DEVELOPING A TRANSGENIC MICROALGA TO ENHANCE ALGAL STARCH
UTILIZATION FOR FUEL AND CHEMICAL PRODUCTION
By
Xiaoqing Wang

A DISSERTATION
Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of
Biosystems Engineering – Doctor of Philosophy
2015

ABSTRACT
DEVELOPING A TRANSGENIC MICROALGA TO ENHANCE ALGAL STARCH
UTILIZATION FOR FUEL AND CHEMICAL PRODUCTION
By
Xiaoqing Wang
Due to the high growth rate, high photosynthetic efficiency, and relatively high
content of energy-rich carbohydrate, green microalgae have been viewed as one of the
most promising non-food-crop-based feedstocks for bioethanol production. However,
two main issues have to be addressed in order to commercialize microalgal bioethanol.
The first one is how to achieve the cost-effective and highly productive cultivation
process for microalgal carbohydrate production, and the second one is how to acquire
fermentable sugars from carbohydrate-rich microalgal biomass efficiently and
economically. To this end, a two-stage cultivation strategy was first evaluated to
enhance microalgal carbohydrate production. In addition, a transgenic microalga
producing thermophilic amylase in its chloroplast was constructed. By using the
transgenic microalga, a novel process of converting microalgal carbohydrate into
fermentable sugars was developed.

Dedicated to my lovely family and friends

	
  

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ACKNOWLEDGEMENTS

My utmost gratitude and respect to my advisor, Dr. Wei Liao, for his strong
support and patient guidance during my doctoral degree; for his optimistic attitude and
hardworking spirit that gave me a lot of motivation. Dr. Liao brought me into the
doctoral program and gave me the opportunity to work on diverse projects. His
persistence and constant encouragement kept me going through the difficulties and
challenges of my research work. I would like to thank Dr. Liao first and most
importantly.
A sincere thank you to my co-advisor, Dr. Yan Liu, for her a lot of guidance and
time; for being so kind and helpful not only in my research work but also in my life.
Thanks to her great help and effort, I can be accompanied with my family and
concentrate on my research work.
Very special thanks go to my committee member, Dr. Barbara B. Sears, for her
precious guidance, suggestion and time during the period of constructing the transgenic
microalga; for her kindly supplying the plasmid P699 that is the basis of gene
expression system developed in this work.
I would also like to acknowledge the contribution of another committee member,
Dr. David Hodge, for his valuable comments and suggestions during my dissertation
process. I am very thankful for his input.
I am sincerely grateful for Dr. Christoph Benning and his lab members Chia-Hong
Tsai and Benshen Liu for their contribution in the transgenic microalga project.
Many thanks go to my group members: Zhenhua Ruan, Danielle Boileau and Pat
Sheridan for their valuable input and help. Thanks are also extended to Julie Xu, Robert

	
  

iv	
  

Kraemer, Zhiguo Liu, Yuan Zhong, Rui Cheng, Tongjun Liu, Zhenlun Li, Yingkui
Zhong. Mauricio Bustamante.
My deepest gratitude goes to my parents and my husband. Without their
unconditional love and unending support, my accomplishments in life could not have
been achieved. I am so grateful for all of my family, especially my little daughter,
Emily, who gives me endless motivations and tremendous happiness that I could never
imagine.

	
  

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TABLE OF CONTENTS

LIST OF TABLES ..........................................................................................................ix
LIST OF FIGURES.........................................................................................................xi
KEY TO ABBREVIATIONS AND SYMBOLS ........................................................ xiii
CHAPTER 1 INTRODUCTION .................................................................................... 1
1.1 Bioethanol from renewable sources ....................................................................... 1
1.2 First-generation bioethanol .................................................................................... 2
1.2.1 Second-generation bioethanol ......................................................................... 2
1.2.2 Microalgae as a potential source for bioethanol production ........................... 3
1.3 Challenges in microalgae cultivation for carbohydrate accumulation ................... 4
1.3.1 Reactor configuration ...................................................................................... 4
1.3.2 Cultivation techniques for microalgal carbohydrate accumulation ................ 6
1.3.2.1 Nutrient control ........................................................................................ 6
1.3.2.2 Cultivation strategies ................................................................................ 9
1.4 Microalgal carbohydrate conversion into fermentable sugars ............................... 9
1.4.1 Chemical saccharification ............................................................................. 10
1.4.2 Enzymatic saccharification ........................................................................... 10
1.4.3 “Self-processing” saccharification ................................................................ 11
1.5 Recombinant protein expression in transgenic microalgae .................................. 12
1.5.1 Nuclear transformation.................................................................................. 13
1.5.2 Chloroplast transformation............................................................................ 14
1.5.2.1 Codon optimization ................................................................................ 16
1.5.2.2 Promoters ............................................................................................... 16
1.5.2.3 Transformation methods ........................................................................ 18
1.6 Goal and Objectives ............................................................................................. 18
CHAPTER 2 TWO-STAGE PHOTOAUTROPHIC CULTIVATION TO IMPROVE
CARBOHYDATE PRODUCTION IN CHLAMYDOMONAS REINHARDTII ............ 20
2.1 Abstract ............................................................................................................. 20
2.2 Introduction ....................................................................................................... 20
2.3 Methods ............................................................................................................ 22
2.3.1 The microalga strain and growth medium .................................................... 22
2.3.2 Experimental setup ........................................................................................ 23
2.3.2.1 Optimization of culture conditions in batch operation ........................... 24
2.3.2.2 Operation of semi-continuous cultivation in Stage I.............................. 25
2.3.2.3 Stage II of batch culture for carbohydrate accumulation ....................... 26
2.3.3 Ethanol fermentation ..................................................................................... 27
2.3.4 Analytical methods........................................................................................ 27
2.3.4.1 Cell growth measurement ...................................................................... 27
2.3.4.2 Carbohydrate, protein and ethanol analysis ........................................... 27
2.3.4.3 Statistical analysis .................................................................................. 28
2.4 Results and discussion ...................................................................................... 29

	
  

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2.4.1 Optimization of culture conditions in batch operation .............................. 29
2.4.2 Stage I of semi-continuous cultivation ...................................................... 34
2.4.3 Stage II of batch culture for carbohydrate synthesis.................................. 38
2.5 Conclusion ........................................................................................................ 42
CHAPTER 3 TRANSGENIC EXPRESSION OF A BACTERIAL TRANSGENIC
EXPRESSION OF A BACTERIAL THERMOPHILIC AMYLASE IN THE
CHLAMYDOMONAS REINHARDTII CHLOROPLAST .............................................. 43
3.1 Abstract ................................................................................................................ 43
3.2 Introduction .......................................................................................................... 43
3.3 Materials and methods ......................................................................................... 45
3.3.1 Strains and culture conditions ....................................................................... 45
3.3.2 Plasmid construction ..................................................................................... 46
3.3.3 Chloroplast transformation............................................................................ 50
3.3.4 DNA isolation and PCR amplification .......................................................... 50
3.3.5 Protein extraction and Western blot analysis ................................................ 51
3.3.6 Enzyme purification ...................................................................................... 51
3.3.7 Enzyme assay ................................................................................................ 52
3.4 Results .................................................................................................................. 52
3.4.1 Expression of arAmyBH in the C. reinhardtii chloroplast ............................ 52
3.4.2 Western blot analysis .................................................................................... 54
3.4.3 Purification and characterization of His6-tagged arAmyBH protein ............ 55
3.5 Conclusions .......................................................................................................... 56
CHAPTER 4 IN VIVO ENZYMATIC HYDROLYSIS OF TRANSGENIC
CHLAMYDOMONAS REINHARDTII STARCH ....................................................... 57
4.1 Abstract ................................................................................................................ 57
4.2 Introduction .......................................................................................................... 57
4.3 Materials and methods ......................................................................................... 59
4.3.1 Strains and culture conditions ....................................................................... 59
4.3.2 Cell growth measurement ............................................................................. 60
4.3.3 Starch analysis ............................................................................................... 60
4.3.4 Enzyme assay ................................................................................................ 61
4.3.5 Effects of pH, temperature and metal ions on arAmyBH activity ................ 61
4.3.6 Hydrolysis of microalgal starch .................................................................... 62
4.3.7 Hydrolysate profile analysis .......................................................................... 62
4.4 Results .................................................................................................................. 63
4.4.1 Enzyme properties of arAmyBH ................................................................... 63
4.4.2 Starch hydrolysis in microalgal cells ............................................................ 65
4.4.3 Cell growth and starch accumulation of wild-type and recombinant strains in
................................................................................................................................ 68
4.5 Conclusion............................................................................................................ 70
CHAPTER 5 CONCLUSION AND FUTURE WORK ................................................ 71
5.1 Conclusions .......................................................................................................... 71
5.2 Future work .......................................................................................................... 73
APPENDIX .................................................................................................................... 75

	
  

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BIBLIOGRAPHY .......................................................................................................... 90

	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  

	
  

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LIST OF TABLES

Table 1 Carbohydrates of selected microalgal species (adopted from Markou et al.,
2012)(Markou et al., 2012) ........................................................................................... 6
Table 2 Carbohydrate content of green microalgae species in N-deprivation ...................... 7
Table 3 Recent successes in recombinant protein production in C. reinhardtii chloroplast15
Table 4 Chloroplast promoters of C. reinhardtii used for the expression of heterologous
proteins (adopted from (Rosales-Mendoza et al., 2012)) ............................................ 17
Table 5 Worksheet for the 18-run, 3-factor CRD optimization experiment ....................... 24
Table 6 Response of 18-run CRD experiment design ......................................................... 30
Table 7 Performance of biomass production and carbohydrate content of semi-batch
culture of C. reinhardtii CC125 with different medium replacement ratio under
optimal conditions obtained from batch tests.............................................................. 37
Table 8 Codon usages of wild type AmyB gene and synthesized arAmyBH gene ............ 46
Table 9 Purification of arAmyBH from transformed C. reinhardtii. arAmyBH activity was
determined at 75 °C in 100 mM sodium phosphate buffer (pH 6.5) with 3,5dinitrosalicylic acid method. ....................................................................................... 55
Table 10 Biomass concentrations under different culture conditions for Figure 3 ............ 76
Table 11 Biomass concentrations before and after medium replacement for Figure 4 ...... 80
Table 12 Cell density and biomass concentration during the Stage II cultivation for Figure
5 ................................................................................................................................... 81
Table 13 Biomass compositions with different CO2 contents in the Stage II cultivation for
Figure 6 ....................................................................................................................... 82
Table 14 Enzyme activities at different pH for Figure 11a ................................................. 83
Table 15 Enzyme activities at different temperature for Figure 11b .................................. 84
Table 16 Enzyme activities after incubation at different temperature for Figure 11c ........ 85
Table 17 Amount of sugars from starch hydrolysis for Figure 12a .................................... 86
Table 18 Hydrolyzed starch with or without alpha-amylase from two strains for Figure
12b ............................................................................................................................... 87
Table 19 Comparison of biomass concentration between wild-type and transgenic strains
for Figure 13 ................................................................................................................ 88
	
  

	
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Table 20 Comparison of starch content between wild type and transgenic strains for Figure
13 ................................................................................................................................. 89

	
  

	
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LIST OF FIGURES

Figure 1 Annual ethanol use in U.S. fuels (data from the website of American Coalition
for ethanol) .................................................................................................................... 1
Figure 2 Schematic representation of the two-stage microalgal cultivation ....................... 26
Figure 3 C. reinhardtii CC125 growth curve in batch culture under different culture
conditions: (A) light intensity of 10 klux and NH4+ concentration of 4.7 mM (B) 10
klux light intensity and 9.4 mM NH4+ concentration (C) 10 klux light intensity and
18.8 mM NH4+ concentration (D) 5 klux light intensity and 4.7 mM NH4+
concentration (E) 5 klux light intensity and 9.4 mM NH4+ concentration (F) 5 klux
light intensity and 18.8 mM NH4+ concentration. ..................................................... 31
Figure 4 Semi-continuous cultivation of C. reinhardtii CC125 under the optimal condition
obtained from batch tests. The culture broth was replaced by 45 %, 70 %, 83 % and
91 % with fresh medium in different semi-continuous tests, each of which was
operated for six repeated cycles. Data are average of two experimental replicates. ... 35
Figure 5 Changes of biomass and cell density of C. reinhardtii CC125 during the Stage II
cultivation. Data are average of two experiment replicates ........................................ 39
Figure 6 Biomass compositions of C. reinhardtii CC125 cultured with (A) 5 % CO2 and
(B) 0.04 % CO2 in the Stage II cultivation. Data are average of two experimental
replicates. .................................................................................................................... 40
Figure 7 Flow chart of plasmid construction. (a) Plasmid p699. (b) Plasmid pBluescript II
KS (+). (c) Plasmid pBp699. (d) arAmyBH chloroplast expression cassette. (e)
Plasmid p666 with primer sites indicated as P1, P3, and P9. (f) Homologous
integration site in the cpDNA segment. ...................................................................... 49
Figure 8 PCR assays of expression cassettes (a) PCR assays of arAmyBH cassette (2.3 kb)
using primers P1 and P3. Lane 1: negative control (PCR products from wild-type
strain); lane 2-5: the PCR products from C. reinhardtii transformants; lane 6: positive
control (PCR products from plasmid p666); lane 7: 1 kb DNA marker (NEB). (b)
PCR assays of the adjacent arAmyBH and aadA cassettes (3.8 kb) using primers P1
and P9. Lane 1: negative control (wild-type); lane 2: PCR products from C.
reinhardtii transformant 7-C-1; lane 3: positive control (PCR products from plasmid
p666); lane 4: 1 kb DNA marker (NEB). (c) Primers P1 and P9 mapping to the
endogenous gene sequence. Dashed line indicates the insertion site for arAmyBH and
aadA cassettes. ............................................................................................................ 53
Figure 9 Western blot of wild-type and transformed C. reinhardtii and E. coli JM109.
Equal amounts of protein (25 µg) were loaded in each lane. Lane1: C. reinhardtii
transformant 7-C-1; lane 2: transgenic E. coli JM109 with p666; lane 3: wild-type C.
reinhardtii; lane 4: wild-type E. coli JM109 ............................................................... 54

	
  

	
   xi	
  

Figure 10 Partial purification of recombinant arAmyBH protein from transformed C.
reinhardtii. Lane 1: crude enzyme extracts; lane 2: partially purified arAmyBH; lane
3: protein marker. ........................................................................................................ 56
Figure 11 Effects of pH and temperature on the activity of arAmyBH. (a) Effect of pH on
arAmyBH amylase activity. (b) Effect of temperature on amylase activity. (c)
Thermostability of amylase activity of arAmyBH. Data points are the means of two
replicate determinations. The amylase activity of a pre-incubated sample at 4 °C was
regarded as 100%. ....................................................................................................... 63
Figure 12 (a) Amount of sugar obtained from starch hydrolysis of transgenic strain 7-C-1.
(b) Enzymatic hydrolysis of microalgae starch with alpha-amylase. Data are the
average of two replicates with standard error. ............................................................ 65
Figure 13 Comparison of growth condition and starch accumulation between wild-type
and transgenic strains in autotrophic culture. (a) Growth curves of wild-type and
transgenic strains in HS medium with 5 % CO2 under 10 klux continuous light (b)
Starch accumulation in wild-type and transgenic strains cultured in HS-N medium
with 5 % CO2 under 10 klux continuous light. Data are the average of two replicates
with standard error. ..................................................................................................... 68

	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  

	
  
	
  

	
  xii	
  

KEY TO ABBREVIATIONS AND SYMBOLS

Abbreviations:
NADPH: Nicotinamide adenine dinucleotide phosphate
ATP: Adenosine triphosphate
DNA: Deoxyribonucleic acid
RNA: Ribonucleic acid
mRNA: Messenger RNA
RUBISCO: Ribulose-1,5-bisphosphate carboxylase/oxygenase
CCM: carbon concentrating mechanism
w w-1: weight/weight
ATGC (NUCLEAR TIDE): Adenine,Thymine, Guanine, Cytosine
3’-UTR / 5’-UTR: three prime untranslated region/five prime untranslated region
vvm: volume/volume/minute
vv-1: volume/volume
mM: mili-mole
rpm: Revolutions per minute
HPLC: High performance liquid chromatography
DCW: Dry cell weight
Ni-NTA: Nickel nitrilotriacetic acid
PCR: Polymerase chain reaction
Kb: Kilobase
Bp: Base pair
DE value: Dextrose equivalent
DP1 DP2 DP3 DP4: Degree of polymerization
	
  

	
  xiii	
  

	
  
	
  
CHAPTER 1 INTRODUCTION

1.1 Bioethanol from renewable sources
Environmentally friendly bio-based fuels and power can make important contributions
to U.S. energy security, rural economic development, and environmental quality. As
outlined by the federal government in the Advanced Energy Initiative (AEI), the national
goal is to displace up to 30% of the nation’s gasoline consumption by 2030 (The White
House National Economic Council, 2009). Biomass, derived from photosynthesis,
including plants and microbes, has been viewed as a great alternative source to produce
renewable fuels, such as bioethanol, biodiesel, and biohydrogen, which can displace a
significant amount of fossil fuel consumption, and make major contribution to realizing
the national goal (Chen et al., 2013).
Bioethanol is one of the most successful and widely used biofuels that has been
developed by biorefining industry to date. Fig.1 illustrates ethanol consumption as a
transportation fuel in the U. S. since 2000.

Figure 1 Annual ethanol use in U.S. fuels (data from the website of American
Coalition for ethanol)

	
  

	
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It has undergone an exponential increase from 1.6 billion gallon per year in 2000 to 13.9
billion gallon per year in 2011. Bioethanol is mainly produced by yeast fermentation of
sugar-based feedstocks (Bai et al., 2008). Based on the type of feedstock, bioethanol can
be generally classified into “First-generation bioethanol” and “Second-generation
bioethanol” (Nigam & Singh, 2011).
1.2 First-generation bioethanol
First-generation bioethanol refers to yeast ethanol production on sugar and starch
crops. Corn grains and wheat grains as starch crops and sugar canes and beets are the
common feedstocks. It has been commercially proven that the yeast fermentation process
on sugar and starch crops for fuel ethanol production is technically and economically
feasible. However, the issue with the first-generation bioethanol is on the feedstock side.
Starch or sugar based crops are also important food sources for human consumption (John
et al., 2011; Nigam & Singh, 2011). With rapid increase of world population, the food
production industry is facing serious challenges of how to improve food production to
satisfy the demand. Using food resource to produce bioethanol creates a food vs. fuel
competition, which is certainly not a sustainable approach that biofuel production should
take. Therefore, other abundant non-food feedstocks must be investigated for
biofuel/bioenergy production.
1.2.1 Second-generation bioethanol
Due to the food vs. fuel issue that the first-generation bioethanol is encountering,
lignocellulosic biomass and other non-food starch materials are targeted for bioethanol
production. The ethanol from non-food based feedstock is defined as the secondgeneration bioethanol (John et al., 2011; Kim & Dale, 2004; Nigam & Singh, 2011).
Lignocellulose including perennial energy crops and crop residues is one of the most
	
  

	
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abundant biomass that can be used for biofuel production. Using them to produce
bioethanol not only addresses food vs. fuel problem, but also enhance the land use
efficiency and reduce carbon footprint from crop operations. Nevertheless, the recalcitrant
structure of lignocellulosic biomass requires severe chemical/thermal pretreatment,
complicated sugar composition and toxic byproducts from the pretreatment need new and
robust microbes to carry on the fermentation, and low fuel concentration demands energy
intensive post-processing. Many studies are on the way to address these barriers, though
the commercial production of second-generation bioethanol is not there yet to date (Hadar,
2013).
1.2.2 Microalgae as a potential source for bioethanol production
Considering the aforementioned issues related to bioethanol production, aquatic
microalgae are gaining more attention as a feedstock for the third-generation bioethanol
production. Microalgae, as unicellular photosynthetic organisms, are able to harness
sunlight and assimilate carbon dioxide to form plastid starch and cellulose-based cell walls
as the main carbohydrate sources. Both starch and cell wall polysaccharides can then be
processed to produce bioethanol (Buleon et al., 1997; Chen et al., 2013; John et al., 2011).
Additionally, many microalgae species are oleaginous in nature and thus could also be
exploited as the raw materials for biodiesel production (Chisti, 2007). Microalgae have
exhibited several advantages over traditional feedstocks for renewable bioethanol
production. Compared to terrestrial plants, microalgae have high growth rate and
volumetric productivity. Harvesting cycle of microalgae is usually from 1 to 10 days
depending on different species, whereas crop feedstocks require half or one year. Since
microalgae do not have structural biopolymers such as hemicellulose and lignin, they are
capable of accumulating more energy storage materials in cells than most productive land

	
  

	
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plants, and severe pretreatment steps required for the second-generation bioethanol
production can be eliminated. In addition, microalgae using CO2 as the carbon source not
only mitigate greenhouse emission, but also enhance global carbon circle and further
reduce the carbon footprint of biofuel production. These features apparently demonstrate
that microalgae are an ideal alternative feedstock for the development of third-generation
bioethanol (John et al., 2011).
1.3 Challenges in microalgae cultivation for carbohydrate accumulation
Many factors can influence the growth of photosynthetic microalgae, including light
intensity, carbon dioxide, inorganic salts, temperature and pH. How to optimize and
efficiently integrate these factors to achieve the processes with high productivity and costeffective cultivation is critical to commercializing microalgal biomass production.
1.3.1 Reactor configuration
Two major cultivation configurations of open pond and photobioreactor as discussed
as follow are being intensively studied to produce algal biomass at large scales. The Open
pond system simulates natural environment where microalgae growth occurs The open
pond system includes three basic types of raceway pond, circular pond and sloped pond
(John et al., 2011; Ugwu et al., 2008). Among them, raceway open pond is the most
common system for algal cultivation due to its characteristics of low running cost,
continuous mixing, and relatively simple to scale up. It has also been reported that the
open pond system is cheaper to build and easier to operate compared to closed
photobioreactor system (Shen et al., 2009). Besides these advantages, the open pond
system has some limitations for biofuel production as well, such as poor light availability
to cells, water loss due to evaporation, low concentration of dissolved CO2, contamination

	
  

	
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issues, lower biomass productivity, as well as requirement of a large amount of water
(John et al., 2011; Ugwu et al., 2008).
The limitations of the open pond system lead to increasing attention on the
development of enclosed photobioreactors such as tubular, flat-plate and vertical-column
photobioreactors. Flat-plate photobioreactors are usually made of transparent materials to
provide large illumination surface area. They have short light path and are good for algal
immobilization. Nevertheless, there are some limitations such as algal biofilm formed on
the walls and difficulty for scale-up (Ugwu et al., 2008). Tubular photobiorectors also
have large illumination surface area. With pumping system, microalgae culture can be recirculated and highly turbulent flow could prevent biomass sedimentation in tubes. During
the operation, due to the fact that oxygen generated from photosynthesis is built up in the
tubes and would cause algal cell damage under intense sunlight, the culture must be
degassed periodically and the degassed device has to be installed on the tube (Chisti,
2007). Compared to tubular and flat-plate photobioreactors, vertical-column bioreactor is
the most common system for microalgal culture. Vertical-column photobioreactor is
compact, economic, simple and suitable for large-scale cultivation. The only major
drawback for the vertical-column photobioreactor is the illumination surface area. An
increase in the size of the vertical-column photobioreactors has to sacrifice the
illumination-surface-to-volume ratio. Many efforts have been paid to increase illumination
surface area, such as designing internally illuminated photobioreactor (Ugwu et al., 2008)
and using optic fibers to enhance intensity and distribution of solar light (Matsunaga et al.,
1991).

	
  

	
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1.3.2 Cultivation techniques for microalgal carbohydrate accumulation
Photosynthetic microalgae absorb solar energy to split the water into protons, electrons
and oxygen. The electrons and protons are then used to synthesize energy carriers of
NADPH and ATP that drive the reduction of fixed carbon dioxide to carbohydrates by the
Calvin cycle (Markou et al., 2012). The functions of synthesized carbohydrates can
generally be classified into two types: structural components in the cell walls and storage
components mainly in the chloroplasts. The composition of these carbohydrates varies
regarding not only their functions but also different algal species. For instance,
cyanobacteria, red algae, and green algae use glycogen, combined starch and glycogen,
and starch, respectively, as their storage compounds (Markou et al., 2012; Nakamura et al.,
2005). Table 1 lists the major carbohydrates in selected microalgae, and shows that the
most abundant carbohydrates in green microalgae Chlamydomonas reinhardtii are glucose
polymers (starch) (Choi et al., 2010; Markou et al., 2012).
Table 1 Carbohydrates of selected microalgal species (adopted from Markou et al.,
2012)(Markou et al., 2012)

1.3.2.1 Nutrient control
Algal carbohydrate accumulation depends on both microalgal species and culture
conditions. Many studies have elucidated that depletion of some essential nutrients such as
	
   6	
  
	
  

	
  
	
  
nitrogen, phosphorous and sulfur can increase algal carbohydrate content (Dragone et al.,
2011; Ho et al., 2013a; Markou et al., 2012; Philipps et al., 2012; Siaut et al., 2011a).
Among then, nitrogen, as the basic element in the synthesis of vital compounds such as
proteins, coenzymes, DNA, RNA, and photosynthetic pigments, can count to around 10 %
of dry biomass weight in most microorganisms, which is much higher than phosphate and
sulfur. Nitrogen starvation is known to have a fast and strong influence on cell
composition. Philipps et al. reported that nitrogen deprivation leads Chlamydomonas to
accumulate more starch than does sulfur deprivation (Philipps et al., 2012). Table 2 shows
some green microalgal carbohydrate accumulation under nitrogen depletion condition.
Table 2 Carbohydrate content of green microalgae species in N-deprivation
Microalgae species

Carbohydrate (%)

References
(Branyikova

Chlorella vulgaris

38
et al., 2011)
(Dragone et

Chlorella vulgaris P12

41
al., 2011)
(Siaut et al.,

Chlamydomonas reinhardtii

40
2011a)
(Ji et al.,

Tetraselmis subcordiformis

35
2011)
(D'Souza &

Tetraselmis suecica

57

Kelly,
2000)
(Sassano et

Spirulina platensis

55-65
al., 2010)

	
  

	
   7	
  

	
  
	
  
Besides nitrogen, CO2 as the unique carbon source plays a critical role in autotrophic
microalgal growth. CO2 concentration not only affects their growth rate but also
influences biomass composition. Although Xia and Gao claimed that carbohydrate
contents in C. pyrenoidosa and C. reinhardtii increased with the elevation of CO2
concentration (Chen et al., 2013; Xia & Gao, 2005), more researchers believe that limiting
CO2 is advantageous for carbohydrate accumulation in microalgae due to the CO2concentrating mechanism (Giordano et al., 2005; Izumo et al., 2007; Ramazanov et al.,
1994; Thyssen et al., 2001). In some autotrophic organisms, atmospheric CO2 and O2
compete with each other for the active sites of RUBISCO, in which CO2 is located in
inferior position because of relatively low affinity of RUBISCO for CO2 and low
concentration of atmospheric CO2. In coping with this situation, many algae and some
higher plants develop CO2-concentration mechanisms (CCMs) that can increase CO2
concentrations around RUBISCO (Giordano et al., 2005). When cells are grown in
ambient air, a high-affinity CCM is induced so that cells can concentrate dissolved
inorganic carbon around RUBISCO sites (Ramazanov et al., 1994). Meanwhile, an
increase in the starch sheath associated with the pyrenoid was observed from microalgal
cells grown with limited CO2 concentration for the reason that increasing starch sheath
might be helpful in hindering the efflux of CO2 from the pyrenoid (Giordano et al., 2005).
As a result, a significant improvement in carbohydrate accumulation was observed by
dropping CO2 concentration from 3 to 0.04 % (Izumo et al., 2007). Considering the effects
of both nitrogen and CO2 on algal carbohydrate accumulation, it would be interesting to
know the effects of combination of nitrogen deprivation and CO2 limitation on algal
carbohydrate accumulation and corresponding growth conditions. However, to our
knowledge, such studies haven’t yet been reported.

	
  

	
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1.3.2.2 Cultivation strategies
As aforementioned, applying nutrient starvation/limitation is an effective approach for
the production of microalgae rich in carbohydrate. However, nutrient deprivation is
always associated with decreased growth rate and biomass productivity, and finally poor
carbohydrate productivity. Engineering solutions are urgently needed to address this
problem. Hsieh et al (Hsieh & Wu, 2009) optimized initial nitrogen concentration for the
microalgal oil production to balance cell growth and lipid productivity. Moreover, they
applied semi-continuous process to maintain the continuous culture. Maeda et al adopted
semi-continuous operation by alternatively feeding medium with two different nitrogen
concentrations to control turbidity and carbohydrate accumulation from Chlamydomonas
cultures (Maeda et al., 2006). Although these strategies are relatively easy to operate and
are able to maintain the cell growth while cells are accumulating storage compounds
(starch or lipids), they require longer culture time to acquire both cell growth and
carbohydrate accumulation. An alternative strategy is two-stage culture where nutrientrich medium is applied in the first stage to promote cell growth, and nutrient-deprived
medium is then used in the second stage to enhance carbohydrate accumulation (Ho et al.,
2010). Two-stage cultivation strategy has been used on a few microalgal strains such as
Scenedesmus obliquus CNW-N, Nannochloropsis oculata and C. reinhardtii for
carbohydrate and lipids accumulation (Ho et al., 2010; Ho et al., 2013c; Siaut et al., 2011b;
Su et al., 2011).
1.4 Microalgal carbohydrate conversion into fermentable sugars
In order to produce bioethanol from algal carbohydrates, the carbohydrates must be
converted into fermentable sugars that can be utilized by ethanol producing strains such as
yeast Saccharomyces serevisiae. Various methods have been applied to produce sugars

	
  

	
   9	
  

	
  
	
  
from microalgal biomass, and these methods could be categorized into two major groups:
chemical saccharification and enzymatic saccharification (Chen et al., 2013).
1.4.1 Chemical saccharification
Acid / alkali hydrolysis has been wildly applied in the pretreatment of lignocellulosic
biomass where the carbohydrates are mainly composed of cellulose and hemicellulose
linked with lignin (Ruan et al., 2013). Considering that microalge do not have lignin and
their carbohydrates are mainly in the form of starch, it is relatively easier to hydrolyze
microalgal carbohydrates compared to hydrolysis of lignocellulosic materials. Chemical
hydrolysis could thus be carried out under relatively mild conditions, such as lower
temperature and less duration time (Chen et al., 2013). For example, C. reinhardtii UTEX
90 has been utilized as feedstock for bioethanol fermentation and maximum 58 % (w w-1
algal biomass) glucose was released from the biomass after dilute sulfuric acid
pretreatment at 110 °C for 30 min (Nguyen et al., 2009). However, by comparing dilute
acid pretreatment and enzymatic pretreatment of C. reinhardtii UTEX 90 biomass, Seung
et al. found sugar yield from chemical saccharification process was still lower than that
from enzymatic saccharification (Chen et al., 2013; Choi et al., 2010; Nguyen et al., 2009).
In addition, low pH of the algal acid hydrolysate require neutralization operation in order
to enable yeast to carry on next step of ethanol fermentation, which makes acid
pretreatment not a ideal solution for algal carbohydrate hydrolysis.
1.4.2 Enzymatic saccharification
An alternative approach to obtain fermentable sugars from microalgae is enzymatic
saccharification. Different enzymes are used to carry on enzymatic saccharification
according to different carbohydrate composition of microalgae, such as microalgal
cellulose can be hydrolyzed by employing endo-β-1,4-D-glucanase, exo- β-1,4-D	
  

	
  10	
  

	
  
	
  
glucanase and β-glucosidase; microalgal starch can be hydrolyzed by amylase and
glucoamylase (Chen et al., 2013). It is apparent that enzymatic hydrolysis of microalgal
biomass exhibits several advantages over chemical treatment, such as higher sugar yield,
less by-products, and mild reaction conditions and environmental friendly (Choi et al.,
2010). However, in order to make the enzymes accessible to their target substrates and
increase hydrolysis efficiency, mechanical disruption of algal cells is often applied before
enzymatic hydroysis. Moreover, high temperature treatment of disrupted algal biomass is
necessary to gelatinize starch for the enzymatic hydrolysis. Harun and Danquah employed
a Vibra-cell ultrasonicator to rupture Trichoderma reesei culture and then used cellulase to
treat ruptured cells to release cellobiose and glucose (Harun & Danquah, 2011). Ho et al.
autoclaved microalgal biomass at 121 °C for 20 min and then mixed pretreated biomass
with fermentation broth of Pseudomonas strain containing a mixture of cellulases and
amylases to carry out enzymatic hydrolysis (Ho et al., 2013a). Both mechanical disruption
and autoclave require a significant amount of energy input and make the process a bit
complicated. Besides, enzyme cost is also an important factor for microalgal bioethanol
production.
1.4.3 “Self-processing” saccharification
Expression of heterologous amylases into algae enabling carbohydrate degradation in
vivo and releasing mono-sugars out of the cells without mechanical disruption could be
effective and economical approach to address the issues related with enzymatic
saccharification of algal biomass. Several studies have reported the successful expression
of heterologous amylases in corn and rice kernels to facilitate starch degradation. Pen et
al. (Pen et al., 1992) first reported that a bacterial α-amylase (from Bacillus licheniformis)
was expressed in tobacco, and the enzyme in the tobacco seeds was applied to liquefy

	
  

	
  11	
  

	
  
	
  
potato and corn starch. DE values of 16 and 13 were achieved from potato starch and corn
starch respectively, which exhibited the feasibility of producing an active industrial
enzyme in plants and replacing commercial enzymes for starch hydrolysis. Xu et al. (Xu et
al., 2008) introduced an α-amylase gene from Bacillus stearothermophilus into rice and
then applied the transgenic rice seeds in corn starch liquefaction without extraction and
enzyme purification. DE value of 17 for corn starch liquefaction was achieved after the
mixture was heated at 90 °C for 10 min and then incubated at 70 °C for 2 h. However, few
studies have focused on developing an amylase-producing microalga to facilitate
microalgal starch hydrolysis.
1.5 Recombinant protein expression in transgenic microalgae
Microalgae have been viewed as a potential platform for the recombinant protein
expression in these years due to the fact that microalgae exhibit all the advantages of
plants coupled with microbial features such as fast growth and high cell productivities
(Rosales-Mendoza et al., 2012; Surzycki et al., 2009; Walker et al., 2005). Over the last
decades, significant progress in development of transgenic microalgae has been achieved.
Several green microalgae such as C. reinhardtii and Dunaliella salina, and diatom
Phaeodactylum tricornutum have been intensively studied (Walker et al., 2005). Among
them, C. reinhardtii is the model microalga with complete chloroplast and nuclear
genomes. In addition, a wide range of chloroplast and nuclear promoters and their 5’ and
3’ UTRs in C. reinhardtii have been characterized, and various transformation
methodologies for its chloroplast and nuclear genomes have been developed (Boynton &
Gillham, 1993; Kindle, 1990; Kindle et al., 1991b).

	
  

	
  12	
  

	
  
	
  
1.5.1 Nuclear transformation
The Chlamydomonas nuclear genome has around 18 chromosomes containing 108 base
pairs. Like all eukaryotic genomes, the ability to perform post-transcriptional and posttranslational modification makes Chlamydomonas nuclear expression a preferred platform
for some complicated protein expression (Merchant et al., 2007). Nuclear expression
enables recombinant proteins to be targeted, attached and displayed on cell membrane, so
that the recombinant proteins can serve as vaccine to induce immune response (Kumar et
al., 2013). The introduction of heterologous DNA into nuclear genome can be achieved by
various approaches, such as biolistic bombardent with DNA-coated microprojectiles,
electroporation, as well as Cre/lox recombination system (Heitzer & Zschoernig, 2007;
Kindle, 1990; Kindle et al., 1989; Specht et al., 2010). Nevertheless, the most popular and
simplest method is to agitate cell-wall-deficient cells and DNA solutions with glass beads
(Kindle, 1990). As for the selection of transformants, wild type genes including NIT1
encoding nitrate reductase and ARG7 encoding argininosuccinate lyase have been used as
selectable marker to rescue auxotrophic mutants (Walker et al., 2005). Bacterial antibioticresistance genes, such as ble gene conferring zeocin resistance (Stevens et al., 1996) and
aadA gene conferring resistance to spectinomycin (Cerutti et al., 1997), have also been
applied for the selection of transformants. Except for these reporter proteins, several other
proteins from diverse sources have been expressed successfully and were found to be
functional, such as human glycoprotein erythropoietin (Epo) and AHL lactonase enzyme
(Siripornadulsil et al., 2002).
In spite of substantial progress, a major obstacle of developing Chlamydomonas
nuclear expression system is posed by the poor expression of foreign proteins from
nuclear genome. The possible reasons for such poor expression are: 1) Many genes from

	
  

	
  13	
  

	
  
	
  
bacteria are AT rich, while nuclear genome of Chlamydomonas is GC rich, therefore
codon usage may limit transcription of heterologous genes in nuclear genome (Merchant
et al., 2007); 2) Transgene silencing that is assumed to be evolved as a protective measure
against intracellular pathogens or viruses leads to instability of exogenous protein products
(Specht et al., 2010); and 3) Heterologous promoters that are commonly used in transgene
expression of plants result in insufficient transcription (Tang et al., 1995).
1.5.2 Chloroplast transformation
Compared to nuclear transformation, gene expression in chloroplast genome
exhibits many advantages and has gained much attention in these years. The ability to
integrate heterologous genes into specific sites on the chloroplast genome through
homologous recombination can enhance expression stability and avoid uncertain influence
on chloroplast gene functions (Kumar et al., 2013). Directed-site integration also enables
multiple-gene transformation into multiple insertion sites or one site by constructing
polycistronic transcripts (Specht et al., 2010). Due to the lack of gene silencing
mechanisms, much higher expression level can be acquired in chloroplast expression
system than nuclear expression system (Specht et al., 2010). Without post-translational
modification, proteins produced within the chloroplast will not be glycosylated, which is
preferred for the production of native recombinant antibodies (Franklin & Mayfield, 2005;
Tran et al., 2009). Furthermore, the features of Chlamydomonas chloroplast gene
expression, including the AT-rich codon bias, and eubacterial RNA polymerase and 70S
ribosome, demonstrated its prokaryotic nature, which make Chlamydomonas chloroplast a
suitable expression system especially for bacterial genes (Purton, 2007). Various reporter
proteins, recombinant vaccines, fully functional antibodies, as well as industrial enzyme

	
  

	
  14	
  

	
  
	
  
proteins have been successfully expressed in the Chlamydomoans chloroplast (Table 3)
and considerable progress has been made in the following aspects.
Table 3 Recent successes in recombinant protein production in C. reinhardtii
chloroplast
Gene expressed

Application

Reference

aadA

Reporter of gene expression

(Goldschmidt-Clermont, 1991)

uidA

Reporter of gene expression

(Ishikura et al., 1999)

gfp

Reporter of gene expression

(Franklin et al., 2002)

luxCt

Reporter of gene expression

(Mayfield & Schultz, 2004)

HSV8-1sc

Pharmaceutical

(Mayfield et al., 2003)

CTB-VP1

Vaccine

(Sun et al., 2003)

HSV8-scFv

Pharmaceutical

(Mayfield & Franklin, 2005)

hMT-2

Pharmaceutical, UV-protection (Zhang et al., 2006)

M-SAA

Therapeutics, oral delivery

(Manuell et al., 2007)

CSFV-E2

Vaccine

(He et al., 2007)

hGAD65

Diagnostics and therapeutics

(Wang et al., 2008)

83K7C

Therapeutics

(Tran et al., 2009)

IgG1

Therapeutics

(Tran et al., 2009)

VP28

Vaccine

(Surzycki et al., 2009)

CTB-D2

Oral Vaccine

(Dreesen et al., 2010)

M-SAA-Interferon β1 Therapeutics

	
  

(Rasala et al., 2010)

	
  15	
  

	
  
	
  
1.5.2.1 Codon optimization
It is well known that codon biases vary from genomes of different organisms, or even
different genomes of the same organisms (Potvin & Zhang, 2010). The C. reinhardtii
chloroplast genome exhibits prokaryotic nature, and codon bias is one of the most critical
factors of protein expression (Surzycki et al., 2009). It was reported that translation rates
of microalgal-destined transgenes could be increased if they share the same codon bias
with microalgal genes (Heitzer & Zschoernig, 2007). Codon optimization combined with
gene synthesis has been viewed as an effective and necessary step to acquire high-level
expression in Chlamydomonas chloroplast. Franklin et al. reported that accumulation of
green fluorescent protein (GFP) in Chlamydomonas chloroplast increased more than 80fold through codon optimization (Franklin et al., 2002).
1.5.2.2 Promoters
Numerous studies have been done to identify the 5’-UTR and 3’-UTR sequences of
endogenous genes in Chlamydomonas chloroplast, as well as their functions on gene
expression. The 3’-UTR regions were found to be less important for gene expression than
5’-UTR, and changing the 3’-UTR had little impact on translation and transcription, as
long as the 3’-UTR was present (Barnes et al., 2005). Whereas, regions within the 5’-UTR
can not only control the translation but also dramatically affect mRNA stability (RosalesMendoza et al., 2012). Moreover, some studies claimed that 5’-UTRs play a critical role in
translational regulation that is a rate-limiting step for chloroplast gene expression
(Rosales-Mendoza et al., 2012). Therefore, influence of different endogenous genes 5’UTR regions on heterologous gene expression have been studied. Goldschmidt-Clermont
(Goldschmidt-Clermont, 1991) constructed different expression cassettes to express aadA
gene coding for aminoglycoside 3’ adenyl transferase in C. reinhardtii chloroplast. Results

	
  

	
  16	
  

	
  
	
  
demonstrated that specific activity of AAD was higher when the expression of aadA gene
was controlled by 5’- UTR of atpA gene than 5’-UTR of rbcL gene. Barnes et al. in 2005
assessed the expression of the gfp reporter gene in Chlamydomonas chloroplast under
various endogenous promoters, and concluded that 5’-UTR of atpA and psbD genes led to
the higher levels of chimeric mRNA and protein accumulation when compared to the 5’UTR of rbcL and psbA genes (Barnes et al., 2005). Interestingly, it was found that the
psbA promoter had good potential to control the heterologous gene expression when the
endogenous psbA gene was replaced (Manuell et al., 2007). However, the absence of psbA
gene product, which is an important protein in photosystem II, will disrupt photosynthesis
function, and re-introduction of psbA gene into chloroplast genome will increase the
complexity of the whole process. As shown in table 4, Rosales-Mendoza et al. have
summarized the most popular endogenous promoters for exogenous gene expression in
Chlamydomonas chloroplast (Rosales-Mendoza et al., 2012).

Table 4 Chloroplast promoters of C. reinhardtii used for the expression of
heterologous proteins (adopted from (Rosales-Mendoza et al., 2012))

	
  

	
  17	
  

	
  
	
  
1.5.2.3 Transformation methods
Unlike nuclear transformation, transformation of chloroplasts requires the delivery of
exogenous genes through both the cellular and chloroplast membrane. Some researchers
applied popular nuclear transformation methods including glass beads agitation and
electroporation to deliver DNA in the chloroplast, and only acquired very low efficiency
of chloroplast transformation (Kindle et al., 1991a; Purton, 2007). Considerable progress
in the chloroplast transformation has been made since Boynton et al. employed highvelocity microprojectiles to deliver DNA into the chloroplast compartment (Boynton et al.,
1988). During the biolistic process, the DNA-coated tungsten or golden microparticles are
accelerated under high helium pressure and then shot into cells plated on solid medium
(Gan, 1989). The kinetic energy of these microparticles is sufficient to penetrate both the
cellular and chloroplast membranes and deliver DNA molecules into the organelle. It was
reported that microparticle bombardment can produce103 transformants per µg DNA,
which is at least ten times more efficient than agitation with glass beads (Kindle &
Sodeinde, 1994). Although this method requires specialized equipment, with gold
microparticles being more expensive, microparticle bombardment has been viewed as the
most effective approach for the transformation of chloroplasts in C. reinhardtii. (Franklin
et al., 2002; Goldschmidt-Clermont, 1991; Kindle & Sodeinde, 1994; Rasala et al., 2010).
1.6 Goal and Objectives
Due to the high growth rate, high photosynthetic efficiency, and relatively high
content of energy-rich cell components, microalgae as the potential non-food-based
feedstock for bioethanol production have gained a lot of attention. As aforementioned,
great research progress has been made in improving bioethanol production from
microalgal carbohydrate. However, several barriers still need to be overcome to make

	
  

	
  18	
  

	
  
	
  
microalgal-bioethanol commercially viable: 1) productivity of microalgal carbohydrate is
still low; 2) chemical pretreatment of microalgal biomass needs high temperature, high
pressure and a large amount of chemicals; and 3) enzymatic pretreatment of microalgal
biomass requires relatively expensive enzymes and complicated mechanical disintegration.
Therefore, the project goal is to develop a novel and highly efficient process for
microalgal carbohydrate production and utilization. The objectives of this project are as
follow:
•

Optimize culture condition and operation mode to improve the microalgal biomass
productivity

•

Investigate the influence of nitrogen depletion combined with CO2 limitation on
microalgal carbohydrate accumulation

•

Construct a novel process to facilitate microagal carbohydrate utilization by using
a “self-processing” microalga.

	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  

	
  19	
  

	
  
	
  
CHAPTER 2 TWO-STAGE PHOTOAUTROPHIC CULTIVATION TO IMPROVE
CARBOHYDATE PRODUCTION IN CHLAMYDOMONAS REINHARDTII
	
  
Xiaoqing Wang, Zhenhua Ruan, Patrick Sheridan, Danielle Boileau, Yan Liu*, Wei Liao
2.1 Abstract
A two-stage strategy was developed to improve the microalgal carbohydrate
production for advanced biofuel production. In the first stage, model microalga C.
reinhardtii CC125 was cultivated in a photo-bioreactor on two different operation modes
to improve the biomass production. Biomass productivity was increased from 0.23 g L-1 d1

to 0.47 g L-1 d-1 through the semi-continuous operation with 70 % medium replacement

ratio under 5 % CO2. In the second stage, reducing CO2 content to 0.04 % led to a high
carbohydrate content of 64.5 %, showing more than 8 times improvement compared to
that in the biomass from the first stage culture. These results suggested that
photoautotrophic two-stage cultivation was an effective approach to accumulate
microalgal carbohydrates as a feedstock for biofuel production.
2.2 Introduction
Microalgae as one of the most promising non-food-crop-based feedstocks exhibit
several advantages over terrestrial energy crops for next-generation biofuel production.
They can grow on marginal lands and use wastewater as nutrient sources, thereby
avoiding competition with food crops on arable land and reducing freshwater
consumption. Photosynthetic microalgae are capable of fixing CO2 per unit area severalfold more than terrestrial plants, which leads to an annual biomass yield (kg biomass/acre)
about two or three orders of magnitude greater than corn or soybean (Huntley & Redalje,
2007). The algal biomass rich in carbohydrate and lipid has great potential to produce

	
  

	
  20	
  

	
  
	
  
biofuels and chemicals (Chen et al., 2013; Ho et al., 2013c; Markou et al., 2012).
Currently, majority of microalgae research and development have been focused on lipid
accumulation for biodiesel production (Cakmak et al., 2012; Chisti, 2007; Halim et al.,
2012; Siaut et al., 2011a). The other energy component in microalgae, carbohydrate, has
been largely overlooked. Many microalgae prefer to accumulate carbohydrate as structural
components in the cell wall and energy storage in the plastids (Chen et al., 2013; Markou
et al., 2012). Compared to lignocellulose, microalgae do not have lignin, and have higher
starch and less cellulose and hemicellulose contents (Chen et al., 2013). Therefore, the
microalgal carbohydrates could be a good substrate for the production of methane,
bioethanol and biobutanol (Markou et al., 2012; Yao et al., 2013).
Nutrient depletion has been accepted as the most common approach to increase algal
carbohydrate content. However, microalgal growth slows down under the stressed
environmental conditions, and corresponding carbohydrate production is low (Markou et
al., 2012). Developing effective culture strategies to realize both carbohydrate
accumulation and biomass growth is critical to increase efficiency of microalgal
carbohydrate production, which could make major contribution towards microalgal
carbohydrate based biofuel production (Dragone et al., 2011). Nutrient limitation strategy
is considered to be a feasible approach for the production of microalgae rich in
carbohydrates through limiting and controlling the concentration of key nutrients instead
of omitting the nutrients from the beginning of batch cultivation (Markou et al., 2012). For
example, Maeda et al developed a semi-continuous cultivation method which could
control turbidity and dilution rate by feeding medium with two different nitrogen
concentrations alternatively to evaluate starch production in Chlamydomonas sp (Maeda et
al., 2006). However, it is still difficult to achieve the highest growth rate and biomass

	
  

	
  21	
  

	
  
	
  
production under nutrient limitation. To handle this situation, a strategy of two-stage
culture was applied, in which the microalgae was first cultivated using nutrient-rich
medium to enhance cell growth and then switched to stressed condition to trigger
carbohydrate accumulation (Ho et al., 2013c).
In this study, Chlamydomonas reinhardtii, as one of the most extensively studied
unicellular green microalgae and a model organism for the study of photosynthesis
function, as well as the one which is capable to accumulate a large amount of
carbohydrate, mainly starch, for biofuel production (Choi et al., 2010), was used to
develop a highly efficient microalgal carbohydrate production system. A two-stage
cultivation approach was applied, in which culture conditions were optimized and semicontinuous culture was conducted to maximize biomass productivity in the first stage and
nitrogen was depleted to promote carbohydrate production in the second stage. Moreover,
the influence of CO2 concentration in mixed air on the carbohydrate accumulation in
related with CO2 concentrating mechanism (CCM) was also discussed in the second stage.
Finally, ethanol fermentation by Saccharomyces cerevisiae using microalgal biomass
hydrolysates was evaluated.
2.3 Methods
2.3.1 The microalga strain and growth medium
C. reinhardtii CC-125 mt+ (137c) was the microalga strain for this study. Solid TrisAcetate-Phosphate (TAP) medium (Harris, 1989) was used to store the strain, and liquid
TAP medium was used for seed culture. In photoautotrophic culture, strain was cultured in
the liquid Sueoka's high salt (HS) medium (Sueoka, 1960) for biomass growth and
transferred into HS depleting N (HSN) medium for carbohydrate accumulation.

	
  

	
  22	
  

	
  
	
  
2.3.2 Experimental setup
All experiments were performed in the 500 mL photobioreactors (PBRs) with 300 mL
loading volume. The PBRs were operated at room temperature (around 25 °C) under
continuous light with an agitation rate of 200 rpm. Sterilized mixed air with different CO2
concentrations (v v-1) were supplied into the liquid culture in the PBRs with fixed flow
rate of 0.7 vvm.
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  

	
  

	
  23	
  

	
  
	
  
2.3.2.1 Optimization of culture conditions in batch operation
In order to reach the highest growth rate during the exponential phase, a completely
randomized design (CRD) was used to analyze three factors of CO2 concentration (10 %,
5 % and 0.04 %, v v-1), light intensity (10 klux and 5 klux) and initial NH4+ concentration
(4.5 mM, 9 mM and 18 mM) (Table 5).
Table 5 Worksheet for the 18-run, 3-factor CRD optimization experiment

	
  

No.
1

Light intensity (klux)
5

NH4+ concentration (mM)
4.7

CO2 content (%)
Air

2

5

4.7

5

3

5

4.7

10

4

5

9.4

Air

5

5

9.4

5

6

5

9.4

10

7

5

18.8

Air

8

5

18.8

5

9

5

18.8

10

10

10

4.7

Air

11

10

4.7

5

12

10

4.7

10

13

10

9.4

Air

14

10

9.4

5

15

10

9.4

10

16

10

18.8

Air

17

10

18.8

5

18

10

18.8

10

	
  24	
  

	
  
	
  
Eighteen treatments with two replicates were applied for a total of 36 individual samples.
C. reinhardtii CC-125 mt+ (137c) was inoculated into each sample with initial cell
number around 3 × 105 cells mL-1. Samples were taken during the culture to monitor pH
and determine biomass concentration. Biomass was spun down using a Beckmann AR-12
® centrifuge at 8,000 g for 10 min, washed twice using deionized water and then dried at
70 °C overnight to estimate dry biomass weight. The optimal culture condition was
determined based on the specific growth rate and biomass productivity.
2.3.2.2 Operation of semi-continuous cultivation in Stage I
The semi-continuous cultivation was carried out with 5 % CO2 aeration with initial
biomass concentration around 0.6 g L-1. Different volumes of the culture broth were
replaced with fresh HS medium. Before and after medium replacement, liquid samples
were collected to measure dry biomass weight and pH. Optimal medium replacement ratio
was selected based on the highest biomass productivity. The biomass collected from the
replaced broth was washed twice with deionized water and then transferred to the stage II
cultivation. Carbohydrate and protein contents of the collected biomass were analyzed as
well.
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  

	
  

	
  25	
  

	
  
	
  

2.3.2.3 Stage II of batch culture for carbohydrate accumulation
Biomass harvested from the semi-continuous cultivation was transferred into HSN
medium for inducing carbohydrate synthesis. The effect of carbon dioxide content on
carbohydrate accumulation was investigated under 5 % and 0.04 % (v v-1) CO2,
respectively. Samples were collected during the culture to monitor the changes of biomass
concentration, cell number, carbohydrate, and protein contents. The scheme of two-stage
cultivation was shown in Fig.2.

Figure 2 Schematic representation of the two-stage microalgal cultivation

	
  

	
  26	
  

	
  
	
  
2.3.3 Ethanol fermentation
Hydrolysate of microalgal biomass harvested from the 2nd cultivation stage was
centrifuged at 8,000 g for 10 min to remove the cell debris. The aerobically pre-cultured
cells of yeast S. cerevisiae were centrifuged at 10,000 rpm for 10 min and then added into
the solution of hydrolysate. Fermentation was conducted at 30 °C until all the sugars were
consumed (Ho et al., 2013b).
2.3.4 Analytical methods
2.3.4.1 Cell growth measurement
Cell mass was separated from the culture by centrifugation at 8,000 g for 10 min, and
then washed with deionized water twice. Cell pellets were then dried overnight and
weighted to calculate dry biomass concentration.
The specific growth rate (µ, d-1) was determined from the dry biomass concentration
(g L-1) in the time interval of exponential phase, as shown in Eq. (1):
µμ = (𝐿𝑛(𝑋! ) − 𝐿𝑛(𝑋! ))/(𝑡! − 𝑡! )

(1)

Where X2 and X1 are the dry biomass concentration at times t2 and t1, respectively.
The biomass productivity (g L-1 d-1) was calculated by the Eq. (2):
𝑃! = (∆𝑋)/∆𝑡

(2)

Where Px is the biomass productivity, ∆X is the change of dry biomass concentration in
the cultivation time ∆t.
2.3.4.2 Carbohydrate, protein and ethanol analysis
Modified quantitative saccharification method from National Renewable Energy
(NREL) (Moxley & Zhang, 2007) was used to determine carbohydrate content in
microalgae. A small amount of biomass was suspended in 72 % (w w-1) H2SO4 solution

	
  

	
  27	
  

	
  
	
  
and incubated at room temperature for 1 hour. Deionized water was then added to the
mixture to dilute the H2SO4 concentration from 72 % (w w-1) to 4 % (w w-1). Mixture was
then incubated at 121 °C for 1 hour. After incubation, the hydrolysate was neutralized
with CaCO3 and analyzed using 3,5-Dinitrosalicylic acid (DNS) method for the sugar
assays (Miller, 1959).
For crude protein test, microalgal biomass was broken by sonication (Ultrasonic
Liquid Processors, Qsonica, LLC., Newtown, CT) in a buffer solution containing 100 mM
Bis-Tris-Propane (pH 7.0). Concentration of crude protein was then tested following BCA
kit analysis method (Ruan et al., 2012).
The concentration of the produced ethanol was detected by HPLC as described by
Ruan, et al. (Ruan et al., 2012). Agilent 1100 was equipped with a Bio-rad Aminex HPX87H and a refractive index detector. 5 mM sulfuric acid was used as the mobile phase and
the column temperature was set to 65 °C. 200 proof, HPLC grade ethyl alcohol was used
as standard.
2.3.4.3 Statistical analysis
A general linear model using R software (R Version 2.15.0, Vienna, Austria) was
applied to perform an analysis of variance (ANOVA) and multiple comparisons on the
experimental data. The significance of individual factors including light intensity, NH4+
concentration as well as CO2 concentration and their interactions on specific growth rate
and biomass productivity, was also identified using ANOVA analysis as described by
Ruan et al (Ruan et al., 2013).

	
  

	
  28	
  

	
  
	
  
2.4 Results and discussion
2.4.1

Optimization of culture conditions in batch operation

Continuous illumination and CO2 bubbling can produce rapid and logarithmic growth
of microalgae in liquid culture. Additionally, microalgae growth also relies on an
sufficient supply of essential macronutrient elements such as nitrogen (Harris, 1989).
Therefore in the present study, we examine the growth response of C. reinhardtii CC-125
mt+ (137c) to physical control by light intensity, CO2 concentration in the mixed air and
initial NH4+ concentration, in order to maximize growth rate and biomass production. The
experimental results obtained by the cultivation of C. reinhardtii under different cultural
conditions are shown in Table 6, and growth curves are exhibited in Fig 3. Cell growth
was repressed with 0.04 % CO2 (ambient air), whereas increase of CO2 concentration from
0.04 % to 5 % significantly enhanced growth rate. It was clear that specific growth rate,
final biomass concentration and biomass productivity were enhanced averagely by 56 %,
103 %, and 136 % respectively when compared to 0.04 % CO2 (Table 6). Further
increasing CO2 content to 10 % led to apparently growth inhibition. Specific growth rate,
final biomass concentration and biomass productivity at 10 % CO2 were dropped to an
average of 53 %, 25 % and 37 % respectively from those at 5 % CO2 (Table 6). Therefore,
the maximum specific growth rate of 0.1 d-1, 0.1 d-1 and 0.08 d-1, and highest biomass
productivity of 0.25 g L-1 d-1, 0.27 g L-1 d-1 and 0.26 g L-1 d-1, were achieved under 5 %
CO2 with NH4+ concentration of 4.7 mM, 9.4 mM and 18.8 mM (Table 5, 6), respectively.

	
  

	
  29	
  

	
  
	
  
Table 6 Response of 18-run CRD experiment design
Biomass
Specific growth rate
No.

Biomass concentration

-1

-1

(h )

(g L )

1

0.03 ± 0.003

0.40 ± 0.010

0.10 ± 0.003

2

0.03 ± 0.001

0.48 ± 0.030

0.14 ± 0.008

3

0.03 ± 0.003

0.15 ± 0.015

0.04 ± 0.005

4

0.03 ± 0.001

0.35 ± 0.010

0.10 ± 0.003

5

0.03 ± 0.003

0.53 ± 0.001

0.15 ± 0.008

6

0.03 ± 0.005

0.17 ± 0.005

0.05 ± 0.002

7

0.03 ± 0.005

0.34 ± 0.010

0.10 ± 0.003

8

0.04 ± 0.005

0.48 ± 0.010

0.14 ± 0.006

9

0.03 ± 0.009

0.15 ± 0.030

0.05 ± 0.009

10

0.07 ± 0.013

0.35 ± 0.035

0.10 ± 0.010

11

0.10 ± 0.016

0.74 ± 0.004

0.25 ± 0.001

12

0.05 ± 0.011

0.53 ± 0.017

0.15 ± 0.005

13

0.06 ± 0.005

0.41 ± 0.075

0.12 ± 0.021

14

0.10 ± 0.001

0.79 ± 0.034

0.27 ± 0.007

15

0.04 ± 0.015

0.58 ± 0.006

0.17 ± 0.002

16

0.05 ± 0.001

0.37 ± 0.020

0.11 ± 0.006

17

0.08 ± 0.003

0.77 ± 0.017

0.26 ± 0.006

18

0.04 ± 0.007

0.61 ± 0.006

0.17 ± 0.002

The data are average of two replicates with standard deviation

	
  

productivity

	
  30	
  

(g L-1 d-1)

	
  
	
  
When cultured under light intensity of 10 klux, almost the same growth modes were
achieved with NH4+ concentrations ranging from 4.7 mM to 18.8 mM (Fig 3A, 3B and
3C).

Biomass concentration g L-1

1

A"

0.8
0.6
0.4
0.2
0
0

20

40
60
Time (h)
Air

5% CO2

80
10% CO2

1
Biomass concentration g L-1

100

B"

0.8
0.6
0.4
0.2
0
0

20

40
60
Time (h)
Air

5% CO2

80

100

10% CO2

Figure 3 C. reinhardtii CC125 growth curve in batch culture under different culture
conditions: (A) light intensity of 10 klux and NH4+ concentration of 4.7 mM (B) 10
klux light intensity and 9.4 mM NH4+ concentration (C) 10 klux light intensity and
18.8 mM NH4+ concentration (D) 5 klux light intensity and 4.7 mM NH4+
concentration (E) 5 klux light intensity and 9.4 mM NH4+ concentration (F) 5 klux
light intensity and 18.8 mM NH4+ concentration.

	
  

	
  31	
  

	
  
	
  
Figure 3 (cont’d)

Biomass concentration g L-1

1

C"

0.8
0.6
0.4
0.2
0
0

20

40
60
Time (h)
Air

5% CO2

80
10% CO2

1
Biomass concentration g L-1

100

D"

0.8
0.6
0.4
0.2
0
0

20

40
60
Time (h)
Air

	
  

5% CO2

	
  32	
  

80
10% CO2

100

	
  
	
  
Figure 3 (cont’d)

Biomass concentration g L-1

1

E"

0.8
0.6
0.4
0.2
0
0

20

40
60
Time (h)
Air

5% CO2

80
10% CO2

1
Biomass concentration g L-1

100

F"

0.8
0.6
0.4
0.2
0
0

20

40
60
Time (h)
Air

5% CO2

80

100

10% CO2

Same with the light intensity of 10 klux, no significant difference of growth conditions
was observed between various NH4+ concentrations under light intensity of 5 klux (Fig 3D
to Fig 3F), whereas dropping light intensity from 10 klux to 5 klux obviously inhibited
cell growth.

	
  

	
  33	
  

	
  
	
  
Additionally, unlike the culture under higher light intensity (10 klux), no significant
differences of specific growth rate and biomass productivity could be observed under 5
lux light intensity with different CO2 contents, which means increase of CO2 supply from
ambient air level to 5 % can’t promote cell growth without sufficient light supplement.
ANOVA analysis also indicated that specific growth rate and biomass productivity were
dependent on the main factor of light intensity (P<0.05), but independent of NH4+
concentration and CO2 content (P>0.05). However, there were significant two-way
interaction between light intensity and CO2 content on cell growth (P<0.05). Additionally,
the broth pH under different culture conditions was quite stable, ranging from 6.5 to 7.0
without any control (data was not shown). The result demonstrated that culture condition
of 10 klux light intensity combined with 5 % CO2 was optimal for microalgal growth,
which was therefore selected in the section of semi-continuous cultivation.
2.4.2

Stage I of semi-continuous cultivation

Unlike heterologous microorganisms for which high-density cultivation has a positive
impact on production of storage metabolites, biomass accumulation of photosynthetic
microalgae is limited by the self-shading effect. Thus, semi-continuous operations with
different medium replacement ratios were applied for the cultivation of C. reinhardtii
CC125, in order to avoid the self-shading effect and maintain the growth in the
exponential phase as well as enhance biomass productivity.

5 % CO2 was applied in the semi-continuous operation since it exhibited the highest
specific growth rate and biomass productivity in the batch culture. C. reinhardtii CC125
were first cultivated in batch system for 35 hours to reach the mid-late exponential phase

	
  

	
  34	
  

	
  
	
  
and then switched to semi-continuous mode with four medium replacement ratios (45 %,
70 %, 83 % and 91 %, v v-1) every 24 hours. Fig. 4 shows biomass growth with different

Biomass Concentration g L-1

Biomass Concentration g L-1

medium replacement ratios for six repeated cycles.

1
0.8
0.6
0.4
0.2

45% replacement ratio

0
0

1

2

3
Time (d)

4

5

6

1
70% replacement ratio

0.8
0.6
0.4
0.2
0
0

1

2

3
Time (d)

4

5

6

Figure 4 Semi-continuous cultivation of C. reinhardtii CC125 under the optimal
condition obtained from batch tests. The culture broth was replaced by 45 %, 70 %,
83 % and 91 % with fresh medium in different semi-continuous tests, each of which
was operated for six repeated cycles. Data are average of two experimental
replicates.
	
  

	
  35	
  

	
  
	
  

Biomass Concentration g L-1

Figure 4 (cont’d)

1
91% replacement ratio

0.8
0.6
0.4
0.2
0

Biomass Concentration g L-1

0

2

3
Time (d)

4

5

6

1
83% replacement ratio

0.8
0.6
0.4
0.2
0
0

	
  

1

1

2

3
Time (d)

	
  36	
  

4

5

6

	
  
	
  
The data demonstrated that higher medium replacement ratios resulted in a lower biomass
concentration (Table 7).
Table 7 Performance of biomass production and carbohydrate content of semi-batch
culture of C. reinhardtii CC125 with different medium replacement ratio under
optimal conditions obtained from batch tests
Carbohydrate
Replacement ratio Biomass concentration Biomass productivity
(%)

-1

-1

(g L )

-1

content

(g L d )
(%, of total DCW)

45

0.81 ± 0.042

0.36 ± 0.019

7.11 ± 0.185

70

0.67 ± 0.051

0.47 ± 0.023

6.52 ± 0.081

83

0.54 ± 0.047

0.45 ± 0.021

7.66 ± 0.307

91

0.31 ± 0.035

0.28 ± 0.016

7.64 ± 0.613

The data are average of six repeated cycles with standard deviation.
While, biomass productivity increased from 0.36 to 0.47 g L-1 d-1 (P < 0.05) with the
increase of medium replacement ratio from 45 % to 70 %, and maintained the same
productivity (0.45 g L-1 d-1) at the replacement ratio of 83 % (P > 0.05). With further
increase of replacement ratio to 91 %, the biomass productivity dropped obviously to 0.28
g L-1 d-1 (P<0.05). The highest biomass productivity of 0.47 g L-1 d-1 reached at 70 %
replacement ratio in semi-continuous operation was doubled compared to that from batch
operation at 5 % CO2 (0.23 g L-1 d-1). Therefore, the culture conditions of 70 %
replacement ratio under 5 % CO2 was selected for Stage I cultivation for biomass
accumulation. However, carbohydrate analysis demonstrated that only a small amount of
carbohydrate was accumulated in C. reinhardtii biomass under the semi-continuous
culture conditions (with nitrogen source in the medium and high CO2 content in the mixed

	
  

	
  37	
  

	
  
	
  
gas), and there were no significant differences in carbohydrate content among different
medium replacement ratios (ranging from 6.52 % to 7.66 %, P > 0.05).

2.4.3

Stage II of batch culture for carbohydrate synthesis

Many studies investigated the effect of nitrogen starvation on carbohydrate synthesis
in C. reinhardtii (Ball et al., 1990; Cakmak et al., 2012; Gardner et al., 2013; Maeda et al.,
2006; Siaut et al., 2011a), and almost all of them cultivated C. reinhardtii under either
mixtrophic conditions using either acetate as organic carbon source or photoautotrophic
conditions with fixed CO2 concentration ranging from 2 % to 5 %. Limited attention has
been paid to the influence of different CO2 concentration on staged microalgal cultivation
for carbohydrate accumulation. CO2 affects microalgal physiology through the CO2concentrating mechanism (CCM). A high-affinity CCM is induced only in the
photoautotrophic culture when cells are grown at ambient air to concentrate dissolved
inorganic carbon inside the cells (Ramazanov et al., 1994). Ramazanov reported that
pyrenoid starch sheath in C. reinhardtii Dangeard was formed rapidly when supplied with
0.04 % CO2 (Ramazanov et al., 1994), and Izumo et al reported the increase of starch
amount in Chlorella cultured at ambient CO2 (Izumo et al., 2007). However, carbohydrate
accumulation in microalgal biomass cultured photoautotrophically with the depletion of
nitrogen and CO2 has not been fully analyzed yet. In this study, we cultured C. reinhardtii
CC125 in the HSN medium supplied with mixed air containing 0.04% and 5% CO2
respectively. Cell growth and composition were monitored during the cultivation.
Changes of cell density and dry biomass weight in Stage II were presented in Fig. 5.
For 5 % CO2, cell division was not inhibited in the first day during the nitrogen starvation,
which was probably due to continuous nitrogen supply from intracellular proteins. After

	
  

	
  38	
  

	
  
	
  
one-day starvation, nitrogen in the cells was depleted, cell division was inhibited, and the
corresponding growth was stopped. Different from the culture with 5 % CO2, cell division
under the 0.04 % CO2 condition was completely inhibited during the Stage II culture. It is
well known that common response of nitrogen deprivation is the inhibition of cell division
and accumulation of storage compounds (Wang et al., 2008). Cell density demonstrated
that decreasing CO2 concentration from 5 % to the ambient level further enhanced such
inhibition effect (Fig. 5). Unlike the cell density, dry biomass of the culture in both 0.04 %
and 5 % CO2 kept increasing from 0.56 to 1.03 g L-1 and 0.60 to 0.93 g L-1 respectively,
and even kept relatively high growth rates in the first three days during the whole
starvation process. Since there were no significant (P>0.05) difference on biomass
concentration between both cultures in Stage II cultivation, dry weight of a single cell in
the culture with 0.04 % CO2 was higher than that with 5 % CO2, considering less cell
numbers in the former culture.

Figure 5 Changes of biomass and cell density of C. reinhardtii CC125 during the
Stage II cultivation. Data are average of two experiment replicates

	
  

	
  39	
  

	
  
	
  
Changes of biomass composition during the nitrogen starvation are presented in Fig. 6.
Although both cultures with 0.04% and 5% CO2 increased carbohydrate content, C.
reinhardtii CC125 cultured with 0.04 % CO2 showed a more significant increase in
carbohydrate, sharper decrease in protein, and a slighter drop in other cell components
(including lipids) (Fig. 6b).
100"

5%"CO2"

Protein"and"carbohydrate"contents""(%)"

90"
80"
70"
60"
50"
40"
30"
20"
10"
0"
0"

1"

2"

3"

4"

5"

Time"(d)"
Protein"content"

Carbohydrate"content"

Others"

Protein"and"carbohydrate"contents"(%)"

100"

0.04%"CO2"

90"
80"
70"
60"
50"
40"
30"
20"
10"
0"
0"

1"

Protein"content"

2"
Time"(d)"

3"

Carbohydrate"content"

4"

5"

Others"

Figure 6 Biomass compositions of C. reinhardtii CC125 cultured with (A) 5 % CO2
and (B) 0.04 % CO2 in the Stage II cultivation. Data are average of two experimental
replicates.
	
  40	
  
	
  

	
  
	
  
The carbohydrate accumulation with 0.04% CO2 rapidly rose with the increased starvation
time, and reached the peak after 4-day starvation. The highest carbohydrate content and
concentration reached 64.5 % of the dry biomass and 0.66 g L-1 in the culture broth,
showing more than 8 and 16 folds improvemet, respectively, compared to those from the
Stage I cultivation. The resulting carbohydrate-rich microalgal biomass was then
hydrolyzed with an enzyme cocktail of alpha-amylase and amyloglucosidase as described
by Van Wychen and Laurens (Van Wychen & Laurens, 2013), and hydrolysate was then
used for ethanol fermentation by S. cerevisiae. The highest ethanol yield was 0.41 g
ethanol per gram glucose, which accounted for 81% of theoretical yield of glucose to
ethanol fermentation.
The carbohydrate accumulation in C. reinhardtii CC125 with 0.04 % CO2 in Stage II
is higher than other C. reinhardtii strains from the literature. It has been reported that C.
reinhardtii UTEX 90 can accumulate 60 % carbohydrate out of total DCW (Nguyen et al.,
2009), and 0.41 g L-1 carbohydrate was reached in the culture broth of Chlamydomonas sp.
MGA161 (Maeda et al., 2006). Figs. 5 and 6 demonstrated that depleting CO2 to low
levels during the nitrogen starvation can inhibit cell division and significantly enhance the
carbohydrate accumulation, mainly due to the development of pyrenoid starch from CCM
(Izumo et al., 2007). However, it is also possible that the response of algae to low CO2
concentration during nitrogen starvation is dependent on culture condition as well, since
Gardner et al reported that no starch accumulation was achieved in C. reinhardtii after the
aeration shift from 5 % CO2 to air during nitrogen depletion in the batch culture (Gardner
et al., 2013).

	
  

	
  41	
  

	
  
	
  
2.5 Conclusion
C. reinhardtii CC125 was selected and underwent a photoautotrophic two-stage
cultivation. In Stage I of semi-continuous operation, the biomass productivity of 0.47 g L-1
d-1) was achieved. Carbohydrate accumulation was then induced in Stage II of the batch
starvation operation. Low CO2 concentration during the nitrogen starvation substantially
improved carbohydrate accumulation. The highest carbohydrate content of 64.5% dry
weight and the highest carbohydrate concentration of 0.66 g L-1 were reached with 0.04 %
CO2 after 4 days starvation. The two-stage algae cultivation has been proven by the study
as an efficient strategy to accumulate a large amount of carbohydrate in microalgae.
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  
	
  

	
  42	
  

	
  
	
  
CHAPTER 3 TRANSGENIC EXPRESSION OF A BACTERIAL TRANSGENIC
EXPRESSION OF A BACTERIAL THERMOPHILIC AMYLASE IN THE
CHLAMYDOMONAS REINHARDTII CHLOROPLAST

Xiaoqing Wang, Zhenhua Ruan, Danielle Boileau, Barbara B. Sears, Yan Liu, Wei Liao
3.1 Abstract
The objective of this study was to develop a microalga capable of hydrolyzing its own
starch in vivo and overcome the barrier of releasing large molecules for algal fuel and
chemical production. A thermophilic amylase (arAmyBH) from hyperthermophilic
bacterium Thermotoga neapolitana was introduced into Chlamydomonas reinhardtii
chloroplast. Expression and accumulation of recombinant protein in the algal chloroplast
were verified by Western Blot analysis. Partial purification of recombinant arAmyBH was
conducted by using Ni-NTA purification system. Results demonstrated that our newly
developed chloroplast expression system could be used to produce recombinant proteins
from C. reinhardtii chloroplast. Successful expression of arAmyBH in C. reinhardtii
chloroplast established a foundation of developing economic and efficient processes of
converting microalgal starch into free reducing sugars for biofuel production.
3.2 Introduction
Some green microalgae, such as Chlamydomonas reinhardtii, are known to convert
carbon dioxide to carbohydrate, mainly in the form of starch (Choi et al., 2010; Markou et
al., 2012). Microalgal starch can be the main substrate for the production of several
significant biofuels, such as methane, bioethanol and biobutanol through anaerobic
digestion and fermentation processes (Markou et al., 2012; Yao et al., 2013). However,
mono-sugars such as glucose rather than starch would be preferred for most of biofuel
	
  

	
  43	
  

	
  
	
  
producing microorganisms (Harun & Danquah, 2011). Therefore, microalgal starch has to
be converted into mono-sugars before microbial fermentation of biofuel production.
Hydrolysis of microalgal starch is usually conducted by either acid or enzymatic
treatment. Compared to acid treatment, enzymatic hydrolysis exhibits several advantages,
such as higher sugar yield, less by-products and mild reaction conditions, though, enzymes
used for hydrolysis are relatively expensive for large-scale processing. In recent years,
expression of heterologous amylases in the starch-producing plants has been viewed as an
effective and economical way to reduce the cost of biofuel production. Several studies
have reported the successful expression of heterologous amylases in corn and rice kernels
to facilitate starch degradation (Lanahan et al., 2011; Pen et al., 1992; Xu et al., 2008).
However, there is limited information about heterologous amylase expression in
microalgal chloroplasts to hydrolyze their starch for biofuel production.
C. reinhardtii, a unicellular green microalga, has been identified as a model
organism for the production of recombinant proteins. Transformation methodologies for
C. reinhardtii chloroplast and nuclear genomes have been well established, and
pharmaceutical proteins have been successfully expressed in the chloroplast and nuclear
(Franklin et al., 2002; Mayfield & Schultz, 2004; Rosales-Mendoza et al., 2012; Sun et al.,
2003; Yang et al., 2006). Compared to the expression of heterologous genes into the
nucleus, expression in the chloroplast offers advantages such as the absence of gene
silencing, the ability to process polycistronic transcripts, and higher accumulation and less
glycosylation of transgenic proteins (Bock, 2007; Franklin & Mayfield, 2005; Specht et
al., 2010). Besides, expression of heterologous gene into chloroplast can be achieved via
integration of gene fragment into the chloroplast genome by homologous recombination
(Rosales-Mendoza et al., 2012). Thus, we developed a chloroplast expression system that

	
  

	
  44	
  

	
  
	
  
can integrate heterologous genes into the region between psaB and trnG genes without
interruption of any functional gene. Based on this expression system, amylase (arAmyBH)
from hyperthermophilic bacterium Thermotoga neapolitana was expressed in the
chloroplast to construct a C. reinhardtii capable of hydrolyzing its own starch after
cultivation.
3.3 Materials and methods
3.3.1 Strains and culture conditions
Wild-type cells of C. reinhardtii CC-125 mt+ (137c) (obtained from the
Chlamydomonas Resource Center) were cultured on solid Tris-Acetate-Phosphate (TAP)
medium (Harris, 1989) with 1.5% (w·w-1) agar. Recombinant cells were cultured on the
solid TAP medium with 1.5% agar and 100 ug·mL-1 spectinomycin (Sigma-aldrich, St.
Louis, MO). The wild type and recombinant cells from solid cultures were then transferred
to the liquid TAP medium (without spectinomycin). Both solid and liquid cultures were
incubated at room temperature (around 25 °C) under continuous light (1000 lux). Liquid
cultures were conducted on a rotary shaker (Thermal Scientific, Odessa, TX) at 200 rpm.
Cell number was determined by hemocytometer. Cells were harvested by centrifugation at
8,000 g for 10 min at 4 °C. Cell mass separated from the culture was then washed with
deionized water twice, dried overnight and weighted to calculate dry biomass
concentration.
Escherichia coli JM109 [endA1 glnV44 thi-1 relA1 gyrA96 recA1 mcrB+ Δ(lacproAB) e14- [F' traD36 proAB+ lacIq lacZΔM15] hsdR17(rK-mK+)] (Promega, Madison,
WI) was used as the host for expression plasmid construction. E. coli transformants were
grown at 37 °C in the selective Luria-Bertani broth: 1% (w·v-1) tryptone (BD, Franklin

	
  

	
  45	
  

	
  
	
  
Lakes, NJ), 0.5% (w·v-1) yeast extract (DOT scientific Inc., Burton, MI) and 0.5% NaCl
with 100 µg·ml-1 ampicillin and 50 µg·ml-1 spectinomycin.
3.3.2 Plasmid construction
It has been reported that transgenes’ codons need to be optimized based on codon bias
of C. reinhardtii chloroplast genes in order to achieve high levels of heterologous protein
expression (Manuell et al., 2007). Therefore, we changed the codon usage of the gene of
amylase AmyB from T. neapolitana based on its amino acid sequence (GenBank
EU871663) (Park et al., 2010) to reflect the codon bias of C. reinhardtii chloroplast genes
(Table 8).
Table 8 Codon usages of wild type AmyB gene and synthesized arAmyBH gene
	
  

	
  

Amino acid

AmyB (T. neapolitana)

arAmyBH

I

AUU, AUC, AUA

AUU

L

UUA, UUG, CUU, CUC, CUA, CUG

UUA

V

GUU, GUC, GUA, GUG

GUU

F

UUU, UUC

UUU

M

AUG

AUG

C

UGU

UGU

A

GCU, GCC, GCA, GCG

GCU, GCA

G

GGU, GGC, GGA, GGG

GGU

P

CCU, CCC, CCA, CCG

CCU, CCA

T

ACU, ACA, ACG

ACU, ACA

S

AGU, UCU, UCC, UCA, UCG

UCA

Y

UAU, UAC

UAU

	
  46	
  

	
  
	
  
Table 8 (cont’d)

W

UGG

UGG

Q

CAA, CAG

CAA

N

AAU, AAC

AAU

H

CAU, CAC

CAU, CAC

E

GAA, GAG

GAA

D

GAU, GAC

GAU

K

AAA, AAG

AAA

R

CGU, CGA

CGU

Stop

UGA

UAA

A His6-Tag coding sequence was added to the C-terminus of the codon-optimized AmyB
gene and the resulting arAmyBH gene (GenBank KF703932) was synthesized by Epoch
Life Science, Inc (Sugar Landing, TX). Plasmid P-423 (obtained from the
Chlamydomonas Resource Center) containing a 1.9 kb aadA chloroplast expression
cassette (Goldschmidt-Clermont, 1991) was used to create the arAmyBH chloroplast
expression cassette, in which the 0.81 kb aadA gene was replaced with the arAmyBH gene
by using two restriction endonucleases NcoI and PstI and a 437 bp fragment of rbcL gene
3’-UTR was replaced with a 414 bp fragment of atpA gene 3’-UTR by using PstI and
SacI, giving plasmid P-423-arAmyBH. The 3’-UTR of the atpA gene was generated by
PCR with a forward primer (plus pstI site) (5’CTGCAGTTTTAATTAAGTAGGAACTCGGTATATGC-3’) and reverse primers (plus
SacI site) (5’-GAGCTCCTGTATAAACAAAAATTTTTAATGTTAAC-3’) (Barnes et
al., 2005).

	
  

	
  47	
  

	
  
	
  
Plasmid p699 (Fig. 7a) contains the aadA chloroplast expression cassette (5’psbA —
aadA—3’psbA; 1.5 kb) which confers resistance to spectinomycin, flanked by 1.8 kb
fragment of the rbcL and trnG genes and a 0.8 kb fragment with part of the psaB gene
(GuhaMajumdar et al., 2008). pBlueScript II KS(+) replaced the pUC18 part of the
plasmid p699 (Fig. 7b) by cutting both plasmids with restriction enzymes at the indicated
sites, treating with Klenow polymerase to generate blunt ends, and then ligating them
together. A 10-base insertion containing restriction enzyme digestion sites EcoRI and
BamHI was then introduced into the plasmid using the QuikChange™ site-directed
mutagenesis kit (STRATAGENE, Agilent Technologies, La Jolla, CA) with the forward
primer (5’-CAATCAAAGAATGCGAATTCGGATCCCAATTCGTCCTATTTTAATAC3’) and reverse complementary primer (5’-GTATTAAAATAGGACGAATTGGGAT
CCGAATTCGCATTCTTTGATTG-3’). In order to amplify the mutated plasmid DNA,
after the initial denaturation at 95 °C for 5 min, the cycling parameters for 18 cycles were
as follows: 95 °C for 1 minute, 55 °C for 1.5 minutes, and 68 °C for 8 minutes. At the end
of temperature cycling, 10 U (1 µL) of Dpn I restriction enzyme was added, and the
reaction mixture was incubated at 37 °C for 1 hour to digest the parental supercoiled
dsDNA. The resultant plasmid was denoted as pBp699 (Fig. 7c). Plasmid pBp699 was cut
with EcoRI and BamHI, and the arAmyBH chloroplast expression cassette was excised
from P-423-arAmyB plasmid with EcoRI and BamHI to be ligated into the digested
pBp699. The resulting plasmid was named p666 (Fig. 7e), which contains both the
arAmyBH cassette and aadA cassette flanked by segments of the psaB gene and rbcL-trnG
genes for homologous recombination into the C. reinhardtii chloroplast genome between
the psaB and trnG genes without any interruption of chloroplast genes after biolistic
transformation. The integration site is shown in Fig. 7f. All constructs were confirmed by

	
  

	
  48	
  

	
  
	
  
restriction and PCR analyses, and the open reading frames (ORF) of the aadA and
arAmyBH genes were verified by sequencing.

Figure 7 Flow chart of plasmid construction. (a) Plasmid p699. (b) Plasmid
pBluescript II KS (+). (c) Plasmid pBp699. (d) arAmyBH chloroplast expression
cassette. (e) Plasmid p666 with primer sites indicated as P1, P3, and P9. (f)
Homologous integration site in the cpDNA segment.

	
  

	
  49	
  

	
  
	
  
3.3.3 Chloroplast transformation
Wild-type cells of C. reinhardtii strain CC-125 mt+ (137c) were transformed with the
plasmid p666 as described by Boynton et al (Boynton & Gillham, 1993) and Kindle et al
(Kindle et al., 1991b) using a PDS Helium-1000 particle gun (Bio-Rad, Hercules, CA).
The transformant cells were incubated in liquid TAP medium overnight under dim light.
For spectinomycin selection, the cells were then carefully plated on TAP agar containing
100 µg·mL-1 spectinomycin and cultured in moderate light at 25 °C. After 2-3 weeks, the
colonies were picked and restreaked on TAP agar plate with spectinomycin and grown for
about one week. Since the chloroplast contains multiple copies of their genomes, at least
three rounds of selection were conducted to be confident of having a strain in which all
copies of the genome were transformed.
3.3.4 DNA isolation and PCR amplification
Total DNA was extracted from cells in liquid culture as described by GuhaMajumdar
et al (Guhamajumdar & Sears, 2005). A pair of PCR primers was designed based on the
chloroplast genome sequence 5’ downstream of the arAmyBH cassette (P1: 5’CACATAACTCCACGTAAGCGCATTTTCTTACAATC-3’) and the 3’ sequence of
atpA 3’UTR in the arAmyBH cassette (P3: 5’-GACGAATTGGGATCCCCGGGTAC-3’).
To amplify the 2.3 kb expression cassette of the arAmyBH gene (primer sites shown in
Fig. 7e), conditions for the 30 cycles of PCR were: denaturation at 95 °C for 4 min,
followed by 94 °C for 1 min, 1 min at 54 °C to anneal the primers with the template, and 3
min at 72 °C for DNA extension. A final extension was done for 10 min at 72 °C. All of
the PCRs were performed with a mastercycler (Eppendorf, Hauppauge, NY). The PCRamplified products were separated on 1% (w·v-1) agarose gels in Tris-borate-EDTA
buffer.

	
  

	
  50	
  

	
  
	
  
To check the presence of both arAmyBH and aadA expression cassettes, the primer P1
and the chloroplast genome sequence 3’ upstream of the aadA cassette (P9: 5’GTTCGAGTCCCATCATCCGCTAAACC-3’) were used to generate a 3.8 kb PCR
product. After the initial denaturation at 95 °C for 4 min, the PCR cycles were repeated 30
times: 93 °C for 15 s, 30 s at 54 °C to anneal the primers and template, and 4.5 min at 68
°C for DNA extension. The final extension was done for 10 min at 68 °C. The PCR
products were run on 1% (w·v-1) agarose gels.
PCR products from the amplification were submitted for sequencing at the Michigan
State University Genomics Technology and Sequence Facility. Megalign software from
the DNAStar program (Madison, WI) was used for sequence analysis.
3.3.5 Protein extraction and Western blot analysis
For Western blot analysis, cells of C. reinhardtii and E. coli were broken by sonication
(Ultrasonic Liquid Processors, Qsonica, LLC., Newtown, CT) in a buffer solution
containing 100 mM Bis-Tris-Propane (pH 7.0) and 1% (v·v-1) protease inhibitor cocktail
(Sigma, St Louis, MO). Soluble proteins were separated by centrifugation at 13,000 g and
4 °C for 5 min, and the resulting supernatant was used for Western blot analysis. Western
blots were carried out as described by Cohen et al (Yohn et al., 1998), using Monoclonal
Anti-polyHistidine antibody (Sigma, St Louis, MO). Concentration of crude protein was
tested following BCA kit analysis method (Ruan et al., 2012).
3.3.6 Enzyme purification
For partial purification of the recombinant enzyme arAmyBH, the recombinant C.
reinahrdtii strain carrying arAmyBH was grown in liquid TAP medium at 25 °C. Cells
were harvested by centrifugation at 10,000 g and 4 °C for 20 min, re-suspended in lysis
buffer (50 mM NaH2PO4, 300 mM NaCl, pH 8.0), and broken as described in section 2.5.
	
  

	
  51	
  

	
  
	
  
The crude cell extract was centrifuged at 13,000 g and 4 °C for 5 min to remove cell
debris. The clear lysate was then mixed with pre-equilibrated Ni-NTA slurry (QIAGEN)
by gently shaking at 200 rpm for 60 min at 4 °C. The lysate-Ni-NTA mixture was loaded
into a column and rinsed with washing buffer (50 mM NaH2PO4, 300 mM NaCl, 5mM
imidazole, pH 8.0). The His6-arAmyBH was eluted with buffer containing 200 mM
imidazole. The active fractions in the elution buffer were dialyzed 3 times against 50 mM
sodium phosphate buffer (pH 6.5) and concentrated using an Amicon Ultra-15 30K
filtration device. The protein concentration was determined according to the Bradford
method, using bovine serum albumin as a standard (Bradford, 1976). The purity and
molecular weight of the proteins were analyzed by gel electrophoresis in a 12% (w·v-1)
sodium dodecyl sulfate-polyacrylamide (SDS-PAGE) (Laemmli, 1970) and precision plus
protein standards (Bio-rad, Hercules, CA).
3.3.7 Enzyme assay
The amylase activity was analyzed in 50 mM sodium phosphate buffer (pH 6.5) based
on Park’s method (Park et al., 2010). 1 mL reaction system contained 0.25 mL 1% (w·v-1)
soluble starch and 0.75 mL enzyme solution. The reaction mixture was incubated at 75 °C
for 1 hour and the enzyme reaction was terminated by quenching on ice. Reducing sugar
generated in the system was analyzed by 3, 5-Dinitrosalicylic acid (DNS) method (Miller,
1959). One unit of arAmyBH activity was defined as the amount of enzyme required to
release 1 nmol of reducing sugar per minute under described test conditions. 3.4	
  Results	
  
3.4.1 Expression of arAmyBH in the C. reinhardtii chloroplast
Transformation of the C. reinhardtii chloroplast with plasmid p666 was achieved by
biolistic bombardment. After spectinomycin selection, putative transformed clones were
inoculated into liquid TAP medium. PCR amplification of the transformants was
	
  

	
  52	
  

b

	
  
	
  
performed using primers P1 and P3 to amplify the arAmyBH chloroplast expression
cassette. The expected 2.3 kb bands representing the arAmyBH cassette were amplified
from transformants and from plasmid p666 as a positive control, while no band was
amplified in wild-type C. reinhardtii (Fig. 8a).
1

2

3

4

5

6

1

7

3.0 kb
2.0 kb
0.5 kb

2

3

4

3.0 kb
2.0 kb
1.5 kb
1.0 kb
0.5 kb

a

Figure 8 PCR assays of expression cassettes (a) PCR assays of arAmyBH cassette (2.3
kb) using primers P1 and P3. Lane 1: negative control (PCR products from wild-type
strain); lane 2-5: the PCR products from C. reinhardtii transformants; lane 6:
positive control (PCR products from plasmid p666); lane 7: 1 kb DNA marker
(NEB). (b) PCR assays of the adjacent arAmyBH and aadA cassettes (3.8 kb) using
primers P1 and P9. Lane 1: negative control (wild-type); lane 2: PCR products from
C. reinhardtii transformant 7-C-1; lane 3: positive control (PCR products from
plasmid p666); lane 4: 1 kb DNA marker (NEB). (c) Primers P1 and P9 mapping to
the endogenous gene sequence. Dashed line indicates the insertion site for arAmyBH
and aadA cassettes.
To further determine if the integration of the two expression cassettes had occurred at the
intended site in the chloroplast genome, PCR amplification was performed using primers
P1 and P9 to generate a 3.8 kb band including the 2.3 kb arAmyBH cassette and the 1.5 kb
	
  53	
  
	
  

	
  
	
  
aadA cassette. As shown in Fig. 8b, the expected bands were obtained from the transgenic
clone 7-C-1 and plasmid p666, while only a small (~150 bp) product was amplified from
the wild-type strain. Hence, both cassettes were integrated into the correct site of the
chloroplast genome of C. reinhardtii. Clone 7-C-1 was selected for the following
experiments.
3.4.2 Western blot analysis
Since the protein expression apparatus in the chloroplast is prokaryotic in nature
(Fargo et al., 1998), the promoters and their 5’UTRs function in prokaryotic E. coli to
drive gene expression. Hence, E. coli carrying the plasmid p666 was spectinomycinresistant and was able to produce His6-tagged arAmyBH protein. Total proteins from E.
coli JM 109 carrying plasmid p666, C. reinhardtii transformant 7-C-1, wild type E. coli
and C. reinhardtii were extracted and analyzed by Western blot using Monoclonal AntipolyHistidine antibody.

Fig. 9 exhibited that both the transformed C. reinhardtii and E. coli, but not their wildtype strains, accumulated arAmyBH protein.

Figure 9 Western blot of wild-type and transformed C. reinhardtii and E. coli JM109.
Equal amounts of protein (25 µg) were loaded in each lane. Lane1: C. reinhardtii
transformant 7-C-1; lane 2: transgenic E. coli JM109 with p666; lane 3: wild-type C.
reinhardtii; lane 4: wild-type E. coli JM109
	
  

	
  54	
  

	
  
	
  
3.4.3 Purification and characterization of His6-tagged arAmyBH protein
His6-tagged arAmyBH protein was purified from the C. reinhardtii transformant 7-C1. Crude enzyme extracts were solubilized in lysis buffer and loaded on a Ni-NTA
column. Elutes were concentrated and dialyzed by centrifugation in Amicon Ultra-15 30K
filter. Using this procedure, we were able to obtain 120 µg of partially purified fusion
protein from 312 mg of crude protein extracts, about 0.9 g of dry biomass (~35% w·w-1
protein content). After the final purification step, 3.5% of the initial amylase activity was
recovered; a specific activity of 119.2 U·mg-1 was calculated for the recombinant
arAmyBH, with a total purification of 90-fold (Table 9).
Table 9 Purification of arAmyBH from transformed C. reinhardtii. arAmyBH
activity was determined at 75 °C in 100 mM sodium phosphate buffer (pH 6.5) with
3,5-dinitrosalicylic acid method.

	
  

Purification

Total

Total

Specific activity

Yield

Purification

step

activity (U)

protein (mg)

(U·mg-1)

(%)

(fold)

Cell extract

409.3

312

1.3

100

1

Ni-NTA

14.3

0.1

119.2

3.5

91

	
  55	
  

	
  
	
  
The partially purified recombinant protein exhibited a major band with an estimated
molecular mass of 51 kDa by SDS-PAGE analysis (Fig. 10), which was a little higher than
the molecular mass reported by Park et al (Park et al., 2010), due to the presence of the Cterminal His6-tag.

Figure 10 Partial purification of recombinant arAmyBH protein from transformed
C. reinhardtii. Lane 1: crude enzyme extracts; lane 2: partially purified arAmyBH;
lane 3: protein marker.
3.5 Conclusions
The codon optimized intracellular thermophilic alpha-amylase arAmyBH was
successfully expressed in C. reinhardtii chloroplast by the newly developed expression
system. PCR amplification and Western blot analysis confirmed the heterologous gene
integration in chloroplast genome and protein accumulation in transformed strain. The
recombinant arAmyBH was partially purified to further confirm the production of
recombinant arAmyBH in chloroplast. The hydrolysis performance of the transgenic
microalgal C. reinhardtii 7-C-1 was investigated in the next chapter.
	
  

	
  

	
  56	
  

	
  
	
  
CHAPTER 4 IN VIVO ENZYMATIC HYDROLYSIS OF TRANSGENIC
CHLAMYDOMONAS REINHARDTII STARCH

Xiaoqing Wang, Zhenhua Ruan, Danielle Boileau, Barbara B. Sears, Yan Liu, Wei Liao
4.1 Abstract
Transgenic microalgal C. reinhardtii 7-C-1 with the capability of producing a
thermophilic amylase arAmyBH in chloroplast was used to develop a novel bioprocess of
microalgal starch utilization. Properties of recombinant enzyme arAmyBH were
investigated for the enzyme application. The optimum pH and temperature of the partially
purified enzyme were 5.5 and 60 °C, respectively. After incubation at 60 °C for 3 h,
enzyme retained 60% of its maximal activity. Enzymatic hydrolysis of the algal starch
demonstrated that without breaking the cell wall, transgenic cells could completely
convert the starch into reducing sugars in vivo at an elevated temperature in the presence
of commercial alpha-amylase. Otherwise, no major differences of cell growth and starch
accumulation between transgenic strain and wild type strain were observed. The studied
process provides a new approach that can reduce enzyme usage, simplify pretreatment
step of microalgal biomass and improve the efficiency of carbohydrate-based algal biofuel
production.
4.2 Introduction
Microalgae rich in carbohydrate have great potential to be the non-food-basedfeedstock for bioethanol production. During algal bioethanol production, microalgal
biomass has to be pre-treated with dilute acids or hydrolytic enzymes in order to release
monosugars. Although acid hydrolysis of microalgal biomass has been widely used (Ho et
al., 2013a; Ho et al., 2013b; Ho et al., 2013c; Nguyen et al., 2009), an enzymatic process
	
  

	
  57	
  

	
  
	
  
would be more promising because of its higher sugar yield, mild reaction conditions,
fewer by-products and lower utility cost (Choi et al., 2010). Technology development in
the enzyme industry has provided more choices of enzyme applications towards different
microalgal carbohydrate compositions. For instance, endo-β-1,4-D-glucanase, exo- β-1,4D-glucanase and β-glucosidase are used to hydrolyze microalgal cellulose (Chen et al.,
2013); α-amylase and amyloglucosidase can be used to hydrolyze microalgal starch (Choi
et al., 2010). However, there are two main obstacles that limit the application of enzymatic
hydrolysis in the pretreatment of microalgal biomass. One is that relatively expensive
enzymes are still too costly for large-scale processing. The other is that starchaccumulating microalgal cells need to be disrupted in order to enable amylolytic enzymes
to access the cytoplasm and hydrolyze intracellular starch. Considering the rigid cell walls
of many microalgae, a significant amount of energy will be consumed during cell
disintegration (Choi et al., 2010; Xu et al., 2008). Therefore, it would be desirable to
develop a cost-effective way that can reduce the time, energy, and capital expenses of the
process of microalgal biomass pretreatment.
Utilization of transgenic crop grains that enable the production of thermophilic
amylases has been viewed as effective and economical approach to reduce the commercial
enzymes input for cost-saving purposes. Transgenic tobacco that can produce bacterial αamylase was applied to the liquefaction of potato starch and corn starch, where DE values
of 16 and 13 were achieved respectively (Pen et al., 1992). Transgenic rice seeds that are
able to produce bacterial α-amylase were mixed with corn powder to carry out corn starch
liquefaction, and a DE value of 17 was achieved (Xu et al., 2008). As for photosynthetic
microalgae, the intracellular starch granule is located in the chloroplast (Harris, 1989).
Expression of thermophilic amylase in microalgal chloroplast could be an effective way to

	
  

	
  58	
  

	
  
	
  
not only reduce the cost of pretreatment process but also enable starch degradation in vivo
and release mono-sugars out of the cells without mechanical disruption. In our previous
work, we developed a transgenic microalga C. reinhardtii 7-C-1 that is able to produce
thermophilic alpha-amylase arAmyBH in its chloroplast. As a follow-up, in this study we
investigated the characteristics of recombinant enzyme arAmyBH and developed a
process to facilitate microalgal carbohydrate hydrolysis by using transgenic microalga 7C-1.
4.3 Materials and methods
4.3.1 Strains and culture conditions
Wild-type cells of C. reinhardtii CC-125 mt+ (137c) (obtained from the
Chlamydomonas Resource Center) were cultured on the solid Tris-Acetate-Phosphate
(TAP) medium (Harris, 1989) with 1.5% (w·w-1) agar. Recombinant cells of C. reinhardtii
7-C-1 constructed as previously mentioned in chapter 3 were cultured on the solid TAP
medium with 1.5% agar and 100 ug·mL-1 spectinomycin (Sigma-aldrich, St. Louis, MO).
Solid cultures of both wild type and recombinant cells were incubated at room temperature
(around 25 ℃) under continuous light (10 klux).
For autotrophic culture, the wild type and recombinant cells were cultivated at 25 °C
in Sueoka's high salt medium (HS) medium (Sueoka, 1960). The experimental conditions
were: initial cell number around 0.3 × 106 cells mL-1, photon flux density of 10 klux and
continuous aeration with the injection of air with 6% (v·v-1) carbon dioxide. Samples were
taken during the cultivation for calculation of specific growth rate and biomass
productivity. For starch accumulation, the wild type and recombinant cells were pelleted
by centrifugation at 8,000 g for 10 min; cell pellets were then transferred into the HS
medium depleted of N (HS-N) with the initial cell number around 1.5 × 107 cells mL-1 and
	
  

	
  59	
  

	
  
	
  
cultured for 72 hours under the same condition as described above. Cells were harvested
for starch quantification and hydrolysis.
4.3.2 Cell growth measurement
Cell mass was harvested from the culture by centrifugation at 8,000 g for 10 min,
pellets were then washed twice with deionized water, dried overnight and weighted to
calculate dry biomass concentration.
The specific growth rate (µ, d-1) was determined from the dry biomass concentration
(g L-1) in the time interval of exponential phase, as shown in Eq. (1):
µμ = (𝐿𝑛(𝑋! ) − 𝐿𝑛(𝑋! ))/(𝑡! − 𝑡! )

(1)

Where X2 and X1 are the dry biomass concentration at times t2 and t1, respectively.
The biomass productivity (g L-1 d-1) was calculated by the Eq. (2):
𝑃! = (∆𝑋)/∆𝑡

(2)

Where Px is the biomass productivity, ∆X is the change of dry biomass concentration in
the cultivation time ∆t.
Statistical analysis using R software (R version 2.15.0, Vienna, Austria) was applied to
the experimental data in order to perform pair-wise comparisons using a type I error rate
of 0.05, to compute the differences between wild-type and transgenic strain regarding
specific growth rates, the volumetric growth rate and starch contents under the same
culture condition.
4.3.3 Starch analysis
Modified quantitative saccharification method from National Renewable Energy
(NREL) (Moxley & Zhang, 2007) was used to determine starch content in microalgal
biomass. A small amount of biomass was suspended in 72 % (w w-1) H2SO4 solution and
incubated at room temperature for 1 hour. Deionized water was then added to the mixture
	
  

	
  60	
  

	
  
	
  
to dilute the H2SO4 concentration from 72 % (w w-1) to 4 % (w w-1). The mixture was then
incubated at 121 °C for 1 hour. After incubation, the hydrolysate was neutralized with
CaCO3 and analyzed by HPLC (Agilent 1100) equipped with a Bio-rad Aminex HPX-87H
organic acid column and a refractive index detector. 5 mM sulfuric acid with a flow rate of
0.6 ml·min-1 was used as the mobile phase and the column temperature was set at 65 ºC.
High purity glucose from sigma was used as the standard (Ruan et al., 2012).
4.3.4 Enzyme assay
The amylase activity was analyzed in 50 mM sodium phosphate buffer (pH 6.5) based
on Park’s method (Park et al., 2010). 1 mL reaction system contained 0.25 mL 1% (w·v-1)
soluble starch and 0.75 mL enzyme solution. The enzymatic reaction was carried out at 75
°C for 1 hour and then terminated by quenching on ice. Reducing sugar generated in the
system was analyzed by 3, 5-Dinitrosalicylic acid (DNS) method (Miller, 1959). One unit
of arAmyBH activity was defined as the amount of enzyme required to release 1 nmol of
reducing sugar per minute under described conditions.
4.3.5 Effects of pH, temperature and metal ions on arAmyBH activity
Partially purified enzyme arAmyBH was used for the property analysis. The effect of
pH on the activity of arAmyBH was evaluated at 75 °C in 50 mM citrate-phosphate
(pH4.0 to 6.0), potassium phosphate (pH 6.0 to 8.0) and Tris-HCl (pH 8.0 to 9.0). In order
to find the optimal temperature of arAmyBH, activities of the partially purified enzyme
were determined at temperatures varying from 30 °C to 90 °C in 50 mM sodium
phosphate buffer (pH 5.5). Regarding the thermostability of arAmyBH, enzyme solutions
were incubated at 50 mM sodium phosphate buffer (pH 5.5) for 3 h at 40 °C, 50 °C, 60 °C
and 70 °C. Aliquots were taken hourly and placed immediately on ice to halt enzymatic
reaction. The residual activities of the aliquots were measured at the optimal temperature.
	
  

	
  61	
  

	
  
	
  
4.3.6 Hydrolysis of microalgal starch
Recombinant cells were cultured in HS-N medium to accumulate starch and were used
to test the capability of starch hydrolysis in vivo. Wild type cells were treated in the same
way for comparison purpose. Starch hydrolysis was conducted by maintaining cell pellets
at 60 °C for 2 hours, followed by cell disruption using an ultrasonic processor
(QSONICA, LLC, USA). The ultrasonic processor was used to release all intracellular
substances for sugar profile analysis (monosaccharide, disaccharide and oligosaccharide).
The hydrolysate was then centrifuged at 8,000 g for 5 minutes and the supernatant was
used for sugar profile analysis.
In addition, two treatments with and without a commercial alpha-amylase were
conducted in order to evaluate the performance of recombinant enzyme arAmyBH on
algal starch hydrolysis. The alpha-amylase from Bacillus licheniformis (0.28 U ml-1) was
added into wild-type and transgenic microalgae biomass (dry biomass concentration of 17
g L-1), respectively. The hydrolysis was conducted under the same conditions as
aforementioned except the ultrasonic processor of cell disruption. After hydrolysis, the
hydrolysate was also centrifuged at 8,000 g for 5 minutes and the supernatant was used for
reducing sugar analysis.
4.3.7 Hydrolysate profile analysis
The monosaccharide and disaccharide were analyzed by HPLC (Agilent 1100)
equipped with a Bio-rad Aminex HPX-87H organic acid column and a refractive index
detector (Ruan et al., 2012). 5 mM sulfuric acid with a flow rate of 0.6 mL·min-1 was used
as the mobile phase, and the column temperature was set at 65 ºC. High purity glucose and
maltose from sigma were used as the standards. Analysis of oligosaccharides was using a
QTRAP 3200 mass spectrometer (AB/Sciex) equipped with binary LC-20AD pumps

	
  

	
  62	
  

	
  
	
  
(Shimadzu), a SIL-HTc autosampler, and column oven. All mass spectrometric analyses,
including data processing, were performed using Analyst v. 1.4.2 software (AB/Sciex).
Oligosaccharides were separated using an Acquity UPLC BEH Amid column with a twosolvent (0.1 aqueousformic acid and acetonitrile) binary gradient. Total solvent flow was
maintained at 150 µL·min-1 and column temperature was set at 55 °C. Ions were generated
using electrospray ionization in Negative mode. Electrospray capillary voltage and
desolation temperature were 4.5 KV and 500 °C, respectively. Ions were analyzed using
Enhanced MS Scan (EMS) from 50 to 1500 Da. Sugar polymers were characterized based
on their deprotonated ([M-H]-) or phosphate adduct ions.
4.4 Results
4.4.1 Enzyme properties of arAmyBH
The amylase activity of arAmyBH was determined at various pH values. The results in
Fig.11a indicated that maximum activity was observed at pH 5.5.

a	
  

Relative activity (%)

120
100
80
60
40
20
0
3

4

5

6
pH

7

8

9

10

Figure 11 Effects of pH and temperature on the activity of arAmyBH. (a) Effect of
pH on arAmyBH amylase activity. (b) Effect of temperature on amylase activity. (c)
Thermostability of amylase activity of arAmyBH. Data points are the means of two
replicate determinations. The amylase activity of a pre-incubated sample at 4 °C was
regarded as 100%.
	
  

	
  63	
  

	
  
	
  
Figure 11 (cont’d)

c	
  

Relative activity (%)

100
80
60
40°C
50°C
60°C
70°C

40
20
0
0

0.5

1
1.5
2
2.5
Incubation time (h)

3

3.5

The activities were measured as a function of temperature from 30 to 90 °C and indicated
that the optimum temperature was 60 °C (Fig.11b). As shown in Fig.1c, the residual
amylase activities were 80 and 90% of the highest activity after treatment at 50 and 40 °C
for 3 h, respectively. Although the amylase activity was lower at 60 °C, the residual
activity was still about 60% after 3-hour incubation at 60 °C, but only 40% activity
remained at 70 °C, indicating that the enzyme was relatively stable up to 60 °C.

	
  

	
  64	
  

	
  
	
  
4.4.2 Starch hydrolysis in microalgal cells
Investigation of arAmyBH properties provided a general idea regarding recombinant
enzyme application in microalgal starch hydrolysis. Considering the optimum temperature
of arAmyBH is 60 °C and gelatinization temperature of C. reinhardtii granule starch was
at 55.3 °C (± 0.2 °C) (Buleon et al., 1997), 60 °C was thus selected to study the hydrolysis
capacity of arAmyBH in vivo. Wild-type cells were used as a control. Results in Fig. 12a
showed that 19.6 mg glucose (DP1), 84.8 mg maltose (DP2), 84.3 mg maltotriose (DP3),
and 10.3 mg maltotetraose (DP4) were produced from hydrolysis of 1 g intracellular
starch in transgenic cells.

Amount of sugar produced (mg·g-1
starch)

100.0
90.0
80.0
70.0
60.0
50.0
40.0
30.0
20.0
10.0
0.0
Glucose

Maltose

Maltotriose

Transgenic strain 7-C-1

Maltotetraose

Wild-type strain

(a)
Figure 12 (a) Amount of sugar obtained from starch hydrolysis of transgenic strain
7-C-1. (b) Enzymatic hydrolysis of microalgae starch with alpha-amylase. Data are
the average of two replicates with standard error.

	
  

	
  65	
  

	
  
	
  
Figure 12 (cont’d)

Hydrolyzed starch (%)

100
80
60
40
20
0
Heated cells

Heated cells + α-amylase

Wild-type strain

Transgenic strain 7-C-1

(b)
A trace amount of maltopentaose (DP5) was found (data was not shown), and no
oligosaccharides with higher degree of polymerization (DP≥6) were detected. Based on
the quantification of hydrolysis profile, about 20% of the starch was digested by
arAmyBH to oligosaccharides (DP ≤ 5). In comparison, the hydrolysis of wild-type
microalgal biomass generated only 2.98 mg and 0.87 mg maltotriose and maltotetraose
from 1 g starch, and there were no other degradation products found in the free cell
extracts (Fig. 12a).
The hydrolytic profile of recombinant arAmyBH was found to be the same with wildtype enzyme AmyB, where maltose was the major product during the starch degradation
followed by maltotriose and glucose (Park et al., 2010). AmyB from hyerthermophilic
bacterium T. neapolitana has been viewed as a new type intracellular amylase that
contains four highly conserved regions of glycoside hydrolase family (GH) 13 α-amylase.
Unlike typical α-amylase, AmyB hydrolyzes starch from non-reducing ends and mainly
liberates maltose and small amount of glucose, which is quite similar to the hydrolytic

	
  

	
  66	
  

	
  
	
  
pattern of maltogenic amylase (EC 3.2.1.133) (one of GH 13 α-amylases) (Park et al.,
2010).
The experimental results also demonstrated that endogenous amylase in microalgal
cells did not significantly contribute to the starch hydrolysis (Fig. 12a). It is well known
that starch hydrolysis of reducing sugar production requires both endo-acting and exoacting amylases. Since the recombinant enzyme arAmyBH worked as an exo-acting
amylase in the current study, a commercial alpha-amylase was added into the hydrolysis
system to investigate whether the recombinant amylase arAmyBH could complete the
starch hydrolysis when cooperating with the external endo-acting amylase. Furthermore,
in order to explore the feasibility of developing a novel bioprocess that could carry out
starch degradation in vivo and release reducing sugars out of the cells without mechanical
disruption, intact cells instead of cell extracts from the recombinant strain were used. As
depicted in Fig. 12b, hydrolysis of transgenic cells without supplement of commercial
alpha-amylase as a control showed that 15 % of starch was converted into reducing sugars
and released into the supernatant after a 2-hour incubation, whereas only a small amount
of reducing sugar (0.4 % of starch equivalent) was detected from the wild-type cells. For
the hydrolysis with the supplemental alpha-amylase, the reducing sugar yield reached
100% from the transgenic biomass compared to 70 % from the wild-type biomass. The
results indicated that recombinant arAmyBH alone was not sufficient to completely
degrade algal starch in the transgenic cells. However, the recombinant arAmyBH showed
a sufficient exo-amylase (such as β-glucosidase) activity to carry on the conversion of all
starch into reducing sugars when supplied with another endo-amylase, such as alphaamylase. In addition, unlike other microalgal starch hydrolysis, releasing reducing sugars
from cells in our process does not require cell disruption. It was assumed that high

	
  

	
  67	
  

	
  
	
  
temperature during starch hydrolysis denatured membrane protein, increased the fluidity
of membrane lipids, and even ruptured the membrane structure, as a result, the
permeability of cell membrane was increased, intracellular starch and oligosacchrides
could be accessed by external starch-hydrolysis enzymes, and reducing sugars could be
released (Lee, 1974). Undoubtedly, recombinant amylase arAmyBH located in the
chloroplast can directly work on starch granule in vivo and make this process more
efficient.
4.4.3 Cell growth and starch accumulation of wild-type and recombinant strains in
autotrophic culture
The two-stage cultivation strategy depicted in chapter 2 was employed to
investigate the influence of the genetic modification on the cell growth and starch

Biomass concentration g L-1

accumulation under phototrophic conditions (Fig.13)
a

0.80
0.60
0.40
0.20
0.00
0

20
Wild-type strain

40
60
80
Time (h)
Transgenic strain 7-C-1

Figure 13 Comparison of growth condition and starch accumulation between wildtype and transgenic strains in autotrophic culture. (a) Growth curves of wild-type
and transgenic strains in HS medium with 5 % CO2 under 10 klux continuous light
(b) Starch accumulation in wild-type and transgenic strains cultured in HS-N
medium with 5 % CO2 under 10 klux continuous light. Data are the average of two
replicates with standard error.

	
  

	
  68	
  

	
  
	
  
Figure 13 (cont’d)

Starch content (%, w w-1)

30

b

20

10

0
0

1

2

3
Time (d)

Wild-type strain

4

5

6

Transgenic strain 7-C-1

Wild type and transgenic 7-C-1 strains were first cultured in the HS medium for
cell growth, and then biomass were transferred into HS-N medium for starch
accumulation. Transgenic strain 7-C-1 exhibited similar growth properties as the wild-type
strain under the same culture conditions (Fig. 13a). After lag phase in the first 12 hours,
both strains started exponential growth, and reached stationary phase at 60 hours. The data
also showed that the transgenic strain had slightly lower mean specific growth rate and
biomass productivity than the wild-type strain. Mean specific growth rates in exponential
phase of wild-type and transgenic strains were 0.096 h-1 and 0.073 h-1, respectively, and
corresponding mean biomass productivities were 0.25 g L-1 d-1 and 0.21 g L-1 d-1.
It was assumed that lower specific growth rate and biomass productivity of the
transgenic strain were mainly caused by different nutrient demand of protein production
by the heterologous gene that contributed to the growth difference of two strains.
However, generally heterologous protein in C. reinhardtii chloroplast only accounts for
0.5 % - 3 % of total soluble protein in microalgal biomass (Mayfield et al., 2007), which
requires a relatively small amount of nutrient for the heterologous protein expression
	
  

	
  69	
  

	
  
	
  
during cell growth. This is why there was difference in growth between two strains,
though, the difference was not large. Wild-type and transgenic cells also demonstrated
some differences in the pattern of starch accumulation. The starch content in wild-type
cells peaked after four days of starvation (Fig. 13b), whereas the starch content in
recombinant cells grew much faster in the first two days than wild-type cells, and then
slowed down until starch content reached the peak at fifth day (Fig. 4-3b). However, the
maximum starch contents of wild-type and transgenic strains were 26.4 % and 26.8 %
respectively of the dried biomass weight, with no significant difference between them
(P>0.05).
4.5 Conclusion
Properties of recombinant amylase arAmyBH from C. reinhardtii 7-C-1 chloroplast
were investigated. The optimum pH and temperature of arAmyBH were 5.5 and 60 °C,
respectively. The hydrolysis profile of intracellular starch by arAmyBH indicated that 20
% of starch was hydrolyzed into oligosaccharides (DP ≤ 5). With the addition of the
commercial alpha-amylase, C. reinhardtii transformant 7-C-1 can digest all of
intracellular starch under elevated temperatures without cell disruption. Additionally, cell
growth and starch accumulation of transgenic strain were investigated and compared with
wild type strain. The results demonstrated that exogenous expression in the algal
chloroplast had slight influence on cell growth and the pattern of starch accumulation.
There were no signficant differences in the maximum starch contents of wild-type and
transgenic strains. Our novel process provides a new approach that can reduce the enzyme
cost, simplify the pretreatment step of microalgal biomass and improve the efficiency of
carbohydrate-based algal biofuel production.

	
  

	
  70	
  

	
  
	
  
CHAPTER 5 CONCLUSION AND FUTURE WORK
5.1 Conclusions
There are several barriers that limit the utilization of microalgal-based fermentable
carbohydrates for bioethanol production, such as poor productivity of microalgal
carbohydrate, high cost and complicated pretreatment process of microalgal biomass. In
order to make microalgal-bioethanol commercially viable, the objective of this dissertation
was to design a cultivation strategy to enhance microalgal carbohydrate production and
develop a transgenic microalga to facilitate microalgal starch hydrolysis. Unicellular green
microalga Chlamydomonas reinhardtii was selected in our research due to following
advantages: 1) it is non-toxic, non-pathogenic and food-grade safe microorganism; 2) it
has a relatively high growth rate; 3) it is a model organism for the study of photosynthesis
function; 4) its chloroplast genome can be easily transformed; and 5) it is one of the most
extensively studied unicellular green microalgae for which a lot of references are
available.
Chapter 2 provided a process strategy that accomplished the continuous culture and
harvest of microaglal biomass in order to maximize microalgal carbohydrate production.
In the first stage, microalgal culture began with batch operation under optimal culture
conditions of 10 klux light intensity, 5 % CO2 content and 9.4 mM initial NH4+
concentration. Once the cell growth reached the late exponential phase, semi-continuous
operation was carried out, which is the simplest approach to dilute and harvest cultures
periodically so that microalgal self-shading effect can be diminished, nutrients can be
refreshed, extracellular metabolites can be removed, and cell growth can be finally
maintained in the exponential phase with the highest specific growth rate. Investigation of
different culture replacement ratio concluded that the highest biomass productivity of 0.47

	
  

	
  71	
  

	
  
	
  
g L-1 d-1 was reached, which was about two-fold high as that from batch operation.
Harvested biomass was then transferred into another photobioreactor containing nitrogen
depletion medium to conduct starvation culture in the second stage. Since CO2 is an
important nutrient factor for autotrophic organism and its content can intrigue a highaffinity CO2 concentrating mechanism, the effect of CO2 content on carbohydrate
accumulation was evaluated in the second stage. The highest carbohydrate content of 71
% (percentage of total DCW) was achieved from the 4-day culture with 0.04 % CO2,
showing more than 9 times improvement compared to that from the first stage cultivation.
The two-stage strategy of microalgal cultivation developed in this chapter significantly
enhanced the cell mass productivity and promoted carbohydrate accumulation in
microalgal cells. Carbohydrates accumulated in green microalgae cells mainly exist in the
form of starch, whereas small molecular reducing sugars such as glucose or maltose rather
than starch would be preferred for microbial fermentation. Hence, the question regarding
how to effectively and economically hydrolyze intracellular carbohydrate (mainly as
starch) arose.
The most common approaches to hydrolyze microalgal carbohydrate include dilute
acid treatment and enzymatic treatment. Chemical treatment is not environmentally
friendly and results in toxic by-products and equipment corrosion. Enzymatic treatment
exhibits several advantages over chemical approach, such as higher sugar yield, fewer byproducts, and mild reaction conditions. However, enzyme consumption increases the cost,
and pre-requisite cell disruption requires large energy input and makes the process
complicated. Therefore, the chapter 3 and chapter 4 developed a transgenic microalga
along with a novel process to facilitate microalgal starch hydrolysis. Chapter 3 constructed
a transgenic microalga C. reinhardtii 7-C-1 that is able to produce recombinant

	
  

	
  72	
  

	
  
	
  
thermophilic amylase arAmyBH in its chloroplast, which was confirmed by PCR and
Western blot analysis. In chapter 4, we used partially purified arAmyBH to investigate its
properties and evaluated its hydrolysis performance at 60 °C. The hydrolysis profile
showed that around 20 % of the intracellular starch was degraded into oligosaccharides
(DP ≤ 5), and among them maltose was the major product followed by maltotriose and
glucose, which was in accordance with the hydrolytic pattern of wild type enzyme AmyB
from hyerthermophilic bacterium T. neapolitana. Because AmyB works as an exo-acting
amylase, we mixed intact transgenic microalgal cells with one commercial endo-acting
amylase and incubated the mixture at 60 °C for 2 hours. Results showed that all of the
intracellular starch in transgenic cells was degraded into reducing sugars, compared to the
70 % hydrolysis ratio in wild type cells, which demonstrated that recombinant arAmyBH
provided a sufficient exo-amylase (such as β-glucosidase) activity to accomplish starch
hydrolysis when supplied with another endo-amylase, such as alpha-amylase.
Furthermore, cell disruption before enzymatic hydrolysis does not seem to be required and
recombinant amylase in the chloroplast can significantly enhance hydrolysis efficiency.
Additionally, a growth kinetics study exhibited that there was no big difference between
transgenic strain and wild type strain regarding cell growth and starch accumulation.
5.2 Future work
Completely converting starch into fermentable sugars requires both endo-acting and
exo-acting amylases. Endo-acting amylase such as typical α-amylase attacks α-1,4 bonds
at random points in the starch molecular to break down long-chain carbohydrates into
short-chain oligosaccharides and dextrins. Exo-acting amylase such as β-amylase
hydrolyzes oligosaccharides and dextrins from the non-reducing ends to degrade them into
fermentable sugars. In the current study, recombinant enzyme arAmyBH worked as an

	
  

	
  73	
  

	
  
	
  
exo-acting amylase, and another endo-acting alpha-amylase was required to completely
degrade starch. Hence, in order to completely eliminate the necessity of commercial
enzymes during the algal biofuel production, the following research should be carried out
next steps:
•

Constructing another expression plasmid to co-express endo-acting amylase in C.
reinhardtii 7-C-1 chloroplast;

•

Optimizing expression system to achieve higher enzyme activity;

•

Conducting enzyme-directed evolution to achieve better cooperation between two
enzymes;

•

	
  

Knocking out antibiotic gene to remove environmental concern.

	
  74	
  

	
  
	
  

APPENDIX

	
  

	
  75	
  

	
  
	
  
APPENDIX A: DATA
	
  
A.1 CHAPTER 2 DATA
Table 10 Biomass concentrations under different culture conditions for Figure 3
Light
intensity
(klux)

NH4+
concentration
(mM)

CO2
content
(%)

Hours
0
12
37
49
61
72
84
0
12
24
36
48
60
70
0
12
24
36
48
60
72
84

Air

10

4.5

5

10

	
  

	
  76	
  

Biomass
concentration 1
(g/L)
0.00
0.01
0.07
0.22
0.28
0.25
0.31
0.00
0.01
0.17
0.42
0.56
0.73
0.73
0.00
0.01
0.03
0.12
0.17
0.28
0.54
0.55

Biomass
concentration
2 (g/L)
0.00
0.03
0.09
0.26
0.30
0.27
0.38
0.00
0.03
0.19
0.38
0.54
0.72
0.74
0.00
0.02
0.07
0.10
0.22
0.31
0.48
0.52

	
  
Table 10 (cont’d)
0
12
37
49
61
72
84
0
12
24
36
48
60
70
0
12
24
36
48
60
72
84
0
12
37
49
61
72
0
12
24
36
48
60
70
0
12
24
36
48
60
72
84

Air
10

9.0

5

10

Air

10

18.0

5

10

	
  

	
  77	
  

0.00
0.02
0.03
0.189
0.211
0.32
0.33
0.00
0.02
0.15
0.49
0.66
0.82
0.81
0.00
0.01
0.14
0.17
0.17
0.31
0.57
0.58
0.00
0.04
0.11
0.23
0.28
0.29
0.00
0.04
0.15
0.45
0.72
0.90
0.79
0.00
0.02
0.12
0.14
0.17
0.31
0.61
0.60

0.00
0.02
0.17
0.267
0.278
0.25
0.48
0.00
0.02
0.18
0.42
0.68
0.76
0.77
0.00
0.01
0.03
0.14
0.22
0.28
0.56
0.59
0.00
0.05
0.16
0.28
0.28
0.24
0.00
0.05
0.18
0.45
0.75
0.81
0.76
0.00
0.03
0.06
0.12
0.19
0.36
0.58
0.61

	
  
Table 10 (cont’d)
0
13
24
37
48
62
73
85
96
0
13
24
36
48
61
70
85
0
12
24
36
48
60
73
79
0
13
24
37
48
62
73
85
0
13
24
36
48
61
70
85
0
12
24
36
48
60
73
79

Air

5

4.5
5

10

Air

5

9.0

5

10

	
  

	
  78	
  

0.00
0.05
0.08
0.16
0.17
0.21
0.37
0.48
0.39
0.00
0.05
0.11
0.21
0.29
0.34
0.38
0.51
0.00
0.00
0.04
0.06
0.15
0.17
0.23
0.16
0.00
0.01
0.13
0.17
0.10
0.21
0.47
0.34
0.00
0.08
0.07
0.12
0.19
0.33
0.32
0.53
0.00
0.00
0.07
0.03
0.10
0.19
0.21
0.16

0.00
0.08
0.08
0.15
0.16
0.23
0.4
0.38
0.41
0.00
0.05
0.14
0.14
0.19
0.35
0.32
0.45
0.00
0.00
0.06
0.07
0.14
0.22
0.25
0.13
0.00
0.01
0.09
0.16
0.19
0.25
0.49
0.36
0.00
0.05
0.10
0.15
0.26
0.35
0.36
0.53
0.00
0.00
0.06
0.09
0.11
0.17
0.28
0.17

	
  
Table 10 (cont’d)
0
13
24
37
48
62
73
85
0
13
24
36
48
61
70
85
0
12
24
36
48
60
73
79

Air

5

18.0

5

10

	
  

	
  79	
  

0.00
0.07
0.13
0.14
0.17
0.22
0.36
0.35
0.00
0.05
0.07
0.15
0.26
0.44
0.30
0.48
0.00
0.00
0.05
0.06
0.10
0.16
0.16
0.12

0.00
0.04
0.05
0.15
0.12
0.26
0.39
0.33
0.00
0.04
0.06
0.20
0.23
0.31
0.35
0.48
0.00
0.00
0.03
0.06
0.10
0.19
0.23
0.18

	
  

Table 11 Biomass concentrations before and after medium replacement for Figure 4
45% replacement ratio
Days
0
1
2
Before* 0.60 0.78 0.67
Biomass concentration (g/L)
After** 0.33 0.43 0.37
Before 0.65 0.77 0.92
Biomass concentration (g/L)
After 0.36 0.42 0.50
70% replacement ratio
Days
0
1
2
Before 0.61 0.66 0.58
Biomass concentration (g/L)
After 0.19 0.20 0.18
Before 0.64 0.73 0.60
Biomass concentration (g/L)
After 0.19 0.22 0.18
83% replacement ratio
Days
0
1
2
Before 0.68 0.57 0.48
Biomass concentration (g/L)
After 0.11 0.09 0.08
Before 0.82 0.59 0.51
Biomass concentration (g/L)
After 0.14 0.10 0.08
91% replacement ratio
Days
0
1
2
Before 0.68 0.36 0.33
Biomass concentration (g/L)
After 0.06 0.03 0.03
Before 0.60 0.37 0.34
Biomass concentration (g/L)
After 0.05 0.03 0.03
*Before: Biomass concentration before medium replacement
**After: Biomass concentration after medium replacement

	
  

	
  80	
  

3
0.74
0.40
0.88
0.48

4
0.77
0.42
0.94
0.52

5
6
0.80 0.63
0.44
0.91 0.87
0.50
-

3
4
5
6
0.64
0.64 0.77
0.19
0.19
0.69 0.75 0.68 0.59
0.21 0.23 0.20
3
0.51
0.08
0.52
0.09

4
0.53
0.09
0.49
0.08

5
6
0.51 0.62
0.08
0.51 0.60
0.08
-

3
0.35
0.03
0.28
0.03

4
0.30
0.03
0.26
0.04

5
6
0.30 0.33
0.03
0.24 0.25
0.02
-

	
  

Table 12 Cell density and biomass concentration during the Stage II cultivation for
Figure 5
0.04% CO2
0
1
13.44 12.75
12.12 10.56
10.57
10.77
0.56 0.67
0.56 0.59
5% CO2
Days
0
1
Cell density (E+06 /ml)
12.69 17.67
Cell density (E+06 /ml)
19.55
Cell density (E+06 /ml)
12.31
Cell density (E+06 /ml)
19.32
Biomass concentration (g/L) 0.60 0.67
Biomass concentration (g/L) 0.60 0.68
Days
Cell density (E+06 /ml)
Cell density (E+06 /ml)
Cell density (E+06 /ml)
Cell density (E+06 /ml)
Biomass concentration (g/L)
Biomass concentration (g/L)

	
  

	
  81	
  

2
12.50
12.02
0.70
0.82

3
10.46
11.24
11.99
10.76
0.95
0.96

4
14.56
11.50
10.09
12.18
0.94
1.06

5
11.47
12.50
15.91
13.79
0.98
1.06

2
20.49
17.34
0.78
0.85

3
4
5
16.04 14.76 16.77
20.27 18.73 15.29
24.50
18.04 15.65 16.14
0.88 1.02 0.93
0.91 0.87 0.93

	
  
Table 13 Biomass compositions with different CO2 contents in the Stage II
cultivation for Figure 6
Days
Protein content (%)
Protein content (%)
Carbohydrate content (%)
Carbohydrate content (%)
Days
Protein content (%)
Protein content (%)
Carbohydrate content (%)
Carbohydrate content (%)

	
  

0.04% CO2
0
1
58.51
39.63
58.95
41.03
7.05
25.89
8.27
26.62
5% CO2
0
1
58.43
50.37
58.59
49.64
6.44
20.96
6.60
18.31

	
  82	
  

2
33.71
33.48
39.87
38.15

3
20.14
20.30
57.15
58.46

4
20.34
20.04
63.57
64.21

5
16.26
15.90
65.35
63.57

2
35.93
36.16
31.84
30.82

3
35.37
35.89
36.39
39.25

4
35.64
35.00
44.79
41.62

5
27.94
27.50
43.15
38.29

	
  

A.2 CHAPTER 4 DATA
Table 14 Enzyme activities at different pH for Figure 11a
pH
4.0
4.5
5.0
5.5
6.0
6.5
7.0
7.5
8.0
8.5
9.0

	
  

Sample 1 (U)
14.02
14.49
17.43
22.83
20.42
20.57
13.60
11.87
12.34
11.03
8.35

	
  83	
  

Sample 2 (U)
10.50
16.22
22.52
26.56
25.14
22.41
17.01
16.27
11.50
5.99
6.20

	
  

Table 15 Enzyme activities at different temperature for Figure 11b
Temperature (°C)
30
40
50
60
65
70
80
90

	
  

Sample 1 (U)
12.74
21.20
31.75
32.48
33.52
19.05
18.42
11.81

	
  84	
  

Sample 2 (U)
16.79
22.52
27.13
33.01
22.16
20.26
15.17
15.01

	
  

Table 16 Enzyme activities after incubation at different temperature for Figure 11c
Hours

40 °C

Sample
1 (U)

Sample
2 (U)

50 °C

Sample
1 (U)

Sample
2 (U)

60 °C

Sample
1 (U)

Sample
2 (U)

70 °C

Sample
1 (U)

Sample
2 (U)

0
34.58
33.69
25.98
22.88
25.98
22.88
34.58
33.69
1
33.27
33.90
23.88
23.36
19.32
18.21
22.46
24.82
2
29.76
32.75
19.16
23.36
16.27
13.02
19.37
14.75
3
29.55
31.80
21.36
16.90
11.13
16.69
17.01
10.97
Note: 50 and 60 °C are tested at the same batch, 40 °C and 70 °C are tested at another
batch

	
  

	
  85	
  

	
  

Table 17 Amount of sugars from starch hydrolysis for Figure 12a

Glucose (DP1)
Maltose (DP2)
Maltotriose
(DP3)
Maltotetraose
(DP4)

	
  

7-C-1 transgenic strain
Sugar amount
Sugar amount
(mg/g starch)
(mg/g starch)
23.87
15.24
98.32
71.20

Wild-type strain
Sugar amount
Sugar amount
(mg/g starch)
(mg/g starch)
N/A
N/A
N/A
N/A

81.82

86.73

0.82

5.15

13.59

7.00

0.58

1.16

	
  86	
  

	
  
Table 18 Hydrolyzed starch with or without alpha-amylase from two strains for
Figure 12b
Wild-type strain
Hydrolyzed
Hydrolyzed
starch (%)
starch (%)
W/O alphaamylase
W alphaamylase

	
  

7-C-1 transgenic strain
Hydrolyzed
Hydrolyzed
starch (%)
starch (%)

N/A

N/A

17.94

10.91

70.55

71.32

99.81

101.29

	
  87	
  

	
  

Table 19 Comparison of biomass concentration between wild-type and transgenic
strains for Figure 13
Wild-type strain
Hours
0
12
24
Biomass concentration (g/L) 0.00 0.01 0.17
Biomass concentration (g/L) 0.00 0.03 0.19
Transgenic strain
Hours
0
12
24
Biomass concentration (g/L) 0.00 0.00 0.10
Biomass concentration (g/L) 0.00 0.00 0.02

	
  

	
  88	
  

36
48
60
70
0.42 0.56 0.73 0.73
0.38 0.54 0.72 0.74
36
48
60
70
0.20 0.29 0.67 0.58
0.28 0.41 0.61 0.67

	
  

Table 20 Comparison of starch content between wild type and transgenic strains for
Figure 13

Days
0
Starch content (%) 3.93
Starch content (%) 4.03
Days
0
Starch content (%) 5.02
Starch content (%) 5.63

	
  

Wild-type strain
1
2
3
12.79 19.42 22.19
11.17 18.80 23.94
Transgenic strain
1
2
3
18.40 23.00 24.77
17.44 17.25 21.10

	
  89	
  

4
5
6
27.32 26.32 25.10
25.39 23.36 22.87
4
5
6
25.20 27.75 26.64
24.57 25.82 25.61

	
  

BIBLIOGRAPHY

	
  

	
  90	
  

	
  
BIBLIOGRAPHY
	
  
	
  
	
  
1. Bai, F., Anderson, W., Moo-Young, M. 2008. Ethanol fermentation technologies from
sugar and starch feedstocks. Biotechnol. Adv. 26, 89-105.
2. Ball, S.G., Dirick, L., Decq, A., Martiat, J.C., Matagne, R.F. 1990. PHYSIOLOGY OF
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