the
14
C method
underestimates the carbon fixation rate because internally recycled carbon dioxide derived from
respiration accounts for over 50% of the photosynthetically fixed carbon in mixotrophic
organisms (Stoecker & Michaels, 1991). Similar results have also been reported in mixotrophic
cultures of cyanobacteria (Scherer & Böger, 1982; Valiente et al., 1992). Valiente et al. (1992)
reported that for cyanobacterium Anabaena variabilis, the rate of
14
CO2 evolved from
radiolabelled fructose was lower in the light than in the dark or in the presence of DCMU.
This study found evidence for the reutilization of CO2, supported by the reduced growth
observed in mixotrophic cultures treated with DCMU. We also found that mixotrophically grown
cells with ambient air accumulated biomass that was similar to that of cells grown with 1% CO2
(Figure 3.5). With 6 g L─1 glucose addition, CO2 derived from metabolism was apparently
sufficient for maximal growth.
In photosynthesis, there is a tendency for rapid O2 accumulation during culturing, particularly
when the photosynthetic rate increases. One would expect that O2 production from
photosynthesis in mixotrophic growth would also result in an increased dissolved oxygen (DO)
56
level, although the level would be lower than that of photoautotrophic cultures due to the oxygen
uptake in glucose metabolism. Yen and Zhang (2011) report that a low DO could retard cell
growth, while an increased DO led to rapid cell growth and a maximum DW in heterotrophic
cultures (Yen & Zhang, 2011). Therefore, O2 production from photosynthesis in mixotrophic
growth can be expected to contribute to rapid growth if the rate of external oxygen supply is not
sufficient to keep up with metabolism. On the other hand, in photoautotrophic cultures, high
oxygen accumulation can impose a more severe impediment for biomass productivity by
inhibiting photosynthesis. Oxygen accumulation is an especially serious problem in closed
photobioreactors (Marquez et al., 1995). O2 reutilization in mixotrophic cultures, however, can
help prevent O2 levels from reaching toxic concentrations and reduce photooxidative damage
(Chojnacka & Marquez-Rocha, 2004).
Our results show that the specific growth rate of C. sorokiniana was strongly affected by the
presence of glucose, light intensity, temperature, and CO2. In the presence of glucose, light
intensity affected the growth rate significantly. At low light intensity, the mixotrophic growth
rate of C. sorokiniana was about equal to the sum of the photoautotrophic and heterotrophic
growth rates (Figure 3.3). This finding on C. sorokiniana accords with previous studies, which
claim that the specific growth rate of Chlorella vulgaris can be enhanced with added glucose as
long as light intensity is not saturating (Ogawa & Aiba, 1981). Similar results have also been
reported for several strains such as unicellular blue-green algae Spirulina platensis (Marquez et
al., 1993), the astaxanthin-producing strain Haematococcus pluvialis (Kobayashi et al., 1992),
and C. vulgaris (Martínez & Orús, 1991).
An especially important finding is that mixotrophic algae exhibited increased specific growth
rates with increased light intensities. It was observed that the mixotrophic growth rate of C.
57
sorokiniana surpassed the sum of the photoautotrophic and heterotrophic growth rates. This
finding accords with results from other strains, such as C. vulgaris (Heredia-Arroyo et al., 2011;
Liang et al., 2009), Chlamydomonas humicola (Lalibertè& de la Noüie, 1993), marine Chlorella
sp. and Nannochloropsis sp. (Cheirsilp & Torpee, 2012). In contrast, photoautotrophic algae
exhibited lower and decreased specific growth rates with increased light intensity (Figure 3.2,
3.3), although they exhibited a linear growth phase at the start, which is a characteristic of
photosynthetic miroorganisms during photoautotrophic growth in batch cultures (Kim et al.,
2002).
Similar results were also observed in marine Chlorella sp. (Cheirsilp & Torpee, 2012;
Vonshak et al., 2000). The decrease in the photoautotrophic growth rate of C. sorokiniana under
high light intensities may be due to photoinhibition, which occurrs when the rate of absorption of
light energy exceeds the rate of its consumption and the CO2 level is low (Takahashi & Murata,
2008). Photoinhibition is greatly reduced in mixotrophic cultures, which can be partly explained
as lower DO levels and lower susceptibility of the mixotrophic cells to light stress due to their
ability to use more light energy and their higher saturation threshold for photosynthetic activity
(Vonshak et al., 2000).
Our results also show that adding DCMU inhibited the noncyclic electron flow of cells due to
the effects of light. However, the specific growth rate of mixotrophic cultured cells was still
higher than that of heterotrophic cells, although the maximum DW was similar to that of
heterotrophic culture (Figure 3.2). Hence, the increased growth rate in the mixotrophic culture
with DCMU reflects cyclic electron flow. Cyclic electron flow has been found to be more
effective in providing energy for sugar uptake, which was higher in the light with DCMU
58
condition than in the dark condition (Valiente et al., 1992), even under anaerobic conditions
(Tanner, 2000).
We found additional evidence that mixotrophy involves a complex combination of both
photoautotrophy and heterotrophy when we evaluated growth performance under different
temperatures. As previously noted, C. sorokiniana showed superior growth at higher temperature
under the heterotrophic condition, and photoautotrophic growth was inhibited at 37 oC (Table
3.3). Photoautotrophic growth depends on the balance between carbon fixation, photorespiration
and respiration. Each of these processes demonstrates different temperature dependencies. This
is especially evident in the oxygenase activity of the Rubisco enzyme (i.e., photorespiration),
which increases with temperature at the expense of its carboxylase activity (Bernacchi et al.,
2001). The increase in photorespiration in combination with enhanced carbon losses by
respiration may explain the observed decline of the photoautotrophic growth rate at higher
temperatures. In contrast, the specific growth rate of mixotrophic algae was stimulated at higher
temperatures, which may be partly attributed to increased heterotrophic metabolic activity. These
results were in accordance with Wilken et al. (Wilken et al., 2013), who showed mixotrophic
organisms became more heterotrophic with increasing temperature.
Studies show that in mixotrophic cultures, the addition of organic substrate increases growth
rates as well as final biomass concentration (Cheirsilp & Torpee, 2012; Heredia-Arroyo et al.,
2011; Marquez et al., 1993). However, this cultivation approach has rarely been applied to study
microalgal oil accumulation. Cheirsilp and Torpee (2012) found that the lipid content of four
microalgal strains did not differ significantly in the three cultivation modes, although the growth
of all strains was improved under mixotrophic conditions. Similar results have also been reported
on Scenedesmus obliquus (Mandal & Mallick, 2009) and Chlorella minutissima (Table 3.4).
59
Table 3.4 Comparison of current research results with previous reports of microalgal cultures under three different culture modes
Strain
Chlorella
minutissima
Chlorella
vulgaris UTEX
2714
Chlorella
vulgaris UTEX
259
60
Chlorella sp.
Chlamydomonas
humicola
Chlorella
sorokiniana
UTEX 1602
Culture
modea
P
H
M
P
H
M
P
H
M
P
H
M
P
H
M
P
H
M
Extraneous
carbon
Extraneous carbon
content (g L─1)
Glucose
Glucose
10
10
Glucose
Glucose
4
4
Glucose
Glucose
10
10
Glucose
Glucose
2
2
Acetateb
Acetate
0.82
0.82
Glucose
Glucose
6
6
Growth
rate (d─1)
Biomass
(g L─1)
Lipid
content (%)
─c
─
─
─
─
─
─
─
─
─
─
─
0.21
0.78
1.66
0.45
1.66
2.68
0.07
0.14
0.38
0.4
0.75
1.4
0.25
1.21
1.70
0.38
0.5
1.38
─
─
─
0.47
2.78
4.57
5.40
14.88
11.79
27.38
30.58
13.82
38
23
21
30
22
26
10.2
5.6
4.8
6.65
12.62
31.58
a
P, photoautotrophic culture; H, heterotrophic culture; M, mixotrophic culture.
b
Sodium acetate.
c
Not detected or mentioned.
Reference
(Bhatnagar
et al., 2010)
(HerediaArroyo et
al., 2011)
(Liang et al.,
2009)
(Cheirsilp &
Torpee,
2012)
(Lalibertè&
de la Noüie,
1993)
This study
For C. vulgaris and Chlamydomonas humicola, the lipid content under photoautotrophic
conditions was higher than those under mixotrophic and heterotrophic conditions. Our results
show that C. sorokiniana achieved the highest lipid content in the mixotrophic condition,
followed by that of the heterotrophic and photoautotrophic conditions. Thus, lipid accumulation
behavior appears to be strain-dependent, although experimental differences could complicate this
interpretation.
The biosynthesis of fatty acids is not only a carbon-demanding process, but also requires
significant metabolic energy. In photoautotrophic cultures, photosynthetic electron transport is
the primary source of ATP and NADPH for fatty acid synthesis, whereas in heterotrophic
cultures, these cofactors are provided by organic carbon. In this study, lipid content increased
with increasing initial glucose concentration in both the heterotrophic or mixotrophic cultures.
Photoautotrophically grown C. sorokiniana supplemented with 1% CO2 showed much higher
lipid content than the cells cultured with air. Clearly, lipid accumulation in C. sorokiniana is
dependent on carbon availability. With the same initial glucose content, mixotrophic algae
displayed much higher lipid content than heterotrophic algae. One explanation for the higher
lipid content in mixotrophic cultures may be the increased availability of carbon from refixation
of CO2 released from organic carbon metabolism process. However, the extra carbon made
available by refixation could not completely explain the increased lipid content. Increased lipid
content also appeared to be a light-correlated process in which the noncyclic electron flow
through photosystem I and photosystem II responsible for ATP and NADPH formation plays a
significant role. The above conclusions were supported by the severe inhibition of DCMU,
which showed that the lipid content of mixotrophic cultures with added DCMU decreased to the
level of heterotrophic cells.
61
Finally, we found that light intensity is another critical factor in this process. As light intensity
increases from 50 to 100 µmol m─2 s─1, the specific growth rate and maximum DW of
mixotrophic algae increases as well. However, the lipid content decreases (Figure 3.3). Similar
results were also reported by Cheirsilp and Torpee (2012). C. sorokiniana may utilize
photosynthetic energy for cell division rather than for lipid accumulation under higher light
intensity. From these results, it can be concluded that the optimal levels of light intensity for
supporting cell growth and lipid accumulation differ. To obtain a high cell dry weight, light
intensity should be increased, although a low light intensity is more favorable for lipid
accumulation. Taken together, these results indicate that increased lipid accumulation in the
mixotrophic condition was not only due to the extra carbon available through refixation, but also
depended on light intensity and the supply of ATP and/or NADPH through the light reactions of
photosynthesis.
For mixotrophic cultivation at large scale, several critical problems still need to be addressed.
First, the significant cost of sugars in the total cost of mixotrophic algae production is a major
drawback. In order to lower production costs, cheaper carbon sources from industrial and
agricultural wastes should be considered (Xu et al., 2006). Second, the appropriate culture
system should be chosen, whether they are open-culture or closed-culture systems called
photobioreactors (PBR). PBRs could offer better control over culture conditions; however,
higher capital and operating costs should be taken into consideration. Open-culture systems are
less expensive to build and operate, yet are normally more susceptible to contamination from
other microorganisms, especially applied with organic carbon. Therefore, further study is
expected to promote the mixotrophic cultivation of C. sorokiniana as a potential technology for
biofuel production at a large scale.
62
3.6 CONCLUSION
In this study, mixotrophic cultures of C. sorokiniana showed the best growth performance, with
the highest growth rates, DW, and lipid productivity compared with other growth modes. The
optimal concentration for biomass and lipid content was 4 to 8 g L─1 of initial glucose. We also
found that efficient utilization of glucose led to an increased biomass yield, which are attributed
to the refixation of CO2 from heterotrophic process. Light intensity had profound impact on
algae biomass and lipid accumulation. The suitable light intensity was found to be 100-200 µmol
m─2 s─1 for all responses. In addition, at higher temperatures, mixotrophic growth showed a
much higher specific growth rate and comparable maximum DW and lipid content compared to
cells grown at 25 oC. This indicates the feasibility of the use of mixotrophic cultures in the
summer season. Therefore, this study provides meaningful insight for exploring efficient cultural
strategy to produce renewable biomass for bioenergy applications.
63
CHAPTER FOUR
PHOTOSYNTHESIS REGULATION IN MIXOTROPHIC CELLS OF
CHLORELLA SOROKINIANA
4.1 ABSTRACT
Mixotrophic growth of microalgae offers great potential as an efficient biomass production
strategy for biofuel production. After cultivation for 1 d, mixotrophic cells of Chlorella
sorokiniana showed the highest photosynthetic activity, with maximum photosynthetic O2
evolution increased by approximately 3 and 4 times, compared with photoautotrophically 1%
CO2-conditioned and air-conditioned cells, respectively. Characteristic of chlorophyll
fluorescence parameters demonstrated that there was no limitation in electron transport
downstream of PSII. The up-regulation of photosynthetic activity was associated with the highest
total ribulose-1,5-bisphosphate carboxylase/oxygenase (Rubisco) carboxylase activity and
expression level of phosphoribulokinase (PRK). Following cultivation after 3 d, photosynthetic
O2 evolution of mixotrophic cells was strongly reduced, accompanied by reduced photochemical
efficiency and reorganization of the PSII complex.
The down-regulation of primary
photosynthetic reactions was accompanied by reduced enzymatic activity levels of Rubisco
carboxylase and mRNA levels of Rubisco and PRK. Importantly, there was almost no nonphotochemical quenching for mixotrophic cells no matter cultured for 1 d or 3 d. It was
interesting that the decline in the quantum efficiency of PSII and oxidized PQ pool was observed
in N-depleted condition during mixotrophic growth. Our results firstly suggest, to the best of our
knowledge, that different regulation pattern of photosynthesis in mixotrophic cells of C.
sorokiniana occurs, which especially is correlated with nitrogen level in cells.
Keywords: microalgae; Chlorella sorokiniana; mixotrophic; photosynthesis.
64
4.2 INTRODUCTION
Mixotrophic growth of microalgae is gaining increasing interest, largely because it offers the
promise of an efficient strategy for commercial production of lipids and other value-added
products. The combination of organic carbon assimilation and simultaneous CO2 fixation in the
light leads to higher growth rates and biomass accumulation in algae cultured mixotrophically
than the same strains cultivated photoautotrophically or heterotrophically (Lee et al., 1996; Li et
al., 2014b). Mixotrophic growth also offers augment lipid productivity (Cheirsilp & Torpee,
2012), which is relevant for the biofuel production from microalgae.
Our previous study proved that the green alga Chlorella sorokiniana shows the best growth
performance in mixotrophic condition (Li et al., 2014b). In mixotrophic growth, CO2 and organic
carbon are assimilated simultaneously and, hence, both photosynthetic and organic substrate
metabolic pathways have to operate concurrently. The question is how to explain the mechanism
of mixotrophic growth, in particular how the photosynthetic and organic substrate metabolic
pathway interacts with each other. The photosynthetic pathways can be divided into the light
reactions, in which light energy is stored as ATP and NADPH, and the dark reactions, in which
the products of the light reactions are used to fuel cell metabolism and in particular reducing CO2
to three carbon sugars. For substrate metabolic pathways, glucose (as a model substrate) is
metabolized to three carbon sugars following ATP-dependent entry into the glycolysis pathway.
Thus, glucose metabolism may exert influences on utilization of carbon compounds arising from
CO2 fixation, and alter electron flux of photosynthesis by modulating the rate of ATP and
NADPH consumption (Kramer et al., 2004; Lee et al., 2012).
It has been reported that acetate inhibits photosynthesis in Chlamydomonas reinhardtii, and
high acetate virtually abolishes photosynthetic carbon gain (Endo & Asada, 1996; Heifetz et al.,
65
2000). Numerous groups have demonstrated that the light reactions and photosynthetic electron
transport of algal cells may be altered by exogenous organic carbon. Lewitus et al. (1991)
showed that glycerol assimilation by Pyrenomonas salina resulted in a reduction of
photosynthetic components associated with light-harvesting. The cell phycoerythrin content,
phycoerythrin to chlorophyll ratio, degree of thylakoid packing, number of thylakoids∙ cell-1,
and PSII particle size were also reduced (Lewitus et al., 1991). The addition of acetate to
photoautotrophically grown C. reinhardtii decreased PSII fluorescence and promoted a transition
from state I to state II, presumably with attendant adjustment of the antenna architecture of the
photosynthetic apparatus (Bulté et al., 1990). Similarly, mixotrophically cultured diatom
Phaeodactylum tricornutum supplemented with different kinds of organic substrate showed a
decline in O2 evolution, accompanied by decreased PSII activity and reduced electron transport
rate, which demonstrated that organic carbon metabolism can have an adverse impact on
photosynthesis beyond the green algae (Liu et al., 2009). However, glucose was shown to
enhance the net photosynthesis rate in the cyanobacterium Synechococcus sp., indicating that this
is not a universal situation (Kang et al., 2004).
Furthermore, exogenous organic substrate metabolism also affected the capacities of
photosynthetic enzymes. Previously studies have reported that organic carbon represses
expression of nuclear-encoded chloroplast proteins involved in light harvesting and inorganic
carbon fixation (Goldschmidt-Clermont, 1986; Kindle, 1987; Steinmuller & Zetsche, 1984). In
the acidophilic red alga Galdieria sulphuraria 074G, addition of glucose led to less abundant of
ribulose 1,5-bisphosphate carboxylase/oxygenase (Rubisco), and reduced levels of the PSII
reaction center protein D1 (Oesterhelt et al., 2007). Thus, there appear to be a number of
different ways in which photosynthetic activity is modulated during mixotrophic growth.
66
In this paper, we investigated the effect of glucose metabolism on photosynthesis and related
processes, and how microalgal cells respond under mixotrophic condition compared to
photoautotrophic condition, in particular, how the photosynthetic activity is regulated as the
algae grow mixotrophically. The effects of externally supplied glucose on multiple aspects of
photosynthesis were studied to assess mixotrophic capacities of C. sorokiniana at different
growth stages. Primary photosynthetic reactions were determined as light-dependent O2
evolution at photosystem (PS) II. Pulse-amplitude modulation (PAM) fluorescence
measurements as well as fluorescence spectroscopy at 77 K was employed to further characterize
the photosynthetic apparatus and energy transformation. Secondary (dark) reactions were
inferred from the total Rubisco carboxylase activities and the expression levels of two important
genes involved in Calvin Cycle: Rubisco large subunit (rbcL gene) and phosphoribulokinase
(PRK, prk gene). To the best of our knowledge, this is the first attempt to depict scenarios of
photosynthetic regulation for mixotrophic cells under different growth stages.
67
4.3 MATERIALS AND METHODS
4.3.1 Organism and medium
The green microalga Chlorella sorokiniana (UTEX 1602) was obtained from UTEX, the Culture
Collection of Algae at the University of Texas at Austin (USA). This strain was maintained at 4
o
C on an agar slant of Kuhl medium. In routine experiments, the inocula were cultured at 25 oC
in a 250-ml flask containing 150 ml Kuhl medium agitated with filter-sterilized air enriched with
CO2 to 1% (v/v). Illumination was provided by cool-white fluorescent lamps to give constant
light intensity of 100 µmol m─2 s─1. Except where indicated, the medium for photoautotrophic
culture was Kuhl medium, agitated with 1% CO2 in air (1% CO2-conditioned) or supplied with
only air (air-conditioned). For heterotrophic and mixotrophic culture the Kuhl medium was
supplemented with 6 g L─1 glucose and the cells were grown with orbital shaking.
4.3.2 Growth analysis
Dry cell weight (DCW) was determined according to our previous methods (Li et al., 2014b).
Nitrate-nitrogen was determined with the H2SO4-salicylic acid method, as described by Cataldo
et al. (1975) (Cataldo et al., 1975).
4.3.3 Photosynthetic oxygen evolution
Photosynthetic O2 evolution by intact cells was measured using a Clark-type oxygen electrode
(Hansatech, UK) using 1.2 mL of cell suspension in the oxygen electrode chamber. In order to
compare the relative photon yield of photosynthesis between the different samples, about the
same chlorophyll concentration (20 µM) was loaded in the oxygen electrode chamber.
Chlorophyll concentration was determined according to Porra et al. (2002) (Porra, 2002) after
extraction of whole cells with 100% methanol. To ensure that oxygen evolution was not limited
by the carbon source available to the cells, 20 µL of a 0.5 M sodium bicarbonate solution (pH 7.4)
68
was added prior to the oxygen evolution measurements. Dark respiration rate was measured with
the same cell suspension before each measurement of net photosynthetic rate. The parameters for
the photosynthetic responses to irradiance curves were analyzed according to Henley (1993)
(Henley, 1993):
𝑃
𝑃𝑚𝑎𝑥 (𝛼𝐼 ⁄√𝑃𝑚𝑎𝑥 + (𝛼𝐼) ) + 𝑅
𝐼𝑘
𝑃𝑚𝑎𝑥⁄𝛼
Where I represents irradiance, P the photosynthetic rate at irradiance I, Pmax the light-saturated
rate of photosynthesis, Ik the saturation light intensity, α-slope the slope of the light-limited part
of the P-I curve, and Rd the dark respiration rate.
4.3.4 Photosynthetic activity measurements
Photosynthetic activities were measured by PAM fluorometry (Walz, Effeltrich, Germany). Prior
to fluorescence measurements, chlorophylls a and b were extracted from cells and their
concentrations were calculated as reference (Porra, 2002). Algal cell suspensions were then
adjusted to a concentration of total 10 mM chlorophyll and dark-adapted for 15 min. Minimal
fluorescence yield (F0) was determined. The saturating light pulse was applied to the darkadapted samples to obtain the maximal fluorescence yield (Fm). The cells were then illuminated
with continuous actinic light for 5 min to obtain a steady fluorescence yield (Fs). A saturating
pulse was applied to obtain a stationary level of maximum fluorescence (Fmʹ). The fluorescence
parameters were calculated according to Baker (2008) (Baker, 2008).
69
4.3.5 77K fluorescence spectroscopy
Fluorescence spectra at 77K were measured to analyze stoichiometry and composition of PSI
and PSII. Low temperature fluorescence spectra were obtained as described by Kirchhoff et. al.
(2007) (Kirchhoff et al., 2007). Cells were adjusted to a chlorophyll concentration of about 2 μM,
shock-frozen in liquid nitrogen, and excited with a broad blue-green light source (400 –550 nm)
produced with a halogen lamp and Schott BG18, Corning 9782, and LOT heat mirror filters.
Emission spectra were recorded with a Horiba Jobin Yvon FluoroMax 4 spectrofluorimeter in
the spectral range of 650-800 nm (slit width, 2 nm) with an excitation wavelength of 435 nm (slit
width, 5 nm).
4.3.6 Measurements of rubisco activity
Rubisco is the initial enzyme of CO2 fixation in the Calvin-Benson cycle, catalyzing the
addition
of
CO2
to
ribulose-1,5-bisphosphate
(RuBP).
Microalgae
were
grown
photoautotrophically (1% CO2, air), heterotrophically, and mixotrophically. After 1 and 3 d, the
cells were harvested by centrifugation, frozen in liquid nitrogen, and subsequently grounded on
ice with a mortar and pestle in 1 ml of extraction buffer [50 mM Hepes, pH 7.8, 10 mM
dithiothreitol (DTT), 1% polyvinylpolypyrrolidone (PVPP), 1 mM ethylenediaminetetraacetic
acid (EDTA), 0.1% Triton] with 10 µL of protease inhibitor cocktail (Sigma, St. Louis, MO,
USA) and briefly centrifuged. The extract was incubated with 15 mM MgCl2 and 15 mM
NaHCO3 for 10 min at room temperature to activate enzymes. Total Rubisco activity was
determined by placing 20 µL of activated enzymes in 1 mL of assay buffer (100 mM EPPSNaOH pH 8.0, 20 mM MgCl2, 1mM EDTA, 0.2 mM NADH, coupling enzymes, 20 mM
NaHCO3) as described in Walker et al. (2013) (Walker et al., 2013). The reaction was initiated
with 0.5 mM ribulose bisphosphate (RuBP) and NADH consumption monitored.
70
4.3.7 Quantitative RT-PCR conditions and analysis
The total RNA was isolated using Trizol reagent (Ambion) from the cell pellet. One microgram
of total RNA was used as a template for each RT-reaction following the manufacturer’s
instructions kit (qScript cDNA SuperMix, Quanta Bioscinces). Gene-specific primers were
designed to amplify fragments of approximately 100–150 bp in length. For the quantification of
gene expression, qPCR was carried out on Mastercycle Realplex 2 (Eppendorf) using the
Perfecta Syber Green Fast Mix (Quanta Bioscinces, Gaithersburg, MD). The 18S ribosomal
RNA gene was served as internal control for the quantification assays. For gene expression
analysis by qPCR, the expression values were calculated according to the 2-∆∆CTmethod (for
each gene, ∆CT = CT,Gene − CT,housekeeping specific ). The ∆∆Ct calculation was validated
using the plot of the log cDNA dilution versus ∆Ct. See Table 4.1 for all primer sequences used
in this work.
Table 4.1 Primers for quantitative real-time PCR detection of expression genes in C. sorokiniana
Gene
18S
rbcL
prk
Primer sequence (5’→3’)
F: ATCAACCTGACAAGGCAACC
R: CCTGCGGCTTAATTTGACTC
F: TGGTCACCACCTGATAAACG
R: GCTCACTACTGCCGTGACAA
F: TGAATGGCTGCTGCTATGAC
R: GTTGAGGCATCTCCAATGCT
71
Length (bp) of production
150
132
150
4.4 RESULTS
4.4.1 Photoautotrophic, mixotrophic and heterotrophic cultivation of C. sorokiniana
Comparison of three different culture modes was carried out in 250-mL flasks. As shown in
Figure 4.1A, mixotrophy showed the most significant promotion in the cell growth, with the
specific growth rate and maximum biomass largely exceeded those of heterotrophy, 1% CO2conditioned photoautotrophy, and air-conditioned photoautotrophy (Table 4.2). After 3 d of
cultivation, nitrogen was assimilated completely in the medium for both mixotrophic and
heterotrophic cells, which went into a stationary phase; but it still remained in the
photoautotrophy, with 60% left in air-conditioned culture (Fig. 4.1B).
4.4.2 Comparison of photosynthesis and respiration for algal cells cultured mixotrophically
and photoautotrophically
The net photosynthetic O2 evolution rate as a function of the probing actinic light intensity (P-I
curve) of mixotrophic and photoautotrophic cultures was compared. In P-I curve, the rate of
photosynthesis first increased linearly with light intensity and then leveled off as the saturation
light intensity (Ik) was approached. The slope of the initial linear increase (α-slope) provides
information about the photon use efficiency of photosynthesis. The rate of photosynthesis
reached saturation at light intensities higher than Ik. This light-saturated rate (Pmax) provides a
measure of the capacity of photosynthesis for the algal sample (Melis et al., 1998).
72
A
Photoautotrophy (air)
Photoautotrophy (1%)
Mixotrophy
Heterotrophy
5
-1
DCW (g L )
4
3
2
1
0
0
1
2
3
4
Time (day)
B
KNO3 remaining in medium (%)
100
Photoautotrophy (air)
Photoautotrophy (1%)
Mixotrophy
Heterotrophy
80
60
40
20
0
0
1
2
3
4
Time (d)
Figure 4.1 Cultivation of C. sorokiniana under different conditions. (A) Growth curves of C.
sorokiniana under photoautotrophic, heterotrophic, and mixotrophic conditions; (B) Time course
of KNO3 consumption under photoautotrophic, heterotrophic, and mixotrophic conditions.
73
Table 4.2 Growth parameters of C. sorokiniana under different culture conditions
Specific growth
rate (d‒1)
Max. biomass
(g L‒1)
Mixotrophy
Heterotrophy
Photoautotrophy
(1% CO2)
Photoautotrophy
(air)
3.13 ±0.22
1.56 ±0.02
1.24 ±0.04
0.60 ±0.01
4.57 ±0.12
2.78 ±0.06
1.70 ±0.22
0.36 ±0.00
Values are the means ±SD
When C. sorokiniana was cultured mixotrophically, there was a drastic change in the level of
respiration and photosynthesis (Table 4.3). After cultivation for 1 d, mixotrophic cells exhibited
dramatic augment in respiration rate (Rd), which was 8- and 5-fold of 1% CO2-conditioned cells
and air-conditioned cells, respectively. At the same time, the photosynthetic O2 evolution rate of
mixotrophically grown cells was significantly higher than that of photoautotrophic cultures (Fig.
4.2). As shown in Table 4.3, mixotrophic cells reached the highest light-saturated rate of
photosynthesis (Pmax=~300 mmol O2 (mmol Chl‒1) h‒1) that is approximately 3 and 4 times
greater than that of 1% CO2-conditioned and air-conditioned cells, respectively. Moreover, the
values of Ik and α-slope were also much higher, pointing to high light requirement for obtaining
high photosynthetic rate and more efficient light utilization, respectively.
After 3 d, mixotrophic cells already entered stationary phase (Fig. 4.1), and exhibited
considerably decreased photosynthetic capability, with the lowest Pmax and Ik value (Fig. 4.2 ,
Table 4.3). At the same time, α-slope showed obvious decrease, which suggests that not all
chlorophyll molecules were photochemically competent in mixotrophic cells (Neidhardt et al.,
1998). Generally, 1% CO2-conditioned cells had higher photosynthetic O2 evolution rate than
air-conditioned cells, with higher values of Pmax, Ik and α-slope.
74
Table 4.3 Characteristic P-I curve parameters (Rd, Pmax, Ik and α-slope) of C. sorokiniana
cultured under mixotrophic (M) and photoautotrophic (1% CO2, air) conditions after 1 and 3 d.
Time
(d)
1
3
Mode
Rd
Pmax
Ik
α-slope
R2
M
1% CO2
Air
M
1% CO2
Air
62.00 ±2.21
7.59 ±0.39
12.06 ±0.45
35.65 ±0.64
13.71 ±0.63
12.47 ±0.40
282.8 ±8.85
99.02 ±9.11
68.49 ±1.91
27.93 ±8.14
191.5 ±8.49
49.05 ±3.51
310.8 ±4.55
183.6 ±19.65
130.1 ±7.30
49.03 ±7.78
250.1 ±28.90
149.2 ±18.03
0.91 ±0.03
0.54 ±0.00
0.53 ±0.04
0.56 ±0.00
0.77 ±0.10
0.33 ±0.05
0.99
0.99
1
0.93
0.99
0.99
Values are the means ±SE derived from the P-I curve.
Pmax, the light-saturated rate of photosynthesis, mmol O2 mmol Chl‒1 h‒1;
Rd, respiration rate, mmol O2 (mmol Chl)‒1 h‒1;
Ik, saturation light intensity, µmol photons m─2 s─1;
α-slope, the slope of the initial linear, (mmol O2 (mmol Chl)‒1 h‒1) (µmol m─2 s─1) ─1.
4.4.3 Fluorescence characterization of mixotrophic and photoautotrophic cells
Light-dependent O2 evolution is strictly dependent on PSII-associated water cleavage and
concomitant linear photosynthetic electron flow. To characterize photosynthetic activity of
mixotrophic and photoautotrophic cells, utilization of absorbed light energy was examined in
more detail via measurement of chlorophyll fluorescence. Potential maximum quantum
efficiency (Fv/Fm), quantum yield of PSII photochemistry (ΦPSII), non-photochemical quenching
(NPQ), energy-dependent quenching (qE), photoinhibitory quenching (qI), and the fraction of
open PSII reaction centers (qL) were determined (Fig. 4.3).
After cultivation for 1 d, Fv/Fm of mixotrophic cells was 0.81-0.86 (Fig. 4.3A), which
indicated healthy cells and efficient PSII primary activity (from water to QA). Moreover,
mixotrophic cells showed no significant differences with respect to 1% CO2-conditioned cells
75
either in ΦPSII or in qL (Fig. 4.3B, 4.3F), thus indicating no limitations in electron transport
downstream of PSII.
250
200
150
2
Oxygen evolution
(mmol O (mmol Chl)-1 h-1)
300
100
50
0
0
200
400
600
800
Light intensity (mol photons m-2 s-1)
Figure 4.2 Light-saturation curves of photosynthesis in mixotrophically (circle) and
photoautotrophically (triangle up, 1% CO2; triangle down, air) cultured C. sorokiniana after 1
(filled) and 3 (open) days, respectively. Rates of oxygen evolution were measured on a per
chlorophyll basis.
After cultivation for 3 d, Fv/Fm of mixotrophic cells dramatically leveled off by nearly 60%,
but there was no difference for 1% CO2-conditioned cells (0.85-0.88) and air-conditioned cells
(0.75-0.78) compared with that after cultivation for 1 d (Fig. 4.3A, 4.3G). The ΦPSII of
mixotrophic cells also largely declined, and showed lower value than that of 1% CO2conditioned cells and air-conditioned cells (Fig. 4.3B, 4.3H), which suggested that less of the
absorbed light energy was directed to functional PSII centers.
76
1.0
0.9
A
G
B
H
C
C
FI
D
J0
E
K
Fv/Fm
0.8
0.7
0.6
0.5
0.4
0.3
0.8
PSII
0.6
0.4
0.2
1.0
0.8
NPQ
0.6
0.4
0.2
0.0
-0.2
0.6
200
400
600
800
600
800
qE
0.4
0.2
0.0
-0.2
0.3
qI
0.2
0.1
0.0
-0.1
-0.2
1.0
F
L
qL
0.8
0.6
0.4
0.2
0.0
0
200
400
600
800
0
200
Light intensity (mol photons m-2 s-1)
77
400
Figure 4.3 Photosynthetic characteristics of C. sorokiniana grown mixotrophically (red square)
and photoautotrophically (black square, 1% CO2; grey square, air) after 1 d (A, B, C, D, E, F)
and 3 d (G, H, I, J, K, L). The light dependency of each photosynthetic parameter is shown (A-L).
(A, G) The potential maximum quantum efficiency (Fv/Fm); (B, H) the quantum yield of PSII
(ΦPSII); (C, I) the non-photochemical quenching (NPQ); (D, J) the energy-dependent quenching
(qE); (E, K) the photoinhibitory quenching (qI); and (F, L) the fraction of open PSII reaction
centers (qL).
On the other hand, the qL of mixotrophic cells tended to be higher than that of
photoautotrophic cells, although it was lower than 1 d- cultured mixotrophic cells at low
irradiance (Fig. 4.3F, 4.3L).
NPQ, a measure of non-photochemical quenching processes (Muller et al., 2001), became
apparent mainly in 1% CO2-conditioned cells and air-conditioned cells, especially for the latter
cultured for 3 d. The air-conditioned cells cultured for 3 d exhibited increased levels of NPQ
across the entire irradiance range. The high-energy quenching qE, component of NPQ shows up
significantly for 1% CO2-conditioned cells. Under these conditions also a slight increase in the qI
parameter occurs that is indicative for photodamage. However, mixotrophic cells cultured for 1
and 3 d displayed almost no NPQ and qE. This indicates no obvious photoprotective regulation,
unlike the appearance of the light-harvesting complex stress-related protein 3 (LHCSR3) seen in
the green alga C. reinhardtii under stressful light conditions (Peers et al., 2009).
4.4.4 ΦPSII vs. qE and ΦPSII vs. qL
To further characterize the influence factor of ΦPSII, we analyzed the dependence of ΦPSII on qE
and on the redox level of the PQ pool (qL). For mixotrophic cells, there was at most a small
78
amount of qE in the range of ΦPSII, and the two factors were not significantly correlated. ΦPSII
values of mixotrophic cells after cultivation for 3 d were distributed in the lower range,
compared with the distribution after 1 d (Fig. 4.4A, 4.4B). On the other side, photoautotrophic
cells generally showed a direct correlation of ΦPSII on qE.
0.9
0.8
B
A
0.7
PSII
0.6
0.5
0.4
0.3
0.2
0.1
0.0
-0.1
0.0
0.1
0.2
0.3
0.4
0.5
-0.1
0.0
0.1
0.2
0.3
0.4
0.5
qE
C
0.8
D
PSII
0.6
0.4
0.2
0.0
0.0
0.2
0.4
0.6
0.8
1.0 0.0
0.2
0.4
0.6
0.8
1.0
qL
Figure 4.4 ΦPSII vs. qE (A, B) and ΦPSII vs. qL (C, D) of C. sorokiniana grown mixotrophically
(red square) and photoautotrophically (black square, 1% CO2; grey square, air) after 1 day (A, C)
and 3 days (B, D).
79
We also studied the dependency of ΦPSII on the redox level of the PQ pool, which are
generally directly correlated. Under all cultivation conditions, ΦPSII showed the expected positive
relationship with qL, which indicates PSII can convert light energy more efficiently with a more
oxidized PQ pool. After cultivation for 1 d, mixotrophic and 1% CO2-conditioned cells had
similar trends of dependency of ΦPSII on qL (Fig. 4.4C). However, in mixotrophic cells after
cultivation for 3 d, the decline in ΦPSII was less pronounced in response to qL. Thus, compared
with 1% CO2-conditioned cells and air-conditioned cells, ΦPSII in mixotrophic cells is less
sensitive to qL, indicating that PSII is controlled by other processes.
4.4.5 77K fluorescence spectra
The above results of photosynthetic regulation prompted us to analyze PSI and PSII with
associated antenna in mixotrophic and photoautotrophic cells. It has been widely accepted that
77K fluorescence spectra indicates the energy flow between the two photosystems. Our results
show that the spectra of the mixotrophic and photoautotrophic cells had major emission peaks
around 688 nm (F688) and 722 nm (F722), shoulder peaks around 695 nm (F695) (Fig. 4.5).
Among these, F688 and F695 are emitted from PSII, F722 are from PSI (Morgan-Kiss et al.,
2002). The fluorescence ratio of F722 to F688 was used to examine variations in the
redistribution of excitation energy between PSI and PSII. This value was 0.97 for 1 dmixotrophic cells, which was much less than that of 1% CO2-conditioned cells (1.63) and airconditioned cells (1.32). The low F722 to F688 ratio for mixotrophic cells indicates that the PSI
antenna system is enlarged relative to the PSII antenna that correlates with transition to state 2.
For 3 d-mixotrophic cells, the fluorescence peak at 695 nm considerably decreased and the peak
at 688 nm shifted to 685 nm. The observed shift is a sign of increased free LHCII component,
80
which has a characteristic emission around 680 nm (Haferkamp et al., 2010), and indicates that
more LHCII is not functionally coupled to PSII or PSI.
A
77K Fluorescence emission (norm.)
1.5
1.0
0.5
0.0
B
1.5
1.0
0.5
0.0
660
680
700
720
740
760
780
Wavelength (nm)
Figure 4.5 77K fluorescence emission spectra of mixotrophic (solid line) and photoautotrophic
(dotted line, 1% CO2; dash line, air) cells after excitation of chlorophylls at 435 nm. Spectra are
normalized to the main PSII emission peak A, 1 day; B, 3 days.
81
4.4.6 Adjustment of Rubisco and PRK
In the photosynthesis process, photosynthetic rate in mixotrophic cells was significantly
enhanced after cultivation for 1 d, and then declined after 3 d, compared with those in
photoautotrophic cells. For efficient utilization of the chemical energy generated from the light
reactions, the corresponding dark reactions must be coordinated. Enzyme activities and
expression levels of mixotrophic and photoautotrophic cells involved in Calvin-Benson cycle
were compared in order to analyze whether the regulation of photosynthesis was accompanied by
major changes on the protein levels. We also included cells cultivated in the heterotrophic
condition in these analyses.
After cultivation for 1 d, total Rubisco activity of the mixotrophic cells was the highest, yet it
declined by 87% and became the lowest one after 3 d (Table 4.4). Interestingly, Rubisco activity
of heterotrophic cells cultured for 1 d was comparable with 1% CO2-conditioned cells. After 3 d,
there still remained 46% of Rubisco activity in heterotrophic cells, yet for 1% CO2-conditioned
cells it was only 30%.
Table 4.4 Effects of different growth conditions on total Rubisco carboxylase activity and
chlorophyll to protein ratio.
Culture condition
Mixotrophy
Photoautotrophy
(1% CO2)
Photoautotrophy
(air)
Heterotrophy
Rubisco carboxylase activity
(µmol CO2·[mg chlorophyll]‒1·s‒1 )
1d
3d
48.02 ±3.62
6.40 ±0.24
41.12 ±1.61
28.66 ±0.19
Chlorophyll per protein
(0.01 mg mg‒1)
1d
3d
5.70 ±0.46 6.81 ±0.25
7.45 ±0.30 8.94 ±0.06
15.76 ±0.03
17.91 ±0.03
8.75 ±0.02
8.07 ±0.01
40.88 ±1.83
22.20 ±0.79
3.49 ±0.16
3.32 ±0.11
Values are the means ±SD
82
The expression levels of Rubisco and phosphoribulokinase (PRK) were examined by
quantitative RT-PCR (Fig. 4.6). Consistent with the result of total carboxylase activity, Rubisco
expression in mixotrophic cells cultured for 1 d was similar to 1% CO2-conditioned cells, but
lower than that of heterotrophic cells. On the other hand, PRK expression level in mixotrophic
cells was the highest, which was approximately 2 times of those in photoautotrophic cells.
Impressively, air-conditioned cells cultured for 1 d exhibited dramatically high value of Rubisco
expression level, indicating that the rbcL gene was obviously up-regulated in this condition.
16
A
Relative mRNA levels
14
12
Mixotrophy
Heterotrophy
Photoautotrophy (air)
Photoautotrophy (1%)
10
8
6
4
2
0
B
Relative mRNA levels
1.0
0.8
0.6
0.4
0.2
0.0
1
3
Time (day)
Figure 4.6 RT-PCR results of Rubisco (A) and phosphoribulokinase (B) involved in the Calvin
cycle.
83
4.5 DISCUSSION
In this study, we comparatively analyzed the response of photosynthetic characteristics of
mixotrophic cells at different growth stages. Compared with photoautotrophic cells, the
mixotrophically grown C. sorokiniana cells exhibited an increase in respiratory metabolism, as
indicated by the large respiratory O2 consumption rates (Table 4.3). This result was in agreement
with glucose-grown C. vulgaris UAM 101 cells (Villarejo et al., 1995), fructose-grown
Anabaena variabilis cells (Valiente et al., 1992), acetate-grown Chlamydomonas reinhardtii
cells (Fett & Coleman, 1994), and Phaeodactylum tricornutum cells supplied with several
organic carbons (Liu et al., 2009).
Meanwhile, the greatest photosynthetic activity occurred in mixotrophic cells after cultivation
for 1 d, which was corroborated by the highest photosynthetic O2 evolution rate. Additionally,
the result of 77K fluorescence spectra was consistent with the results of photosynthetic O2
evolution rate. It has been widely accepted that 77K fluorescence spectra indicates the energy
flow between the PSI and PSII photosystems (Fuhrmann et al., 2009; Lin & Knox, 1991). In
mixotrophic cells after cultivation for 1 d, the ratio of PSI fluorescence to PSII fluorescence was
reduced. These results proved that PSII activity increased in mixotrophic cells, and glucose
enhanced the photosynthetic efficiency. The up-regulation of photosynthetic activity was also
observed in the dark reaction. The increase in photosynthetic efficiency was associated with a
higher Rubisco carboxylase activity (Table 4.4). At the same time, compared with those of
photoautotrophically 1% CO2-conditioned cells, a similar expression level of Rubisco and a
higher expression level of PRK also support the high photosynthetic rates exhibited by
mixotrophic cells.
84
It should be noted that photoautotrophically air-conditioned cells cultured for 1 d showed the
lowest total Rubisco activity (Table 4.4). This is partially due to the relative increase in the ratio
of chlorophyll to protein. It is known that Rubisco exists in two states: activated and inactivated.
Before catalyzing carbon assimilation, the enzyme must be activated via the carbamylation
(Lorimer, 1981). Several phosphorylated metabolites , such as Rubp, 2-carboxyarabinitol 1phosphate or 6-phosphogluconate can strongly bind to carbamylated sites, leading to inactive
Rubisco in the cell (Mouget et al., 1993; Nieva & Valiente, 1996). Thus, the explanation for the
low total Rubisco activity could also concern on the Rubisco regulation, which leads to the
difficulty of achieving complete activation of Rubisco during measurement. Interestingly,
heterotrophic cells displayed higher Rubisco carboxylase activity, and higher expression levels
of Rubisco and PRK. Therefore, the algal cells probably maintain the carbon-fixing capacity in
heterotrophic condition for a period after inoculating photoautotrophic seeds. Similar
observations is also reported in microalga Chlorella protothecoides that Rubisco carboxylase
activity gradually increased after the photosynthetic algal cells transferred to heterotrophic
conditions (Xiong et al., 2010).
Based on current knowledge about the process of photosynthesis adjustment in algal cells,
photosynthetic activity may be down-regulated by exogenous carbon sources (Bultéet al., 1990;
Endo & Asada, 1996; Heifetz et al., 2000; Lewitus et al., 1991; Liu et al., 2009), although this
condition generally increases specific growth rate and biomass concentration (Heredia-Arroyo et
al., 2011; Liang et al., 2009). Photoautotrophically grown algae often have decreased
photosynthetic capacity and reduced Rubisco carboxylase activity after supplementing with
glucose (Liu et al., 2009; Villarejo et al., 1995). Also, in these cases, genes related to
85
photosynthesis were down-regulated, including those for chlorophyll-binding proteins and
Rubisco.
In our study, enhanced photosynthetic capacity was observed in mixotrophic cells cultivated
for 1 d. Instead, mixotrophic cells showed largely decreased photosynthetic efficiency after
cultivation for 3 d. Lowered ΦPSII and Fv/Fm are indicators of a reduced photochemical efficiency
of PSII, which are in agreement with the results of dramatically decreased photosynthetic O2
evolution rate. This conclusion was further supported by the 77K fluorescence spectra. The shift
of F688 to F 685 as well as the change of the relative intensity of F695 point out changes in the
organization of the components of PSII complex. It strongly suggests that the light energy
absorbed by LHCII is not properly transferred to the PSII reaction center. The major difference
of photosynthetic regulation strategy between mixotrophic cells cultured for 1 d and 3 d is
probably correlated with the different physiological state, especially from the aspect of nutrient
availability. After 1 d, the mixotrophic cells grew in optimal nutrient condition; after 3 d, the
acceleration of growth and increase of biomass changed the nutrient status in the cells, especially
for nitrogen, which was completely assimilated from the medium (Fig. 4.1). It’s been proved that
in algae nitrogen assimilation is directly coupled to photosynthetic electron flow and 20% of the
photosynthetic electron flow would be in support of nitrogen assimilation for amino acids
synthesis (Turpin, 1991). In Chlamydomonas at low light intensities, up to 40% of
photogenerated reductant is allocated to nitrate reduction (Curtis & Megard, 1987). As growth
becomes limited by nitrogen, there are large changes in cellular metabolic ATP and NADPH
demands in algal cells. Thus, the down-regulation of photosynthesis in mixotrophic cells
probably serves as a feedback control to guarantee growth and survival of algal cells.
86
In plant leaves after long-term growth at elevated CO2, down-regulation of the photosynthetic
machinery also occurs and is called “sugar signalling” (Paul & Pellny, 2003). Glucose levels are
often high in plants grown at elevated CO2, whereas the expression of genes for photosynthesis is
reduced (Oosten & Besford, 1996). However, mechanistic details of sugar repression of
photosynthesis is not available, due to the complexity, multiplicity and cross-talk with hormone
signalling and nitrogen metabolism (Paul & Foyer, 2001). In sugar feeding experiments or at
elevated CO2 giving rise to high leaf carbohydrate, no loss of photosynthetic gene expression or
photosynthetic capacity is observed where nitrogen is kept high (Geiger et al., 1999; Martin et al.,
2002). Photosynthetic capacity can actually be stimulated by high CO2 under high nitrogen,
however, it is reduced in nitrogen-limited plants (Habash et al., 1995).
It seems that C. sorokiniana cells follow the same photosynthesis regulatory patterns. Here we
further tested this regulation strategy through fluorescence characterization of algal cells after
cultivation under nitrogen-deplete or nitrogen-replete conditions (Table 4.5). Compared with
control, values of ΦPSII, Fv/Fm and qL in nitrogen-deplete condition were clearly reduced with
glucose supplementation; however, they were not significantly influenced without glucose. On
the contrary, they were largely enhanced in nitrogen-replete conditions, even with glucose
addition. Therefore, sugar-signalling appears to be operating under nitrogen-depleted condition;
whereas under nitrogen-replete condition, no down-regulation of photosynthesis is observed.
Collectively, carbon to nitrogen balance rather than purely high carbon level is central to
feedback control of photosynthesis.
87
4.6 CONCLUSION
By systematically analyzing photosynthetic regulation in mixotrophic cells of C. sorokiniana,
we showed that photosynthesis is dynamically adjusted at different growth stages. Additionally,
we provided quantitative evidence that nitrogen limitation is correlated with down-regulation of
photosynthesis in mixotrophic cells. Although elucidation of the precise mechanisms that give
rise to feedback control of photosynthesis in algae still remains sketchy, our study results would
provide a clue on how to keep high photosynthesis efficiency for carbon acquisition and biomass
accumulation, whether through nutrient supply schemes or through molecular engineering.
88
Table 4.5 Chlorophyll fluorescence characteristics of algal cells.
Parameter
Control
Fv/Fm
ΦPSII
qL
0.77 ±0.01
0.57 ±0.01
0.49 ±0.00
+N
Glucose content (g L‒1)
0
6
20
0.81 ±0.01 0.83 ±0.02 0.84 ±0.02
0.72 ±0.02 0.74 ±0.00 0.72 ±0.01
0.67 ±0.02 0.66 ±0.01 0.76 ±0.01
‒N
Glucose content (g L‒1)
0
6
20
0.74 ±0.01 0.73 ±0.01
0.72 ±0.02
0.62 ±0.02 0.48 ±0.01
0.45 ±0.00
0.52 ±0.01 0.34 ±0.00
0.34 ±0.00
Values are the means ±SD
89
Algal cells were grown photoautotrophically in Kuhl medium under light intensity of 100 µmol m─2 s─1, agitated with 1%-CO2.
When grown to the exponential phase, algal cells were utilized as control and inoculated into medium with (+ N) or without (‒ N)
1 g L‒1 KNO3, supplemented with 0, 6, and 20 g L‒1 glucose, respectively. After inoculation, cells were grown for 9 hrs with
orbital shaking, under light intensity of 100 µmol m─2 s─1. For fluorescence measurement, the actinic light intensity was 100 µmol
m─2 s─1.
CHAPTER FIVE
REGULATORY MECHANISM OF STARCH AND LIPIDS ACCUMULATION IN A
MICROALGA STRAIN CHLORELLA SOROKINIANA
5.1 ABSTRACT
Microalgae have attracted growing attention due to their potential in biofuel feedstock
production. However, current understanding of the regulatory mechanisms for lipid biosynthesis
and storage in microalgae is still limited. This work explores the metabolic changes of starch and
lipid biosynthesis in the microalga Chlorella sorokiniana. The organism demonstrated faster
growth, higher maximum biomass dry weight (DW), and higher starch and lipid content under
mixotrophic condition compared to the heterotrophic condition. Time-course analysis revealed
that both the mixotrophic and heterotrophic cells showed sequential accumulation of starch and
lipids. That is, when nitrogen was replete and/or depleted over a short period, starch was the
predominant carbon storage form with basal levels of lipid accumulation. After a period of
nitrogen depletion, lipid accumulation increased considerably, accompanied by starch
degradation. Results show that lipid accumulation in mixotrophically-grown cells under nitrogen
starvation is strongly dependent on the linear electron flow of photosynthesis, peaking at lower
light intensities. Lipid accumulation was also partially due to starch degradation in C.
sorokiniana, as well as the turnover of primary metabolites. Collectively, our results reveal an
interesting pattern of starch and lipid accumulation that is basically controlled by nitrogen levels.
The mixotrophic growth of C. sorokiniana shows promise for biofuel production in terms of
lipid accumulation in the final biomass.
Keywords: microalgae; Chlorella sorokiniana; starch; lipids.
90
5.2 INTRODUCTION
Microalgae offer an alternative feedstock for biofuel production due to their high growth rates,
high photosynthetic efficiency, and relatively high content of energy-rich molecules. Starch and
lipids are two major carbon and energy storage compounds in microalgal cells, particularly under
conditions of stress or nutrient starvation. If biofuel feedstock is to be used at a refinery to make
diesel, jet fuel or gasoline, lipids are a preferable option due to their higher energy density and
lower downstream energy cost from reduced hydrogen requirements (Sivakumar et al., 2010).
Therefore, enhancing lipid production in algae has garnered considerable attention.
Ways to increase lipid production have been explored through genetic and metabolic
engineering of microalgae and plants. Most efforts have focused on stimulating fatty acid
biosynthesis (Dehesh et al., 2001; Dunahay et al., 1995; Klaus et al., 2004), increasing the
availability of glycerol as a backbone of lipids (Vigeolas et al., 2007), and overexpressing the
genes involved in the triacylglycerol (TAG) biosynthesis pathway (Jako et al., 2001; Lardizabal
et al., 2008; Zheng et al., 2008). However, these efforts have shown limited success, typically
yielding less than a 50% increase in lipid content. Another approach to increasing cellular lipid
content is to block starch synthesis pathways. Several studies have reported the
overaccumulation of lipid bodies in starch-devoid mutants of Chlamydomonas reinhardtii (Li et
al., 2010a; Li et al., 2010b; Wang et al., 2009; Work et al., 2010). However, impairing starch
synthesis does not necessarily result in higher lipid content in mutants of C. reinhardtii and
Arabidopsis (Periappuram et al., 2000; Siaut et al., 2011). Therefore, the mechanisms involved in
rerouting carbon metabolism from carbohydrates to fatty acids are still unknown.
Many studies have demonstrated that a variety of stress conditions such as excessive light,
sulfur deprivation and nitrogen deprivation can induce the storage of carbohydrates and/or oil
91
accumulation in microalgae (Dragone et al., 2011; Sharma et al., 2012). The microalgae
Chlorella species are known to accumulate starch and lipids (Mizuno et al., 2013; Takeshita et
al., 2014). Our previous work found that the lipid levels of Chlorella sorokiniana under
mixotrophic conditions are 2- to 3-fold higher than those under heterotrophic conditions,
provided that the same amount of organic carbon is supplied (Li et al., 2014a). Starch synthesis
shares common carbon precursors with lipid synthesis. However, it is still unclear whether starch
or lipids are more likely to be accumulated, and how this accumulation is regulated. More
research on carbon partitioning of algal cells into lipids and/or starch would provide key insights
for cultivation strategies and development for biofuel production.
This study provides a greater understanding of carbon allocation between starch and lipids and
of the regulatory mechanisms controlling this distribution in C. sorokiniana in response to
altered growth environments. In addition to analyzing the time course of starch and lipid content,
quantitative real time (RT)-PCR was used to examine the expression levels of four important
genes involved in starch and lipid metabolism: acetyl-CoA carboxylase (ACCase) beta subunit
(accD gene), ADP-glucose pyrophosphorylase (AGPase) large subunit (AGP-L gene), β-amylase
(BAM gene), and iso-amylase (ISA1 gene). ACCase catalyzes the irreversible conversion of
acetyl-CoA to malonyl-CoA in fatty-acid synthesis (Cronan Jr & Waldrop, 2002); AGPase
catalyzes the rate-limiting step of starch synthesis from glucose 1-phosphate and ATP to ADPglucose and pyrophosphate (Ball & Morell, 2003); β-amylase and iso-amylase play critical roles
in the hydrolysis of the starch granules that hydrolyze the 1,4- and 1,6-glycosidic bonds,
respectively (Zeeman et al., 2010a). This study also included an analysis of metabolites in order
to profile the major metabolite accumulation patterns.
92
Results reveal a nitrogen-level controlled temporal relationship between starch and lipids, with
starch accumulated in the nitrogen (N)-replete condition and lipid accumulated in the N-depleted
condition. Accumulation of both starch and lipids in mixotrophically-grown cells was found to
be strongly dependent on the linear electron flow of photosynthesis, while lipid content peaked at
lower light intensities. Understanding the interdependence of starch and lipids as well as their
regulation is integral to the metabolic engineering of alga C. sorokiniana for improved biofuel
productivity. Results also suggest that converting previously assimilated carbon in the form of
starch and other components may provide carbon/energy for lipid biosynthesis. Our results
demonstrate that mixotrophically-grown C. sorokiniana is a good candidate for lipid-based
biofuel feedstock production.
93
5.3 MATERIALS AND METHODS
5.3.1 Organism and culture conditions
The green microalga Chlorella sorokiniana (UTEX 1602) was obtained from UTEX, the Culture
Collection of Algae at the University of Texas at Austin (USA). This strain was maintained at 4
o
C on an agar slant of the Kuhl medium (KUHL & LORENZEN, 1964). In routine experiments,
the inocula were cultured at 25 oC in a 250-mL flask containing 150 ml Kuhl medium, agitated
with filter-sterilized air enriched with CO2 to 1% (v/v). Cool-white fluorescent lamps gave a
constant light intensity of 100 µmol m─2 s─1. In experimental cultures, 2% (v/v) exponentially
growing seeds were inoculated into each flask, which contained 50-mL Kuhl medium
supplemented with glucose and grew at 25 oC on an orbital shaker at 150 rpm. For mixotrophic
cultures, illumination was provided; for heterotrophic cultures, flasks were covered with foil.
For treatment with the PSII inhibitor 3-(3,4-dichlorophenyl)-1,1-dimethylurea (DCMU), the
compound (in methanol) was added directly to the growth media. Control cells were treated with
methanol alone. To inhibit de novo protein synthesis, cycloheximide (CHX) was added at the
beginning of the experiments. The final concentration in the mineral medium was 1 mg L‒1.
CHX and DCMU were purchased from Sigma-Aldrich.
5.3.2 Analytical procedure
To determine the biomass dry weight (DW), a 5 mL cell suspension sample was centrifuged at a
speed of 1000 ×g for 5 min. Cell pellets were washed twice with distilled water and dried in a
pre-weighed aluminum dish at 105 °C for 3 hrs. They were subsequently cooled to room
temperature in a desiccator before weighing.
Glucose concentrations were analyzed by a Dionex ICS-3000 ion chromatography system
(Dionex Corporation) equipped with a CarboPac TM PA 20 (4×50mm) analytical column and
94
CarboPac TM PA 20 (3×30mm) guard column. Samples were filtered with 0.2μm pore-size filter
before injection and eluted with 0.01 M NaOH at a flow rate of 0.500 ml min‒1. The analytes
were detected and quantified against standard curves by electrochemical detection in a pulsed
amperometric detector.
Nitrate-nitrogen was determined with the H2SO4-salicylic acid method, as described by
Cataldo et al. (1975) (Cataldo et al., 1975).
The lipids were analyzed with gas chromatography using tridecanoic acid (C13:0) as an
internal
standard.
The
corresponding
chromatographic
conditions
and
the
extraction/transesterification method are described in O'Fallon, et al. (2007) (O'Fallon et al.,
2007).
For starch analysis, samples of 1‒2 mL of the algal suspension were harvested by centrifuging
at 3000 g for 5 min. Algal pastes were disintegrated by vortexing with 0.5 mL of glass beads (0.5
mm diameter) in 1 mL of deionized water. The pigments were extracted three times with 5 mL of
80% ethanol for 5 min at 80 oC. After centrifuging, concentrations of starch in the pellets were
measured enzymatically with the Sigma Starch Assay Kit (STA 20, Sigma-Aldrich, St. Louis,
MO, USA) according to the manufacturer’s instructions.
For microscopy analyses, C. sorokiniana cells were stained with a Nile red fluorescent dye
(Sigma-Aldrich) at a final concentration of 1 µg mL‒1 for 10 min in the dark, and then observed
under a Zeiss fluorescence microscope (Carl Zeiss, Jena, Germany).
5.3.3 Quantitative RT-PCR conditions and analysis
Total RNA was isolated with a Trizol reagent (Ambion) from the cell pellet. One microgram of
total RNA was used as a template for each RT-reaction, following manufacturer’s instructions
(qScript cDNA SuperMix, Quanta Bioscinces). Gene-specific primers were designed to amplify
95
fragments of approximately 100–150 bp in length. For the quantification of gene expression,
qPCR was carried out on a Mastercycle Realplex 2 (Eppendorf) using the Perfecta Syber Green
Fast Mix (Quanta Bioscinces, Gaithersburg, MD). The 18S gene served as the internal control
for quantification assays. For gene expression analysis by qPCR, the expression values were
calculated according to the 2-∆∆CT method (for each gene, ∆CT = CT, Gene − CT,
housekeeping specific). The ∆∆Ct calculation was validated using the plot of the log cDNA
dilution versus ∆Ct. See Table 5.1 for all primer sequences used in this work.
Table 5.1 Primers for quantitative real-time PCR detection of expression genes in C. sorokiniana
Gene
accD
AGP-L
BAM
ISA1
18S
Primer sequence (5’→3’)
F: ACGCCACTCAAGAAGGCTTA
R: GAAATTTTTGCCATTTGCATT
F: CCATGAGCAACTGCATCAAC
R: GGTTGAGCGAGGTGGAGTT
F: GTACCCGTCCTACCCAGAGG
R: TGTCGTAGCACTGGAACTGG
F: CCACCGCCTCTGTCAACT
R: CGTTGGCCTCATTGTGCT
F: ATCAACCTGACAAGGCAACC
R: CCTGCGGCTTAATTTGACTC
Length (bp) of production
100
74
72
100
150
5.3.4 Metabolite analysis
Relative levels of amino acids, organic acids and sugars were determined by gas
chromatography- mass spectroscopy (GC-MS). The lyophilized cells were transferred to a 2 ml
eppendorf tube and disrupted with a steel ball (5mm). Then, 0.75 ml of extraction buffer
methanol: chloroform: water (5:2:2) was added to each sample and vortexed for 10 minutes at
room temperature. Cell debris was centrifuged at 16,000 g for 2 minutes. The supernatant was
evaporated and spiked with 2 µL of internal standard mixture solution of methyl esters and C30
96
linear chain-length fatty acids.
Ninety minutes of incubation (5µl pyridine methoxyamine
hydrochloride, added to each sample) was performed in a thermostatic bath at 30°C. Samples
were derivatized by adding 45 µl N-methyl-N-trimethylsilyltrifluoroacetamide (MSTFA+
1%TMCS, Pierce, Rockford, IL, USA) for 30 minutes at 37°C. Time of flight mass spectrometer
(Pegasus 4D, LECO, St. Joseph, MI),
equipped with an MPS-2 autosampler (Gerstel,
Muehlheim, Germany), was used for analysis with an RTX®-5Sil MS with Integra-Guard®
column (30 m × 0.25 mm ID × 0.25 μM film thickness) from Restek (GmbH, Bad Homburg,
Germa ny). The column was held isothermally at 50 °C for 1 min and ramped to at 20 °C min‒1
to 330 °C over 5 min. All injections were performed in splitless mode (1 µL), with helium as a
carrier gas at a constant flow of 1 mL min‒1. The ChromaTOF software version 4.41 equipped
with the LECO/Fiehn Metabolomics database was used for primary metabolite data processing.
97
5.4 RESULTS
5.4.1 Sequential accumulation of starch and lipids
To gain insight into the relationship between starch and lipid accumulation, we monitored the
time course of changes in starch and lipids as a result of mixotrophic and heterotrophic growth.
Following inoculation, starch levels in mixotrophic cells increased rapidly during exponential
growth, with the maximum content of 27% after 2 d, which then steadily decreased (Fig. 5.1A).
The accumulation of lipids lagged behind the starch, with a rapid increase observed after 3d of
cultivation, when starch accumulation declined. The lipid content continued to increase to 44%
by the end of the time course. The increased lipid accumulation was visualized using Nile red
staining after cultivation for 3 d (Fig. 5.1E) and 6 d (Fig. 5.1F). However, there was no oil
droplet formation in algal cells after 1 d (Fig. 5.1D). This finding supports the observation for a
delay in the accumulation of lipids. Similarly, in heterotrophic growth, the maximum
accumulation of starch occurred after 2 d of cultivation, and a marked loss of starch was
followed by enhanced lipid production (Fig. 5.1B). However, the maximum starch and lipid
content during the time course were 14% and 19%, respectively, which was much less than that
observed in the mixotrophic culture.
We also quantified the remaining glucose and nitrate levels in the medium under mixotrophic
and heterotrophic conditions. The assimilation of nitrate occurred much faster than that of
glucose, which was 1 d for the mixotrophic culture and 2 d for the heterotrophic culture (Fig.
5.1C). However, glucose was used up in the medium after 3 d in the heterotrophic culture and 4
d in the mixotrophic culture. Therefore, this situation countered that observed in nitrate
assimilation.
98
Figure 5.1 Mixotrophic and heterotrophic cultivation of C. sorokiniana. (A) Time course of DW,
lipid, and starch in mixotrophic culture; (B) Time course of DW, lipid, and starch in
heterotrophic culture; (C) Time course of glucose and KNO3 in mixotrophic and heterotrophic
culture. (D, E, and F) Fluorescence images of Nile red-stained cells cultured in mixotrophic
condition after 1 d (D), 3 d (E), and 6 d (F).
99
Our results show that lipid content in mixotrophic and heterotrophic cells began to increase as
nitrate ran out. Coincidently, the time in which the lipid level began to increase was exactly
when the starch accumulation slowed down or began to decrease. Therefore, it appears that
nitrate level correlates with both starch and lipid accumulation in reverse directions. In other
words, before nitrate deprivation, starch was largely accumulated, and after nitrate deprivation,
starch accumulation slowed down, while lipid accumulation increased.
5.4.2 Accumulation of starch and lipids depends on nitrogen status
To further characterize the metabolic changes triggered by nitrogen levels, algal cells were
transferred to the nitrogen (N)-free Kuhl medium supplemented with glucose and cultured for 2 d.
In N-starved cultures supplemented with 4 g L‒1 glucose, there was still an increase in starch
content (Fig. 5.2A); however, starch levels peaked much faster (12 h) and showed much lower
values (14%, in the light; 6%, in the dark) than cultures supplemented with nitrogen (Fig. 5.1A,
5.1B). The peak value of starch content for N-starved cells was only 43% of that observed in Nsupplemented cells in the dark, which was approximately half of the values in the light.
We tested whether lower starch levels in the N-free medium were caused by glucose limitation
and found that increasing glucose from 4 to 20 g L‒1 did not lead to a significant difference in the
maximum level of starch in both dark and light grown cells. After excluding the factor of
insufficient carbon availability, another explanation for lower starch levels is nitrogen limitation.
It is possible that the limited endogenous nitrogen of algal cells is depleted quickly in N-starved
culture conditions. This alters protein levels so that starch synthesis and its accumulation are
negatively influenced.
100
Lipid/starch content (%)
30
A
25
20
15
10
5
0
0.0
0.5
1.0
1.5
2.0
Time (d)
Lipid/starch content (%)
30
B
25
20
15
10
5
0
0.0
0.5
1.0
1.5
2.0
Time (d)
Figure 5.2 Time course of lipid and starch accumulation in C. sorokiniana in response to N
starvation. Cells were first grown in Kuhl medium until the exponential phase, and then switched
to N-free Kuhl medium supplemented with 4 g L‒1 (A) or 20 g L‒1 glucose (B) and further
cultured for 2 d. Lipid (closed symbols) and starch (open symbols) content was monitored under
light intensities (µmol m─2 s─1) of 0 (down-triangle), 50 (square), and 100 (circle).
101
Results show that lipid accumulation lagged behind that of starch, and lipid content increased
rapidly after a decrease in starch. Under the same light conditions, algal cells supplied with 20 g
L‒1 glucose in the N-free medium accumulated similar levels of lipids as the N-starved cells with
4 g L‒1 glucose (Fig. 5.2). This result counters the report that lipid accumulation in N-starved
cells depends strictly on increased carbon, and that lipid content increases steadily as acetate
increases (Fan et al., 2012). This suggests that algal cells in N-depleted conditions utilize carbon
sources differently, and that other factors beyond carbon availability determine the upper limits
of lipid accumulation in C. sorokiniana.
5.4.3 The biosynthesis of starch and lipids are both benefitted from active photosynthetic
activity
The aforementioned results indicate lower levels of starch and lipids in the heterotrophic culture
than in the mixotrophic culture. This highlights the significant role played by photosynthesis in
the products of energy storage, and points to ATP and/or NADPH as critical factors in regulating
levels of these storage compounds.
In photoautotrophic cells, photosynthetic electron transport is the only source of ATP and
NADPH for synthesis of energy storage products. However, in heterotrophic cells, the energy
and reducing potential equivalents are supported by the oxidation of organic carbon that also
serves as carbon skeletons in energy storage products.
This study aimed to determine the role of photosynthetic electron transport in storage product
accumulation in response to mixotrophic culture conditions. Therefore, we applied the PSII
inhibitor DCMU, which inhibits photosynthesis by interrupting the flow of electrons from PSII
to plastoquinone (Rippka, 1972). As shown in Fig. 5.3, adding DCMU reduced the accumulation
of both starch and lipids in the mixotrophic culture after 3 and 6 d. Levels remained more or less
102
constant with the corresponding heterotrophic culture. These results support the notion that the
light reactions of photosynthesis are an important source of free energy and reductant for the
synthesis of starch and lipids in C. sorokiniana under mixotrophic conditions.
50
Lipid content (%)
40
Mixotrophy
Mixotrophy + DCMU
Heterotrophy
30
20
10
0
Starch content (%)
25
20
15
10
5
0
3
6
Time (d)
Figure 5.3 Starch and lipid accumulation in C. sorokiniana in response to mixotrophic growth is
dependent on photosynthetic electron transport.
5.4.4 Lipid biosynthesis does not require high light intensity
To further understand the role of light in the formation of storage reserves, we determined the
amounts of starch and lipid in cells starved for N under three light conditions; 0 (dark condition),
50, and 100 µmol m─2 s─1. As shown in Fig. 5.2, N-starved cells grown under light intensities of
103
50 and 100 µmol m─2 s─1 accumulated equal amounts of starch, which were higher than that of
N-starved cells grown in the dark condition.
Generally, lipid content showed a slow increase at the beginning of cultivation. This increase
was enhanced after starch content peaked under all light conditions (Fig. 5.2A, 5.2B). In addition,
light exerted a considerable effect on lipid content. As observed for starch, N-starved cells grown
in the two light conditions showed higher lipid levels than did cells grown in the dark. However,
the maximum lipid level (28%) was obtained under a light intensity of 50 µmol m─2 s─1 instead
of 100 µmol m─2 s─1. It should be also noted that under a light intensity of 100 µmol m─2 s─1, Nstarved cells grown for 2 d underwent a drastic decrease in chlorophyll content and eventually
died (data not shown). This may have been caused by photooxidative damage to the N-starved
cells under a higher light intensity (Solovchenko et al., 2008). Thus, although photosynthetic
light reactions are beneficial for starch and lipid accumulation, lipid biosynthesis does not
require a high light intensity in the N-starved cells.
5.4.5 Declining starch levels are mainly attributed to starch catabolism
To determine whether the declining starch level was due to a decrease in starch anabolism or an
increase in starch catabolism, we investigated the expression levels of three important genes
involved in starch metabolism under the mixotrophic growth of C. sorokiniana (Fig. 5.4).
Transcript levels of AGPase increased after cultivation for 3 and 6 d. This may indicate feedback
regulation by decreased starch content in algal cells. For β-amylase and iso-amylase, the
expression levels after 3 d were increased by 40% and 288%, respectively, compared with those
after 1 d. The mRNA level of iso-amylase continued to increase after 6 d, which was 9.4-fold
higher than that after 1 d. The increased mRNA levels of β-amylase and iso-amylase were
consistent with decreased starch content, which suggests that starch degradation occured.
104
Consistently, the significantly increased levels of glucose and glucose 1-phosphate were
indicators of starch degradation after 3 d (Fig. 5.7). It suggests that the starch degradation rate is
higher than the starch synthesis rate, if we consider that increased mRNA levels of AGPase
indicate enhanced starch synthesis.
12
Relative mRNA levels
10
1 day
3 day
6 day
8
6
4
2
0
ase
AGP
se
se
se
Acca eta-amyla Iso-amyla
B
Figure 5.4 RT-PCR results of critical enzymes involved in the synthesis of starch and lipid and in
the degradation of starch.
5.4.6 Lipid synthesis is largely dependent on starch degradation
The aforementioned results demonstrate that starch degradation occurs as lipid levels increases.
To determine whether starch degradation facilitates carbon partitioning into the fatty acid
synthesis pathway and increases lipid content, we added CHX to the medium. CHX is an
105
antibiotic that inhibits eukaryotic cytoplasmic protein synthesis without significantly reducing
protein synthesis in the chloroplast. It is important to note that CHX inhibits the induction of βamylase mRNAs, which requires de novo synthesis of proteins in the cytoplasm (Ohto et al.,
1992). CHX also greatly reduces amylase activity (Kato et al., 2002; Todaka et al., 2000). In our
study, there was no significant difference in starch and lipid content for algal cells with or
without CHX after 3 d (Fig. 5.5). This indicates that CHX had no influence on the synthesis of
starch and lipids, which reside largely within the chloroplast. However, algal cultures without
CHX showed greatly reduced starch levels after 6 d. At the same time, algal cultures with CHX
showed a much smaller decrease, and contained twice the starch of cells without CHX. On the
other hand, algal cultures without CHX showed a 1.8-fold increase in lipid content after 6 days,
while lipids in algal cultures with CHX did not show an obvious increase.
Thus, the substantially increased lipid levels in algal cells correspond to significantly
decreased starch levels. A similar phenomenon has been observed in microalga
Pseudochlorococcum sp. (Li et al., 2011). The partial inhibition of starch degradation resulted in
a decrease of lipid content, indicating the conversion of starch to lipid in C. sorokiniana, which
was also supported by major metabolite accumulation patterns (Fig. 5.7). After 6 days, levels of
glucose and glucose 1-phosphate in algal cells significantly increased, reflecting drastic starch
degradation (Smith et al., 2005). On the other hand, most fatty acids showed very strong positive
values compared to those measured at 1 and 3 d, indicating enhanced lipid synthesis. However,
the gene expression level of ACCase dramatically declined when lipid content increased. Similar
results have been reported in Chlamydomonas reinhardtii, indicating ACCase gene expression
was not upregulated during lipid accumulation. This suggests that this enzyme may not be
regulated at the transcriptional level (Lv et al., 2013).
106
Taken together, these results suggest that lipid synthesis is largely dependent on starch
degradation, and that C. sorokiniana cells show the strategy of rerouting the carbon skeleton
from starch to lipids.
50
Mixotrophy + CHX
Mixotrophy
Lipid content (%)
40
30
20
10
0
Starch content (%)
25
20
15
10
5
0
3
6
Time (d)
Figure 5.5 Effect of cycloheximide (CHX, 1 mg L‒1) on changes of starch and lipid content in
mixotrophic growth of C. sorokiniana.
5.4.7 Metabolites profiling in C. sorokiniana
In this study, we used gas chromatography-time of flight mass spectrometry in conjunction with
automatic mass spectral deconvolution and database annotation to profile primary metabolites.
107
From a total of 612 unique metabolic peaks positively detected at s/n>5 in at least three extracts,
we structurally identified 124 non-redundant metabolites. We used principal component analysis
(PCA) to determine how metabolite abundance patterns relate to the time course of C.
sorokiniana growth under mixotrophic conditions. The loadings plot from the PCA analysis
showed three distinct groups at each time point (Fig. 5.6). The group including the 1 d samples
was closer to the 3 d samples than to the 6 d samples. Major metabolite accumulation patterns
also reflected this relationship, which accords with differential metabolite profiling within these
three time points (Fig. 5.7). Levels of most of the metabolites involved in the TCA cycle
(organic acids) and amino acids decreased after 6 d and remained suppressed thereafter.
However, two central metabolites, glutamine and malate, showed rapid and high accumulation.
Glutamine is known to play a critical role in C/N partitioning and N assimilation in plant cells.
Considering the N-depleted conditions in the tested cells after 6 d, it appears that the cells
actively maintain high levels of this metabolite to enable rapid response to changing N levels.
Elevated levels of malate suggest suppressed respiration in these cells, resulting from a slowing
of cell growth in response to N deprivation.
108
Figure 5.6 Modeling of metabolite shifts during mixotrophic growth. A PLS-DA model
illustrates differences in metabolite levels between the three time points (1, 3 and 6 d).
109
Figure 5.7 Primary metabolite profiles in C. sorokiniana during the mixotrophic growth with
time course. Time points presented for sugars and sugars phosphate, fatty acids, organic acids,
and amino acids are 1, 3 and 6 d. Data are normalized to the mean standard deviation. Values
presented are means of three replicates.
110
5.5 DISCUSSION
5.5.1 Accumulation patterns of starch and lipids
Our results show that when nitrogen is replete and/or depleted over short periods in the
medium, starch is the predominant carbon storage, with little lipid accumulation. On the other
hand, when nitrogen is depleted, large amounts of lipid accumulation occurs, corresponding to
the gradual decrease of starch content (Fig. 5.1, Fig. 5.2). Li et al. (2011) have reported similar
starch and lipid accumulation patterns in the microalga Pseudochlorococcum (Li et al., 2011),
suggesting that algal cells use starch as a short-term carbon and energy storage product, and that
cells shift the carbon partitioning into neutral lipids as a secondary storage product after
prolonged nitrogen depletion.
Previous studies demonstrate several different kinds of starch and lipid regulation patterns in
microalgae. Both starch and lipid content in C. sorokiniana are enhanced with 2% CO2 as
opposed to air (VanderGheynst et al., 2014). A marine microalgae Tetraselmis subcordiformis
reached 54% of starch immediately following nitrogen deprivation without lipid accumulation
(Yao et al., 2012). Takeshita et al. (2014) found a strain-dependent accumulation pattern. For C.
sorokiniana, starch accumulation was induced and accompanied by decreased lipid accumulation.
For other Chlorella strains, lipid content increased with decreasing starch content (Takeshita et
al., 2014). In C. reinhardtii mutant, which lacks cell walls, starch is the dominant sink for carbon
storage, and rapid lipid synthesis occurs only when the carbon supply exceeds the capacity of
starch synthesis (Fan et al., 2012). Furthermore, a microalga Chlorella zofingiensis possesses
basal levels of starch and lipids under nitrogen-replete conditions, and large amounts of starch
accumulate under nitrogen-starvation conditions. This is followed by increased lipid content with
partially degraded starch (Zhu et al., 2014). This suggests that distinct factors are involved in
111
sensing and rapidly responding to the N-deprived condition in different alga cells. Future
research to identify and characterize the role of these specific regulatory proteins will provide a
clear understanding of how the switch to N-depleted medium is indeed sensed and then
transduced to large metabolic changes, which enables this aquatic, single-celled organism to
survive under large environmental stresses.
5.5.2 Influential factors for lipid synthesis
This study demonstrated that the light reaction of photosynthesis is an important source of free
energy and reductant for lipid synthesis in C. sorokiniana (Fig. 5.2, Fig. 5.3). Our previous work
showed that the refixation of CO2 through photosynthetic activity is critical for biomass
synthesis (Li et al., 2014a). This provides higher carbon availability to produce energy storage
products. Excess carbon from photosynthesis has been found to partition into the lipid synthesis
pathway (Suen et al., 1987). Therefore, photosynthetic activity provides a carbon source for lipid
accumulation along with energy.
On the other hand, significantly higher lipid production may serve as a protective mechanism
in mixotrophic cells. Under optimal growth conditions, large amounts of algal biomass are
produced, partly driven by the ATP and NADPH produced from photosynthesis. When cell
growth and proliferation is impaired by N limitation, the pool of NADP+, the major electron
acceptor for photosynthesis, can become depleted as it becomes over-reduced (due to high levels
of NADPH build-up). This leads to over-oxidation of the PSII reaction center, causing photooxidative damage, reactive oxygen species generation, and overall cellular damage. Since
NADPH is consumed in fatty acid biosynthesis, increased lipid production may be a means used
by the cell to protect itself from oxidative damage (Hu et al., 2008).
112
The contribution of the carbon source from photosynthesis to fatty acid production is
weakened due to the degradation of photosynthetic machinery in N-deficient conditions. A
marked decrease in chlorophyll and enzymes essential for CO2 fixation, such as Rubisco, has
been observed in nitrogen-starved Chlamydomanas cells (Fan et al., 2014; Msanne et al., 2012).
This may be due to that the degradation of photosynthetic machinery intensifies cellular damage
under higher light intensity, reducing lipid accumulation compared with that in the lower light
condition (Fig. 5.2). However, the decreased cellular components are recycled for neutral lipid
biosynthesis (Fan et al., 2014; Msanne et al., 2012). Our results show that conversion of nonlipid cellular components to lipids is supported by changes in the metabolic profile (Fig. 5.7).
Fatty acids increased considerably, amino acids diminished significantly, and high levels of
glutamine caused rapid turnover of metabolites in N-deprived algal cells. It is notable that
substantial accumulation of lipids, mostly in the form of TAG observed through Nile red staining
(Fig. 5.1), were accompanied by starch degradation after prolonged incubation in the N-deprived
medium. Starch degradation contributed to lipid synthesis in C. sorokiniana (Fig. 5.4, Fig. 5.5,
and Fig. 5.7). Taken together, these results show that photosynthesis and turnover of other
cellular components are involved in the lipid synthesis in mixotrophic cells of C. sorokiniana
under N-deficient conditions.
5.5.3 The relationship between starch and lipid contents
Studies indicate a competitive relationship between starch and lipid synthesis due to their
common precursors. Blocking the starch synthesis pathway results in substantially higher lipid
accumulation in Chlamydomonas and Chlorella (Fan et al., 2012; Ramazanov & Ramazanov,
2006). Therefore, this is a promising technique to enhance oil production in microalgae.
However, the complexity of carbon partitioning goes beyond mere competition between starch
113
and lipid synthesis. Starchless mutants of C. reinhardtii cannot over-accumulate lipids (Siaut et
al., 2011), show restricted growth rates (Darzins et al., 2010), and exhibit a decrease in overall
anabolic processes (Work et al., 2010). The supply of carbon from starch is vital for normal plant
growth. In Arabidopsis, like most vascular plants, starch plays an important role in the
carbohydrate metabolism of the leaf. Glucose produced through photosynthesis during the day is
retained mainly in the form of starch granules in the chloroplast. Starch is then degraded at night
to provide a constant supply of carbohydrate in the absence of photosynthesis. Mutant
Arabidopsis plants that cannot synthesize starch or degrade it at night show reduced growth rates
under most conditions (Smith & Stitt, 2007). Therefore, the increase in starch content in C.
sorokiniana during N-replete culturing probably plays an important physiological role and
represents a mechanism to increase the synthesis of a primary bioenergy carrier (starch) at the
expense of other cellular constituents used for cell division (proteins and nucleic acids).
5.5.4 Comparison of starch and lipid accumulation from an energy perspective
Compared with lipids, starch synthesis is an efficient means of storing carbon. The synthesis
of fatty acids, which are major components of lipids, represents a significant loss of carbon due
to the conversion of pyruvate (C3) to acetyl-CoA (C2) by pyruvate dehydrogenase. Furthermore,
if we account for the energy of starch and lipids with the active import of CO2, the ATP and
NADPH requirements for starch synthesis are 50% and 45% lower, respectively than those
required for TAG (C55H98O6) synthesis (Subramanian et al., 2013). In addition, glucose can be
directly converted into starch without prior conversion to triose-phosphates (C3) (Fettke et al.,
2010), which requires only one ATP per glucose.
On the other hand, although starch represents a more accessible form of carbon storage for
plant cells than fatty acids, the energy recovery from fatty acid oxidation is greater than that of
114
starch oxidation. When fatty acids are oxidized via the β-oxidation pathway and the citric acid
cycle, the energy recovery is approximately 6.7 ATP equivalents per carbon for palmitic acid.
For glucose oxidation via glycolysis as well as the citric acid cycle, energy recovery is 5 ATP
equivalents per carbon (Johnson & Alric, 2013). This higher energy/mass ratio for oil may
explain why oils are often a preferred storage reserve in seeds. Namely, smaller and lighter seeds
may confer a selective advantage for dissemination.
In C. sorokiniana, starch as the preferred carbon storage in N-replete condition guarantees the
highest efficiency for energy capture. Furthermore, C. sorokiniana cells can reroute all the
carbon into only one sink (lipids) instead of two sinks (starch and lipids) in the N-depleted
condition. The straight substitution of lipids for starch in the final biomass offers an effective
strategy for energy accumulation. By degrading starch in the chloroplast, algal cells generate
more physical space and have a higher capacity to host lipid bodies. This has been shown to
relate to increased lipid levels in starchless mutants of C. reinhardtii (Goodson et al., 2011).
Therefore, it is not necessary to block the starch synthesis pathway to increase lipid content in C.
sorokiniana. Most studies that successfully increased lipids by blocking the starch synthesis
pathway found decreased growth in the engineered strains (Li et al., 2010a; Li et al., 2010b;
Work et al., 2010).
However, to determine the oil-producing ability of microalgae, both lipid content and biomass
production should be considered simultaneously. Mixotrophic growth of C. sorokiniana
promised to be an efficient lipid production strategy only by optimizing the cultivation
conditions. In addition, engineering this strain by disrupting lipid catabolism shows promise.
Most studies of this strategy focused on higher plants and yeast, and their limited success may
have been due to the requirement of cellular energy from fatty-acid oxidation for proper seedling
115
development (Fulda et al., 2004; Germain et al., 2001). However, the microalga Thalassiosira
pseudonana showed greatly increased lipid content without compromising growth through the
knock-out of lipase genes involved in lipid catabolism (Trentacoste et al., 2013).
116
5.6 CONCLUSION
Our study reveals the dynamic carbon partitioning that occurs between starch and lipids during
cell growth of C. sorokiniana. This species uses starch as a primary carbon and energy storage
compound under nitrogen-replete conditions, and lipids become major storage products
accompanied by starch degradation as nitrogen is depleted. Starch degradation is necessary for
partial lipid accumulation under nitrogen-depleted conditions. The distinct accumulation pattern
of starch and lipids, high growth rate, low light intensity requirement for lipid accumulation,
along with high lipid content, all point to the promise of mixotrophic cultivation of C.
sorokiniana as an efficient production strategy.
117
CHAPTER SIX
SUMMARY
This dissertation study explored different cultivation strategy of a microalga Chlorella
sorokiniana for biofuel production and characterized its photosynthesis regulation and carbon
partitioning between starch and lipid. The major conclusions are:
 The feasibility of C. sorokiniana for heterotrophic cultivation with high cell density was
proved. Under the optimized culture conditions, the algal biomass reached 37.6 g L‒1 in
batch fermentation, with 80 g L‒1 of initial glucose and 4 g L‒1 of initial KNO3. Moreover,
this microalga could tolerate high temperature (up to 42 oC) and showed the best growth
performance at 37 oC.
 Mixotrophy was demonstrated as the most efficient strategy for cultivating this microalga
under the tested culture conditions, with the highest growth rates, the highest maximum
biomass, and the highest lipid productivity. The biomass yield (based on consumed
glucose) in mixotrophic culture (0.82 g g
culture (0.34 g g
─1
─1
) was much higher than in heterotrophic
), which are attributed to linear electron flow and refixation of CO2.
Moreover, photosynthesis also had profound impact on lipid content, with the lipid level
of 45% in mixotrophic cells (50 µmol m─2 s─1) and 13% in heterotrophic cells.
 Photosynthesis in mixotrophic cells of C. sorokiniana is dynamically adjusted according
to algal cell status at different growth stages. Mixotrophic cells in exponential phase
(cultivation for 1 day) exhibited enhanced photosynthetic activity; yet cells in stationary
phase (cultivation for 3 day) showed down-regulation of photosynthesis, which was
proved to be closely correlated with nitrogen limitation. To maintain an efficient
118
photosynthetic activity for biomass accumulation, efficient nitrogen supplementation is
necessary.
 Microalga C. sorokiniana used starch as a primary carbon and energy storage compound
under nitrogen-replete conditions, whereas lipids became major storage products
accompanied by starch degradation as nitrogen was depleted. Starch degradation is
necessary for partial lipid accumulation under nitrogen-depleted conditions. The distinct
accumulation condition for lipids in comparison to starch, together with high growth rate,
low light intensity requirement, maximum biomass, and high lipid content, makes
mixotrophic cultivation of C. sorokiniana as a promising production strategy.
This research implies that mixotrophic culture of C. sorokiniana has great potential to be
adopted for biofuel production. Mixotrophic growth not only keeps the features of organic
substrate metabolism, but also keeps photosynthetic activity. Firstly, mixotrophic growth
enhances photoautotrophic activity and reduces inorganic carbon (CO2) from organic substrate
metabolic pathways into organic carbon, which contributes to efficient utilization of glucose and
finally leads to the increased biomass yield. Secondly, mixotrophic growth of C. sorokiniana at
higher temperature (37 oC) shows similar performance with that at 25 oC, which indicates the
possibility of mixotrophic culture in summer season. Thirdly, oxygen production from the
photoautotrophic part could be consumed in the respiration process; therefore photooxidative
damage could be efficiently alleviated or eliminated in mixotrophic culture. Fourthly, much
higher lipid content could be accumulated in mixotrophic cells, largely contributed by turnover
of starch.
However, several issues about mixotrophic cultivation are still needed to be considered for
large-scale production. 1) Organic carbon is a major energy source for mixotrophic cells, but the
119
cost of substrate would be a major limitation. However, the organic carbon is more efficiently
utilized with mixotrophy than heterotrophy. 2) At the beginning of cultivation, high light
intensity is required for cell growth to guarantee the highest growth rate and maximum biomass
production; while at the last stage of growth, low light intensity is beneficial for
overaccumulation of lipid. Therefore, light intensity must be adjusted during cultivation
processes. Although the sunlight is almost everywhere on the earth for free and seems desirable,
the stability, uniformity, and continuity of sunlight cannot be manipulated, which may impede
the yields of mixotrophic culture. Therefore, the suitable choice should be artificial light sources.
This would result in high illumination costs and may additionally offset the advantages of
mixotrophic growth. 3) Another challenge with mixotrophic cultivation is contamination from
heterotrophic bacteria and other microorganisms. Even if photobioreactors are utilized,
contamination is still a concern and can lead to costly system shut-downs if not properly
managed.
The traditional strain selection and cultural process optimization is one viable approach for
developing high-lipid-containing algal cultivation strategy. The recent development of molecular
techniques for microalgal strain optimization offers an alternative method to increase lipid
production. The final objective is increasing lipid content without decreasing cell growth. There
has been an emerging concept that overaccumulation of lipid bodies can be achieved in starchdevoid mutants. However, not all cases can support this strategy and all successful studies with
increased lipids showed decreased algal growth in the engineered strains. This study showed that
lipid was the only carbon sink in the final algal biomass of C. sorokiniana. Therefore, it is not
necessary to disrupt carbohydrate pool and make engineering efforts to reroute carbon flux. It
has been reported that disrupting lipid catabolism has less impact on the primary carbon
120
pathways associated with growth, and it is a practical approach to increase lipid yields in
microalgae without affecting growth or biomass. Combined with the characteristics of one
ultimate carbon sink in C. sorokiniana, it must be promising to get overaccumulation of lipid
without compensating cell growth in C. sorokiniana mutants of lipid catabolism. At the same
time, this research will also advance knowledge on lipid metabolism and its connected pathways.
Additionally, biomass carbon in mixotrophic cells can be derived from both photosynthesis
and organic carbon metabolism. There exist several CO2 release processes in plants and algae: 1)
tricarboxylic acid cycle, 2) oxidative pentosephosphate pathway, 3) oxidative decarboxylation of
malate through NADP-malic enzyme, 4) conversion of pyruvate to Acetyl-CoA, and 5)
photorespiration in photosynthetic organisms under certain conditions. On the other hand, there
are also several CO2 fixation processes: 1) PEP carboxylation, 2) pyruvate carboxylation, and 3)
Calvin cycle, which is the most important strategy for CO2 fixation in nature. The involvement
and partitions of each pathway would be adjusted differently under heterotrophic and
mixotrophic conditions. The question how the central pathways in mixotrophic cells are
partitioned and adjusted remains to be answered. More in-depth understanding of carbon
metabolisms in mixotrophic cells would facilitate much better utilization and improvement of
mixotrophic cultivation.
121
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