bioprospecting and discovery of novel microalgal isolates...
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Bioprospecting and discovery of novel microalgal isolates with
multiple product potential.
by
Amritpreet Kaur Minhas
Submitted in fulfilment of the requirements for the degree of Doctor of Philosophy
Deakin University
May, 2016
DEAKIN UNIVERSITY
CANDIDATE DECLARATION
I certify the following about the thesis entitled “Bioprospecting and discovery of novel microalgal isolates
with multiple product potential”
Submitted for the degree of Doctor of Philosophy
a. I am the creator of all or part of the whole work(s) (including content and
layout) and that where reference is made to the work of others, due
acknowledgment is given.
b. The work(s) are not in any way a violation or infringement of any
copyright, trademark, patent, or other rights whatsoever of any person.
c. That if the work(s) have been commissioned, sponsored or supported by
any organisation, I have fulfilled all of the obligations required by such
contract or agreement.
d. That any material in the thesis which has been accepted for a degree or
diploma by any university or institution is identified in the text.
i
Aknowledgements I would like to thank my supervisors, Dr. Alok Adholeya and Prof. Peter
Hodgson, for providing me the excellent opportunity, guidance and
throughout support for the challenging research topic i worked with, I am
especially grateful for the passion Dr. Alok have shared in my research
endeavors besides mentorship which helped me sail through and timely
completion of my lab work as well as writing. Not to undermine his
willingness to offer suggestions and constructive criticism throughout
this journey of research and publications so far. My special gratitude
to my associate supervisor, Prof. Colin Barrow, thank you for all your
technical advice and critical support throughout thesis and manuscript
writing for my publications. Your continued encouragement is much
appreciated. I truly appreciate the effort of all the members in the lab,
especially Mr Chandrakant Tripathi, Ms Deeprajni, Ms Priyanka Gupta,
Ms Shikha Chaudhary, Mr. Yaspal, for assisting me with fluorescence
microscopy, SEM, GC and HPLC analysis. All other colleagues, seniors
and lab staff are thankfully acknowledged for their continuous support
and encouragement.
I also would like to thank immensely Dr Adarsh Gupta and Dr
Munish Puri for extending all support and sharing methodologies during
my short but productive stay at Deakin University. I am thankful to
administration of TERI as well as divisional secretaries for their
patience in assisting me throughout and at difficult times. Last but not
the least, I would like to express my most sincere and personal gratitude
to my family members, Dadaji, Dadi ji and Rajat for consistant
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encouragements and love. Special thanks to my elder brother and father
who always stood by me during critical times. My younger Brother and
elder sister always played pep up role. Mother‘s are always special and I
thank my mother for remaining understanding and supportive.
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Table of contents
Acknowledgements………………………………………………………....i
List of Tables………………………………………………………………...ix
List of Figures ……………………………………………………………… xi
Publications/Conference presentations………………………………………xv
List of abbreviations………………………………………………………...xvii
Abstract ............................................................................................................ xx
Chapter 1: Introduction and Literature review ............................................ 1
1.1 Introduction ................................................................................................. 1
1.1.1 High value products from microalgae ................................................. 5
1.2 Microalgal lipids ........................................................................................ 11
1.2.1 Lipid biosynthesis ................................................................................ 12
1.2.2 Cultivation medium ........................................................................... 16
1.2.3 Strategies to enhance stress based lipids changes in microalgae ......... 17
1.3 Microalgae: Collection and isolation ......................................................... 17
1.3.1 Collection of diverse microalgae ......................................................... 18
1.3.2 Isolation and purification techniques .................................................. 19
1.3.3 Microalgal identification strategies ..................................................... 20
1.3.3.1 Morphometric identification ....................................................... 21
1.3.3.2 Molecular identification ............................................................. 21
1.4 Exploring the richness of biodiversity ....................................................... 22
1.5 Autotrophic stress factors .......................................................................... 23
1.5.1 Temperature ....................................................................................... 23
1.5.2 Light ................................................................................................... 25
1.5.3 Nitrogen sources ................................................................................. 26
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1.5.3.1 Cow urine ................................................................................. 28
1.5.4 Phosphate ........................................................................................... 28
1.5.5 Salinity ............................................................................................... 29
1.6 Carotenoids produced by microalgae ......................................................... 30
1.6.1 Bioactive compound .......................................................................... 30
1.6.2 Strategies to enhance microalgal carotenoids production .................. 33
1.6.2.1 Autotrophic growth factors: Temperature ................................. 34
1.6.2.2 Light ......................................................................................... 35
1.6.2.3 Nitrogen sources ....................................................................... 37
1.6.2.4 Salinity ...................................................................................... 38
1.6.2.5 Iron ............................................................................................ 39
1.7 Multiple factors that influence the synthesis of lipids and carotenoids ...... 46
1.8 Solvent compatibility studies for algal milking of live cells during
continuous fermentation. ............................................................................. 49
1.9 Conclusion ................................................................................................. 52
1.10 Aims of the study ..................................................................................... 54
Chapter 2: Isolation, identification and preservation of new strains of
microalgae from different geographical region of India and the World
2.1 Introduction
2.2 Materials and Methods ............................................................................... 57
2.2.1 Microalgal sample collection .............................................................. 57
2.2.2 Isolation and purification of microalga .............................................. 59
2.2.2.1 Direct plating method................................................................. 59
2.2.3 Long term preservation of microalgae ................................................ 60
2.2.4 Characterization of isolates by scanning electron microscopy (SEM) . 61
v
2.2.5 Cultivation of microalgal strains .......................................................... 61
2.2.6 DNA extraction from diverse isolates .................................................. 62
2.2.7 PCR amplification, and 18S rDNA sequencing ................................... 62
2.2.8 Construction of degenerate algal-specific 18S rDNA primers ............. 63
2.2.9 Phylogenetic analysis ........................................................................... 64
2.3 Results and Discussion ............................................................................... 64
2.3.1 Isolation of microalgae ....................................................................... 64
2.3.2 Chemical treatment: Purification of microalga ................................... 67
2.3.3 Streak-plating and Serial dilution ........................................................ 68
2.3.4 Preservation of microalgae .................................................................. 69
2.3.5 Identification of diverse microalgal isolates using SEM and molecular
biological techniques .................................................................................... 71
2.4 Conclusion ................................................................................................. 83
Chapter 3: Characterising new oil and carotenoid producing
microalgal isolates with biofuel potential
3.1 Introduction ................................................................................................ 84
3.2 Materials and Methods ............................................................................... 86
3.2.1 Chemicals used ................................................................................... 86
3.2.2 Culturing of microalgal isolates .......................................................... 87
3.2.3Screening of microalgal isolates........................................................... 87
3.2.4 Qualitative analysis of lipids using BODIPY 505/515 staining .......... 89
3.2.5 Spectral Monograms and Biomass composition using Attenuated total
reflectance (ATR-FTIR) .............................................................................. 90
3.2.6 Lipid extraction and preparation of FAMEs ...................................... 90
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3.2.7 Gas chromatography .......................................................................... 91
3.2.8 Evaluation of microalgal oil properties for biofuel ............................ 92
3.2.9 Determination of microalgal carotenoids ........................................... 93
3.3 Results and Discussion ............................................................................... 94
3.3.1 Lipid quantification using BODIPY 505/515 ............................................................. 94
3.3.2 ATR-FTIR analysis ............................................................................. 97
3.3.3 Comparative analysis of photo-autotrophic growth, biomass and lipid
levels .......................................................................................................... 108
3.3.4 Fatty acid production of microalgal isolates....................................... 115
3.3.5 Estimation of biofuel properties ......................................................... 117
3.3.6 Standard curve preparation for carotenoid analysis ........................... 121
3.3.7 Lutein and astaxanthin productivity .................................................... 122
3.4 Conclusion ................................................................................................ 124
Chapter 4: Influence of rate limiting factors on lipids, carotenoids and
biofuel properties in four selected isolate
4.1 Introduction
4.2 Materials and Methods ............................................................................. 130
4.2.1 Chemicals used ................................................................................. 130
4.2.2Strain selection and inoculum preparation ......................................... 130
4.2.3 Measurement of algal growth ........................................................... 137
4.2.4 Biomass productivity ........................................................................ 138
4.2.5 Lipid extraction and preparation of FAME ........................................ 138
4.2.6 Gas chromatography ......................................................................... 139
4.2.7 Evaluation of microalgal oil properties for biofuel ........................... 139
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4.2.8Determination of microalgal carotenoids ............................................ 139
4.3 Results and discussion.............................................................................. 140
4.3.1 Growth curves of Scenedesmus bijugus ............................................. 140
4.3.2 Different stress induction during pre-stationary and stationary phase145
4.3.3.1 The biomass, lipid production of Scenedesmus bijugus in response
to different stress conditions at pre-stationary phase ................................ 145
4.3.2.2 The biomass, lipid production of Scenedesmus bijugus in
response to different stress conditions at the stationary phase ................. 148
4.3.2.3 Total fatty acid composition and biofuel properties of
Scenedesmus bijugus at pre-stationary and stationary phase ................... 149
4.3.4 Effect of light and photoperiod on the biomass and lipid production
of Coelastrella sp. ...................................................................................... 164
4.3.5 Analyses of carotenoids from Coelastrella sp .................................... 173
4.3.6 Cultivation of Chlorella sp. Using two stage cultivation strategy 175
4.3.7 Effect of salinity, nitrogen and cow urine in biomass and lipid
production………………………………………………………………….179
4.3.7.1FAME analysis ………………………………………………….181
4.3.7.2 Carotenoids analysis ..................................................................... 186
4.4 Conclusion .................................................................................................. 195
Chapter 5: Non-destructive extraction of lipids and recovery of active cell
biomass ...................................................................................................... 198
5.1 Introduction ............................................................................................. 198
5.2 Materials and Methods ............................................................................. 201
5.2.1Organism and culture conditions ....................................................... 201
5.2.2 Experimental design ......................................................................... 202
5.2.3 Algal growth determination ............................................................. 202
5.2.4 Cell count ......................................................................................... 203
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5.2.5 Scanning electron microscopy (SEM) .............................................. 203
5.2.6 Algal lipids analyses ....................................................................... 203
5.3 Results and Discussion ............................................................................. 207
5.3.1 Effect of organic solvents on algal growth and cell viability ............. 208
5.3.2 Effect of n-dodecane on lipid profiles ............................................... 210
5.4 Conclusion ............................................................................................... 215
Bibliography ................................................................................................... 222
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List of tables
Chapter 1:
Table 1.1 Existing market for algae-derived ingredients…………............ 8
Table 1.2 Effect of stress factors on lipid and bioactive production……41
Chapter 2:
Table 2.1 Collection sites of microalgal isolates from diverse habitats…..66
Table 2.2 Molecular determination of microalgal isolates………………..77
Chapter 3:
Table 3.1 Biomass productivities, lipid content, lipid productivities,
lutein content and astaxanthin and lutein productivity of microalgal
isolates…………......................................................................................112
Table 3.2 Comparison of lipid productivity of microalgal isolates with
other published data …………………………………………………..114
Table 3.3 Percentage of SFA, MUFA, PUFA and estimated biodiesel
properties from the FAME profile of twenty two microalgal isolates…..119
Chapter 4:
Table 4.1 Percentage of SFA, MUFA, PUFA and estimated cetane number
from the FAME profile of Scenedesmus bijugus……………………….156
Table 4.2 SFA, MUFA, PUFA and desired biofuel characteristics of
Coelastrella sp. in two stage cultivation process………………………172
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Table 4.3 SFA, MUFA, PUFA and biofuel properties of Chlorella sp.
under tested condition………………………………………………….185
Table 4.4 SFA, MUFA, PUFA and biofuel characteristics of
Auxenochlorella protothecoides………………………………………..195
Chapter 5:
Table 5.1 Treatments performed to evaluate long term milking of
algae……..................................................................................................206
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List of figures
Chapter 1:
Figure 1.1 General scheme of microalgae biorefinery concept……………4
Figure 1.2 Role of various carbon sources and nitrates in lipid biosynthesis
………………………………………………………………………….. .14
Figure 1.3 Schematic diagram showing impact of environmental stresses
on the lipid and carotenoids production…………………………………17
Figure 1.4 Carotenogenesis in Haematococcus…………………………..32
Chapter 2
Figure 2.1 Map of World and India depicting sampling locations………..58
Figure 2.2 Mixed partial purified cultures……………………………......67
Figure 2.3 Pure cultures on nutrient agar and algae agar plates………….68
Figure 2.4 Formation of algal colonies on agar plates……………………69
Figure 2.5 Maintenance of unialgal isolates: Streak plate………………..69
Figure 2.6 Preservation of diverse algae…………………………………70
Figure 2.7Scanning electron micrographs of microalgal isolates showing
some of cell shapes observe……………………………………………..72
Figure 2.8 Phylogenetic analysis of algal isolates by the Maximum
Likelihood method based on the Tamura-Nei model………………….82
Chapter 3:
Figure 3.1 Representative confocal laser scanning micrographs of diverse
microalgal isolates using BODIPY 505/515 under tested growth
conditions………………………………………………………………...97
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Figure 3.2 FTIR microspectroscopic spectra of different microalgal species
from diverse habitats……………………………………………………107
Figure 3.3 Biomass (mg/L/d) and lipid productivity (mg/L/d) of twenty
two microalgal isolates in 9 days old culture………………………… ..110
Figure 3.4 Growth pattern of different microalgal species from diverse
habitats cultivated in multicultivator MC 1000-OD for 10 days at 25 °C
under 16 h of light (120 μE/m2/s) alternating with 8h of darkness…….111
Figure 3.5 Changes in morphology and colour of Scenedesmus sp. at
stages of cultivation……………………………………………………..115
Figure 3.6 Fatty acid composition (wt %) of 22 microalgal isolates under
test conditions. In each bar, fatty acids are shown in descending order of
the number of carbon atoms in each…………………………………...117
Figure 3.7 Calibration curves for each carotenoids standard…………...122
Figure 3.8 Top ten isolates showing lipid and biomass productivity
against lutein productivity (mg/L/d)…………………………………...123
Chapter 4:
Figure 4.1 Combined effect of phosphate and salinity with cow urine on
the growth of Scenedesmus bijugus…………………………………….141
Figure 4.2 Biomass, lipid productivity (mg/L/d) and lipid content (% dry
wt.) of Scenedesmus bijugus under different treatments……………… 147
Figure 4.3 FAME profile of 7 day old culture under different tested
conditions……………………………………………………………….151
Figure 4.4 FAME profile of 13 day old culture under different tested
conditions.................................................................................................152
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Figure 4.5 Lutein and astaxanthin productivity (mg/L/d) against biomass
productivity (mg/L/d) in Scenedesmus bijugus under different
treatments……………………………………………………………….160
Figure 4.6 Growth pattern of Coelastrella sp. under tested treatments in
two stage cultivation process……………………………………………161
Figure 4.7 Biomass, lipid productivity and lipid content of Coelastrella sp.
under different tested treatments in two stage cultivation process……...164
Figure 4.8 Fatty acid composition of Coelastrella sp. in two stage
cultivation process under different tested conditions…………………...168
Figure 4.9 Lutein and astaxanthin productivity of Coelastrella sp. in two
stage cultivation…………………………………………………………173
Figure 4.10 Growth kinetics of Chlorella sp. in two stage cultivation
process…………………………………………………………………...178
Figure 4.11 Lipid content, biomass and lipid productivity of Chlorella
sp. in two stage cultivation process…………………………………….180
Figure 4.12 FAME profile of Chlorella sp. in two stage cultivation
process under tested condition………………………………………..183
Figure 4.13 Lutein and astaxanthin productivity of Chlorella sp. in two
stage cultivation process……………………………………………….189
Figure 4.14 Growth curves for Auxenochlorella protothecoides under
photoautotrophic conditions …………………………………………….191
Figure 4.16 FAME profile of Auxenochlorella protothecoides at 6 and 12
day under tested conditions……………………………………………..194
Figure 4.17 Lutein and astaxanthin productivity of Auxenochlorella
protothecoides under different tested conditions……………………….196
xiv
Chapter 5:
Figure 5.1 Flow diagram showing steps involved throughout the
experiment ……………………………………………………………...207
Figure 5.2 Figure 5.2 Culture grown under three different treatments
in aqueous phase (T1), aqueous –organic phase (T2) and aqueous
(salinity stress)-organic phase at 12 and 18 day ………………………209
Figure 5.3 Effect of n-dodecane on Scenedesmus bijugus growth in an
aqueous– organic and non-organic system……………………………...210
Figure 5.4 Biomass productivity of Scenedesmus bijugus at 12 and 18
day with and without solvent treated algae……………………………211
Figure 5.5 SEM images before and after solvent exposure…………...212
Figure 5.6 Summary of lipid classes identified in solvent phase and
algal cells in long term milking experiments………………………….214
Figure 5.7 Microalgae the bio-refinery concept………………………...215
xv
List of publications
Accepted/Published
1. Minhas, A.K., Hodgson, P., Barrow, C.J., Sashidhar, B., Adholeya, A.
(2016). The isolation and identification of new microalgal strains
producing oil and carotenoid simultaneously with biofuel potential.
Bioresource technology. 211, 556–565.
2. Minhas, A.K., Hodgson, P., Barrow, C.J., Adholeya, A. (2016). A review
on the assessment of stress conditions for simultaneous production of
microalgal lipids and carotenoids. Frontiers in microbiology. 7, 546.
Conference/symposia presentations
1. Minhas, A.K.*, Sharma, S., Beri, S., Ramakrishnan, V., Adholeya, A.
Screening, isolation and characterization of microalgae for high lipid
profile and oil content to achieve enhanced biofuel extraction,
International conference on Association of microbiologist (AMI), 3-6
november 2011. Chandigarh, India.
2. Minhas, A.K.*, Beri, S., Lamba, S., Sharma, S., Adholeya, A. Lipid
profile signatures of multivariate habitat based micro algae and its
potential for optimal aviation fuel, International conference on green
chemistry (ICGC), 7-9 december 2011.Jaipur ,India.
3. Minhas, A.K., Beri, S., Kochar, M.*, Aggarwal, N., Adholeya, A.
Lipid profiling and bioactives signatures of multivariate habitat based
microalgae
as potential sources of optimal aviation fuel, 8th Asia-Pacific Conference
on Algal Biotechnology, 9-12 July 2012, Adelaide, Australia.
4. Minhas, A.K.*, Hodgson, P., Barrow, C.J., Sashidhar, B., Adholeya, A.
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The isolation and identification of new oil and carotenoid producing
microalgal strains with biofuel potential, 9th annual Algae Biomass
Summit, 30 september - 2 october 2015 at Washington D.C, USA.
5. Attended Bio commodity refinery (BioCore) workshop at Amritsar,
India and Czech republic, Prague.
xvii
Abbreviations °C - Degree celsius Cm - Centimeter % - Percentage μm - Micrometre μg - Microgram mg - milligram mL - Millilitre μL - Microlitre mg L-1 - Milligrams per litre
mg L-1 d-1 - Milligrams per litre per day
h - Hours
min - Minutes α - Alpha γ - Gamma ω - Omega Δ - Delta β - Beta C - Carbon SEM - Scanning electron microscopy
GPS - Global positioning system
xviii
DHA - Docosahexaenoic acid
EPA - Eicosapentaenoic acid
AA - Arachidonic acid
GLA - Gamma-linolenic acid
SFAs - Saturated fatty acids
MUFAs - Monounsaturated fatty acids
PUFAs - Polyunsaturated fatty acids
LC-PUFA - Long chain polyunsaturated fatty acids
FA - Fatty acids
TFA - Total fatty acids
TAGs - Triacylglycerols
BBM -Bolds basal medium
FAS - Fatty acid synthase
DNTPs - Deoxynucleotide triphosphates
BHT - Butylated hydroxytoluene
DNA - Deoxyribonucleic acid
rDNA - Ribosomal deoxyribonucleic acid
rRNA - Ribosomal ribonucleic acid
PCR - Polymerase chain reaction
Kb - Kilo base
kHz - Kilo hertz bp - Base pairs sp. - Species FTIR - Fourier Transform Infrared
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SEM - Scanning electron microscope
GC - Gas Chromatography
MS - Mass spectrophotometry HPLC - High-performance liquid chromatography
v/v - Volume by volume
w/v - Weight by volume SD - Standard deviation Mu - Absorbance Units OD - Optical density NADPH - Nicotinamide adenine dinucleotide phosphate
Acetyl Co-A -Acetyl co-enzyme A
FAMEs - Fatty acid methyl esters SC -Secondary carotenoids
ROS -Reactive oxygen species PBRs -Photobioreactors CN - Cetane number IV - Iodine value SV - Saponification value
xx
Abstract Microalgae are capable of producing biomass rich in lipids such as long-
chain polyunsaturated fatty acids (LC-PUFA), vitamins and value added
products such as carotenoids (lutein, zeaxanthin, and astaxanthin). Co-
products from microalgae is very challenging and holds a wide
application in nutraceutical industries as food additives and human
health. Diverse gene pool of microalgae were explored for their potential
to produce multiple products from same biomass. Two hundred samples
were collected from aquatic and terrestrial rocky habitats from Asia, North
America, and the Middle East. Isolated cultures were converted into
unialgal form. The identification of the isolates was confirmed by 18S
rDNA sequencing using universal primers. Twenty two isolates were
selected and further confirmed at species level using newly-designed
primers. Scanning electron microscopy was performed to study the cell
surface morphology in twenty two isolates. Bioprospecting studies
were carried out in twenty two isolates for their potential to produce
lipids and carotenoids. Fourier transform infrared (FTIR) micro-
spectroscopic analysis mdemonstrated the presence of lipids and fats (band
1745 cm-1), including the presence of unsaturated fatty acids (band 3045
- 2796 cm-1). GC analysis showed that over 50% fatty acid from the
twenty isolates were mainly C16:0, C18:1 and rest 50 % were C18:2,
C18:3 with few traces of long chain fatty acids, indicating that these
isolates may be useful as omega-3 producers and has fatty acid profile
suitable for biofuel production. Further biofuel properties in the twenty
isolates were estimated and showed that 18 out of 22 isolates were
xxi
matching with European and US biofuel standard specification. HPLC
analysis showed that twenty two isolates demonstrated the potential to
produce lutein and astaxanthin.The four most superior isolates were
selected on the basis of producing lipids and carotenoids in the same
biomass from single growth cycle. A variety of stress factors were
implemented on the selected four isolates to further enahance the
production of lipids and carotenoids in two stage and continuous mode
system. All the four isolates found to be influenced positively under
stress factors employed, resulting in increased biomass, lipids and
carotenoids. Cow urine as nitrogen source, salinity stress, nitrogen
deficiency in the medium resulted in higher biomass, lipids, carotenoids
and suitable fatty acid profile as compared to earlier found most
optimum conditions, indicating that these isolates can grow on low
cost nutrient sources. Alteration in light intensity (low to high) and
photoperiod (12:12 to 24:0h) resulted in increased biomass, lipids and
carotenoids in two stage system in one of the Coelastrella sp. (V3). Out of
four isolates Scenedesmus bijugus was most suitable for biofuel and
carotenoids production, Coelastrella sp. has more carotenoids potential,
Chlorella sp. useful as omega-3 producer and Auxenochlorella
protothecoides showed both biofuel and carotenoids potential.
Scenedesmus bijugus which showed enhanced production of lipids and
carotenoids under salinity stress was further selected for biocompatible
studies. The same medium with same salinity stress and control condition
was used, and was saturated with biocompatible solvents (n-dodecane).
The isolates were studied for lipid production in organic phase and
algal cells. The isolates produced few traces of palmitic, oleic and
xxii
linoleic acid in the organic phase. The high amount of linoleic acid
(C18:2) was only present in medium saturated with n-dodecane at 18 day.
This work have shown that some of the naturally occurring isolates have
novel properties of producing multiple products such as lipids and
carotenoids from same biomass in one growth cycle. Some of the further
selected isolates have shown ability to respond positively to combination
of stress conditions and altered nutrient sources towards enhanced
production of multiple product. One isolate selected with the objective of
milking lipids while keeping the organism alive using biocompatible
solvent have shown potential possibility and recovered well and shown
indication towards re-synthesis of lipids when attempted to be revived.
Novel approaches employed through this work, amply indicate a
potential of not only recovery of multiple product but also in a
continuous mode which if further explored can contribute in a most
desirable next generation biorefinery concept.
Chapter 1
1
Chapter 1: Introduction and Literature review
1.1 Introduction
Microalgae are potential producers of both energy and chemicals,
particularly lipids and other commercially useful organic compounds,
and so there is considerable research interest in microalgae. Since the
reservoirs of fossil fuels are being depleted, the demand for biomass-
derived fuels and food from microalgae is increasing due to high
productivity per area (Yen et al., 2013) compared with other crops
such as soyabean (Amaral et al., 2010). Alternative technology for
biomass derived fuels is partly driven by concerns regarding increased
levels of carbon dioxide being released into the atmosphere due to
combustion of fossil fuels (Demirbas, 2010). It is well established that
microalgae have a unique ability to tolerate and consume a high level of
carbon dioxide discharged from various industrial operations in the form
of flue gas (Nigam and Singh, 2011). Since the early 1970s,
bioprospecting of microalgae (such as green algae, cyanobacteria and
diatoms) has been carried out for various uses, leveraging their ability to
consume CO during photosynthesis and providing a sustainable source of
renewable biofuels (Jones and Mayfield, 2012). The interest for large scale
cultivation of microalgal biomass that are rich reserves of high value
metabolites is increasing, however detailed literature on its optimal and
economically more sustainable production is not available (Clarens et
al., 2010; Norsker et al., 2011; Soratana and Landis, 2011). Selecting
strains that have a high cellular lipid content and high lipid productivity
is also important for reducing the costs of microalgae-based biofuel
Chapter 1
2
production (Wijffels et al., 2010). Microalgal have high growth rates
and higher productivity (about 50 times) compared with terrestrial crops
and so are considered to be next generation feedstocks (Jacob-Lopes
and Franco, 2010). Highly explored for lipids, microalgae also produce
metabolites such as carotenoids (lutein, zeaxanthin, and astaxanthin),
long-chain polyunsaturated fatty acids (LC-PUFA), that are widely used
in nutraceuticals industries as food additives (Priyadarshani and Rath,
2012). Besides cost, a major issue for the microalgae industry is
preselection of appropriate strains that produce commercially useful
levels of valuable products, as is the development of suitable cultivation
conditions for further enhancement of the above products (Ramírez-
Mérida et al., 2014). The production of lipids, carotenoids and algal
biomass can be enhanced under environmental stress factors (Mata et
al., 2010; Mulders et al., 2014; González et al., 2015). In addition to
stress factors, selection and use of microalgal species and strains is
also important for enhancing the metabolite production. Therefore,
exploration of a wider and more diverse gene pool of microalgae is
required. For economical production, deriving multiple products such as
lipids and high-value by- products from the same biomass in one growth
cycle is one way (Campenni et al., 2013; Nobre et al., 2013). Further
optimizing culture conditions, by selecting organisms that can overcome
the limitations imposed by ambient conditions, and by selecting strains
that produce high lipid content can also lower the unit cost of
microalgae-based biofuels (Wijffels et al., 2010).
Microalgae are useful in aquaculture as sources of biomolecules and
biomass that can improve the nutritional value of food or provide
Chapter 1
3
additional health benefits (Yaakob et al., 2014). Carotenoid production
from microalgae remains an active research area with additional
commercial potential (Wichuk et al., 2014). Carotenoids from microalgae
are already used extensively in food, cosmetics, and pharmaceutical
products as colouring agents and antioxidants (Prieto et al., 2011).
However, synthetic versions of some of these carotenoids are still
lower cost to produce and dominate the larger markets, such as for
astaxanthin in aquaculture feed. Many algae produce a variety of
antioxidants and pigments (carotenoids including fucoxanthin, lutein, β-
carotene, astaxanthin and phycobilliproteins) and also produce long-chain
polyunsaturated fatty Acids (PUFAs) and protein (Gouveia, 2014) with
applications in food, feed, agriculture and pharmaceutical industries
(Markou and Nerantzis, 2013), as shown in Figure 1.1.
Chapter 1
4
Figure 1.1 General scheme of microalgae biorefinery concept
The present review highlights knowledge in algal biodiversity and ways
to improve microalgal production in biotechnology. The impact of the
cultivation strategies used for microalgae growth, mainly focusing on
the environmental stress factors, for increasing the production of lipids
and value-added products (namely, lutein and astaxanthin). We provide
Chapter 1
5
information on developing methods that can maximize the production of
biofuel, biomass, and other value-added byproducts (carotenoids) from
the same species or strains of microalgae that will become a win–win
strategy in the coming decades. The collection, isolation and
identification of microalgae from diverse habitats is also described.
Finally, we discuss methods for the milking of multiple products from
one growth cycle using biocompatible solvents.
1.1.1 High value products from microalgae
Algae can produce a variety of antioxidants and pigments (carotenoids
including fucoxanthin, lutein, β-carotene, astaxanthin, and
phycobilliproteins); LC-PUFA; and proteins (the essential amino acids
methionine, threonine, and tryptophan) (Gouveia, 2014) with wide
applications in food, feed, agriculture, and pharmaceutical industries
(Markou and Nerantzis, 2013) (Figure 1.1). Microalgae are useful in
aquaculture as sources of biomolecules and biomass that can improve
the nutritional value of food or provide additional health benefits
(Yaakob et al., 2014). However, for food and aquaculture applications in
particular, low cost production is important for commercial success. The
production yield of microalgal biomass is higher under heterotrophic
conditions than under photoautotrophic conditions. Although, the relative
cost of substrate to products with respect to energy balances is more cost
intensive in heterotrophic cultivation (Chen et al., 2011) compared to
photoautotrophic.
Many reports suggested that microalgae are too expensive as a source for
biofuel, without the co-production of more valuable compounds. Table 1.1
gives recent estimates of the production cost of microalgal value-added
Chapter 1
6
products along with omega FAs. Currently, three major production
systems are used for microalgal cultivation: open raceway ponds,
horizontal tubular photobioreactors (PBRs), and flat-panel PBRs.
Raceway ponds are much better than open ponds for cultivation of
microalgae (Jacob-Lopes et al., 2014). Several semi-commercial or
commercial projects using open ponds and PBRs for producing value-
added products from microalgae are operational worldwide (Table 1.1).
The omega-3 FA market was worth $690 million in 2004, and the Asian
Omega-3 PUFA market was expected to be worth $596.6 million in 2012
(Seambiotic, Israel 2013). The global market for EPA and DHA is
estimated at $300 million and $1.5 billion, respectively (Table 1.1). A
few industries in Europe and USA have also started producing EPA and
DHA from microalgae (Table 1.1), to be used as food additives. In near
future, the market for PUFA (particularly omega-3) is expected to grow.
PUFA (particularly, γ-linolenic acid (GLA, 18:3 ω-6), (EPA, 20:5 ω-3),
arachidonic acid (ARA, 20:6 ω-6), docosapentaenoic acid (22:5 ω-3) and
(DHA, 22:6 ω-3) is gaining interest because of their application in
health industry (Fraeye et al., 2012). Table 1.1 provides an overview of
the main products from microalgae that are in the early stage of
development (particularly lutein), notably for the cosmetics industry.
Cyanobacteria strains produce various intracellular and extracellular
metabolites that show antibacterial, antifungal, antiviral, antitumor, anti-
HIV, anti-inflammatory, antioxidant, antimalarial, or herbicidal properties
(Semary, 2012).
One of the major producers of Spirulina, Hainan Simai Pharmacy Co.
(China), produces 3000 tonnes of biomass annually (Priyadarshani and
Chapter 1
7
Rath, 2012). Table 1.1 shows that a large group of industries produce β-
carotene, a food additive, from D. salina in Australia. Many species
produce high concentrations of β-carotene, astaxanthin, and
canthaxanthin, which have significant applications as antioxidants and
natural dyes (Table 1.1). The market cost of β-carotene estimated at $261
million by 2010 as shown in Table 1 .1 and expected to grow up to
$334 in 2018. On the other hand, the total market value of
cyanobacteria and phycobiliproteins is estimated at $6–$11 million
(Yaakob et al., 2014). The commercial market for lutein is $150
million a year in USA (Fernández-Sevilla et al., 2010) and was $233
million in 2010 in the poultry industry and as a nutritional supplement
(Borowitzka, 2013) which is expected to grow up to $309 by 2018. The
pigment phycocyanin is produced from Spirulina platensis by a small
number of industries and even lutein and astaxanthin are produced by
many industries worldwide (Table 1.1). Recent trends show that
industries are entering the markets for EPA, DHA, astaxanthin, and
phycocyanin. Products that are well studied include fucoxanthin,
proteins, β-glucan (a polysaccharide) and phycoerithrin (a pigment).
Chap
ter 1
8
Ta
ble
1.1
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Chapter 1
11
1.1 Microalgal lipids
The class of lipids known as polyunsaturated long chains alkanes or
alkanol (PULCA) are the membrane unbound lipids which appear
similar to neutral lipids found to be present in golgi, plasma membrane
and chloroplast thylakoid (Matthew., 2005). Khozin-Goldberg et al.
(2005) suggested that there are two major classes of lipids within the
chloroplastic lipids: Eukaryotic lipid and prokaryotic lipids, the former
contains fatty acid; palmitic or oleic acid at the sn-1 site of their
glycerol skeleton in chloroplast and later contain only C18 fatty acid at
both position (18/18) produced using extracholoroplastic lipids.
Docosahexaenoic (DHA) and arachidonic acid (ARA) are major fatty acid
present in marine algae. The microalgal lipid class under stress conditions
is an basic criterion for selection of microalgae for their further application
in biofuels or in nutraceutical applications (Olmstead et al., 2013).
Microalgal lipids are divided in to two main categories, namely
storage lipid (non-polar) and structural lipids (polar). Storage lipids are
present in form of triacylglycerides (TAGs) mainly SFAs and UFAs which
can be transesterified to biodiesel. Some microalgae also produce large
amounts of lipids in the form of TAGs (Sharma et al ., 2012) . The
synthesis of lipids in microalgae varies with the species (from either
freshwater or marine habitats or in cyanobacteria) used (Hu Q et al.,
2008). Structural lipids on other hand are essential component of cell
memebrane having animal and human application (Sharma et al., 2012).
Green algae share their ancestry with higher plants and their metabolic
mechanisms and photosynthetic pigments are similar to those found in
Chapter 1
12
higher plants (Yu et al., 2011). However, suitable metabolic pathways
need to be designed for engineering strains rich in lipids (Bellou et al.,
2014). An unusual feature of algae is that they can store large quantities
of lipids in the form of oil globules in parts of the cell other than
chloroplasts and they have the capability to link the electron transport
system to hydrogen production (Radakovits et al., 2010). PUFAs contains
huge group of fatty acids mainly ω-3 fatty acids (Lozac'h, 1986). The
molecular structure of omega-3 fatty acids comprises of an (16 to 24) even
number of carbon chains with 3 to 6 double bonds. Docosahexaenoic acid
(DHA) and eicosapentaenoic acid (EPA) are two associates of the ω-3
family having 22 and 20 C-atoms with 6 and 5 cis-double bond respectively
, (Lozac'h, 1986).
This suggests that different evolutionary groups of microalgae may
have evolved different mechanisms for lipid biogenesis. Over the past 20
years research has been carried out in order to understanding mechanism
of lipid accumulation in oleaginous species (Ratledge, 2001).
1.2.1 Lipid biosynthesis
In unicellular organisms, fatty acid biosynthesis follows a complex
pathway where synthesis starts with the formation of acetyl CoA by the
ACCase gene through acetyl CoA carboxylation. This is the key step at
which carbon is assigned for lipid synthesis (Hu Q et al., 2008) as shown
in Figure 1.2 (Modified from Perez-Garcia et al., 2011). Lipid
biosynthesis has been explored most extensively in Chlamydomonas
reinhardtii. For example, nutrient depletion in this microalga brings about
structural and behavioural changes in terms of deposition of fatty acid in
to TAGs and accumulation of starch (Moellering and Benning, 2010).
Chapter 1
13
Moreover, TAGs serves as a protective mechanism under these conditions
(Sharma et al., 2012). A simplified scheme for fatty acid and TAG
biosynthesis is shown in Figure 1.2. Li et al. (2012) showed that
neutral lipid accumulation in the cell occurs through the conversion of
various carbon source or starch to lipids, but the conversion depends on
specific microalgal strains, because different strains have different ways
to transmit the carbon flux from the carbohydrate pathway for
synthesizing lipids. Apart from lipids, FAs can be synthesized in larger
quantities by various means, such as overexpressing the genes that
regulate the enzymes responsible for lipid content, increasing
accessibility of the precursor molecule, such as acetyl CoA, and using
inhibitors that down-regulate the catabolism of FAs to change the FA
profiling through regulatory enzymes such as desaturases (Hannon et al.,
2010). The first metabolic study for increasing FA accumulation was
reported in Cyclotella cryptica in the overexpression of acetyl-CoA
carboxylase gene (ACCase) (Dunahay et al., 1996). Moreover, the
regulation of gene expression has been well studied in cyanobacteria
(Sakamato et al., 1994). For example in Phaedodactylum tricornutum,
overexpression of malic enzyme resulted in 2.5 fold increase in the total
lipids under nitrogen deficient condition as compared to a control (Xue et
al., 2014). However, Ming et al. (2010) determined that the two major
fatty acids, i.e.α-linolenic acid (18:3) and docosahexaenoic acid (22:6),
which formed the intermediates in fatty acid (PUFA) synthesis, were
known to be derived from 18:2 (n-6) through temperature regulated
enzyme omega 3 desaturation (Yongmanitchai and Ward., 1989). This
enzyme is found in marine microorganisms (Chen et al., 1991). There
Chapter 1
14
are two pathways which are evoked during the biosynthesis of fatty
acids, one being the prokaryotic pathway and other being the eukaryotic
pathway. The prokaryotic pathway is mainly chloroplastic, C16 and
C18 fatty acids are desaturated up to 18:3 omega-3 and 16:3 omega 3,
whereas in the eukaryotic pathway, which contain both cytoplasmic and
chloroplastic lipids, 18:1 is desaturated up to 18:2 (Khozin-Goldberg
and Cohen, 2006). Microalgae are known to synthesize very-long-
chain polyunsaturated FAs (VL-PUFA, >20C), and the synthesis is
regulated by desaturases and elongases, which are temperature-sensitive
enzymes (Niu et al., 2013). PUFA found in fish is concentrated from
ingested microalgae. Moreover, EPA and DHA are important valuable
fatty acid in microalgae due to their elevated levels in these
organisms (Spolaore et al., 2006; Yen et al., 2013) (Gimpel et al., 2015).
Omega-6, such as γ- linolenic acid (GLA) and arachidonic acid (ARA),
is also an interesting microalgae PUFA. GLA is produced by Spirulina
(Blanco et al., 2007; Ronda and Lele, 2008) and ARA is produced from
Porphyridium (Durmaz et al., 2007; Guil-Gucrrcro et al., 2000).
Chapter 1
15
Figure 1.2 Role of various carbon sources and nitrates in lipid osphate
NAD+oxidoreducatse; ach 1, 1aconitate; idh 1, isocitrate
dehydrogenase-NADH dependent; idh 2, isocitrate dehydrogenase-
NADPH dependent; ogd 1, ketoglutarate dehydrogenase; fum1, fumarate
hydratase; sdh 1, succinate dehydrogenase; mdh 3, malate dehydrogenase
scla 1, succinate-CoA ligase (ADP forming); FAS, fatty acid synthesis;
TAG, triacylglycerols; id, isocitrate dehydrogenase; rbcL, Ribulose
bisophosphate carboxylase/oxygenase large subunit; rbcS, Ribulose
bisophosphate carboxylase/oxygenase small subunit; prk,
Phosphoribulokinase (Perez-Garcia et al., 2011)
Several series of enzymes may be involved in controlling the level of
production of PUFA in cells. Some microalgae adapt well to various
environmental conditions that affect cellular processes including lipid
metabolism (Juneja et al., 2013). The family of very long chain-PUFA
(VLC-PUFA) holds promise in pharmaceutical and food industries; its
members can store VLC-PUFA in TAGs as storage lipids, which helps
the organism to cope with stress factors such as low temperature and
ultraviolet radiation (Bigogno et al., 2002). The distribution of each FA
is species specific. For example in Phaeodactylum tricornutum,
overexpression of malic enzyme resulted in 2.5 folds increase in the total
lipids under nitrogen deficient condition as compared to control (Xue et
al., 2014). On other hand, some microalgae adapt well to various
environmental conditions that affect cellular processes including lipid
metabolism (Juneja et al., 2013). Therefore, several physicochemical
factors such as nutrients, temperature, salinity, pH and the growth
phase and age of the culture, directly affect the composition of TAGs
Chapter 1
16
in a cell (Juneja et al., 2013). Under abiotic forms of stress, many
microalgae produce TAGs that can serve as feedstock for biofuel
production. However, the amounts of TAGs vary with the species and the
genera (Trentacoste et al., 2013). Various stress factors induce changes in
the metabolic activities of a cell such as activation of starch and
accumulation of TAGs leading to lipid bodies gathering in algae
(Johnson and Alric, 2013). Indeed, several fundamental studies on lipid
synthesis confirm that microalgae can be considered to be a feedstock for
biofuel (Yu et al., 2011). Since lipid biosynthesis process is complex, it
is challenging to increase total lipid levels in microalgae. A full
understanding of metabolic pathways and their manipulation is
necessary to improve lipid biosynthesis yields in microalgae through
metabolic engineering.
1.2.2 Cultivation medium Diversity of culture medium are available for the growth of freshwater and
backwater algae. The major freshwater media are: Bolds basal medium
(BBM) used for freshwater Chlorophyceae, Xantophyceae,
Chrysophyceae, and Cyanophyceae and BG11 used for Freshwater soil
Cyanophyceae (Barsanti and Gualtieri, 2006). These media types can
support the growth of microalgae from specific aquatic environments,
such as fresh water and brackish water. The above cultures in specified
medium are maintained at light and dark photoperiods essential for the
preservation of wide ranges of isolates with 12:12h and 16:8 h light:
dark regime (Lorenz et al., 2005).
Chapter 1
17
1.2.3 Strategies to enhance stress based lipids changes in microalgae Autotrophic and heterotrophic strategies have been studied widely, and
several physicochemical stress factors that affect the metabolism of
microalgae significantly have been identified. Managing environmental
stress is a typical approach used in refining microalgae based lipid
production in the laboratory and at the pilot-scale. The most common
stress factors used for enhancing lipid production are light intensity,
temperature, and nitrates (Figure 1.4).
Figure 1.3 Schematic diagram showing impact of environmental stresses
on the lipid and carotenoids production.
1.3 Microalgae: Collection and isolation
Microalgae are widely distributed and have a longer evolutionary history
than terrestrial plants. They show a rich diversity with more than 200
000 species (Guiry and Guiry, 2014). In recent years, unique strains have
been isolated from a range of habitats in the tropics, including both
Chapter 1
18
aquatic (lakes, streams, and backwaters) and terrestrial habitats. Such
extremes as very high temperatures and prolonged exposure to intense
light found in the tropics have conferred on extremophiles some distinctive
physiological properties. Nevertheless, extremophilic conditions such as
high temperature, longer and higher exposure of light, prevailing in
tropical regions of the world have enabled them to perform better with
distinctive physiological expression (Gouveia and Oliveira, 2009). It has
been reported that many green microalgae can tolerate high salinity as
well as survive in brackish water (Mutanda et al., 2011). Green algae
are rich in chlorophyll a and b, β-carotene, lutein and xanthophyll
(Graham, 2009). Green algae have been used extensively used for
production of secondary metabolites, such as β-carotene and astaxanthin
(Hannon et al., 2010). Some species have the ability to accumulate high
levels of carotenoids under optimum conditions. The haptophytes genera
are well known for excretion of polysaccharides and have wide
application in aquaculture. These haptophytic algae are also rich in
PUFA such as EPA and DHA (Meireles et al., 2003).
1.3.1 Collection of diverse microalgae
Conventional sampling processes for microalgae require specialised
equipment and are time consuming (Anandraj et al., 2007). The
equipment required for sampling includes knife, mesh, vials, scalps and
GPS (Mutanda et al., 2011).
Several methods for sampling algae has been reported such as by chipping
and brushing. The brushing method for collectionof microalgae from
rocks was developed by MacLulich. (1986) and is the most widely
used method. Microalgae collection is effected by several abiotic or
Chapter 1
19
biotic environmental factors (Mutanda et al., 2011). During collection of
microalgae the location latitude and longitude coordinates of the site
should be noted for future collection (Woelfel et al., 2007). Diverse
habitat at different geographical location exhibit nutrient variability and
other climatic conditions that result in additional algal diversity (Bernal
et al., 2008). Tropical and subtropical regions tend to have different
species, for example, with low temperature promoting growth of
Scenedesmus sp., whereas colder temperatures promote growth of
chlorophytes. Therefore collection of isolates from both high and low
temperature environments is important for the isolation of diverse
microalgae (Sheehan et al., 1998).
1.3.2 Isolation and purification techniques
Gravimetric separation, isolation using agar and dilution techniques are
used for the isolation of microalgae. Gravimetric separation techniques
could be more effective in separation of smaller and larger organism. A
mild centrifugation can be used to separate dinoflagellate (loose pellet)
and other smaller algae (Anderson, 2005). Isolation using agar is a
common method for isolating microalgae. The diluted microalgae
samples are spread plated on agar plates and incubated for 2 weeks. Most
of the algae does not grow on the surface of agar and instead grows
embedded in agar. This method can give pure culture without any
further plating or chemical treatments (Brahamsha, 1996). On other
hand, the dilution method provides inoculation of single cells in broth
culture, thereby establishing a single cell isolates. Further, the cultures
are made unialgal by diluting the culture in a distilled water, culture
medium or combination of these two (Anderson, 2005).
Chapter 1
20
Bacterial and fungal contamination should be avoided during the
isolation of microalgae as the isolation temperatures (20-25°C) promote
the growth of faster growing microorganisms. There are various method
to purify algae such as the use of antibiotics, phenol or addition of
germanium dioxide to the growth medium to inhibit growth of other
microbes (Abu et al., 2007). The biocide chloramine-T has been used in
agar plates and liquid media to minimize the growth of unwanted
organisms. Axenic cultures can be obtained by treating isolated algae to
an extensive washing procedure further.
1.3.3 Microalgal identification strategies
Identification of specific microalgae is foremost challenge. Biochemical
and molecular techniques have been used to identify algae. Microscope-
based identification methods are the standard protocols used for quick
screening of algal isolates. However, conventional methods sometimes
gives misleading result due to their often change in cell morohology during
their growth cycle (Godhe et al., 2002; Bertozzini et al., 2005). The
identification of microalgae using microscopy based technique is very
tedious and time consuming process and requires lot of taxonomical skills
(Godhe et al., 2002). Morphometric identification of microalgae has been
carried out using scanning electron microscopy (Mutanda et al., 2011)
Moreover, species-level taxonomical identification through 18S rDNA
sequencing is considered authentic for identification of algae up to genera
and species level (Moro et al., 2009).
Chapter 1
21
1.3.3.1 Morphometric identification The morphological examination of microalgae has conventionally been
recognised as an important tool for identification of species (Fletchner,
1998, 2008). However it is limited since small algae like Chlorella are
difficult to identify under light microscopy. Scanning electron
microscope (SEM) has been developed to improve microalgal
morphological identification (Gärtner et al., 2015). This method is also
widely used in studies of diatoms (Bacillariophyceae) (Henderson and
Reimer 2003; Houk et al. 2010).
1.3.3.2 Molecular identification
The molecular identification of microalgae is based on 18S rDNA
sequencing, which defines the taxonomic association of unknown
isolates by comparison with known isolates (Pr¨oschold and Leliaert,
2007, Fawley, 2004). Confirming an organism phylogeny is important
in terms of knowing the adaptive traits that have been collected through
natural selection and their usefulness in biotechnology. It involves PCR
based amplification of 18S rDNA gene, utilising a set of universal
primers as well as designed primer for different strains and comparative
phylogenetic analysis (Auinger et al., 2008). The multiple sequence
alignment of the isolated 18S rDNA sequences and available sequences
give a branching relationship between the new isolates, which were
previously reported and deposited with the NCBI database as complete
sequence from different algae collection centres/resource databases. The
use of multiple alignment was achieved using Bio-Edit Sequence
Alignment Editor version 7.2.5
(http://www.mbio.ncsu.edu/Bioedit/bioedit.html). The algorithms
Chapter 1
22
available at MEGA6 ver. 6: Maximum Likelihood (ML), Neighbour-
Joining (NJ) and Maximum Parsimony (MP) were employed to
construct phylogenetic relationships among the algae isolates.
1.4 Exploring the richness of biodiversity Biodiversity is characterized by organisms taxonomical and morphological
traits, as well as through the physiology and biochemistry of the
isolates (Litchman and Klausmeier, 2008). Microalgae play a major role
in maintaining the balance between abiotic stresses and living organism
(Nasser and Sureshkumar, 2012). The habiat and diversity relationship of
eukaryotic microalgae is not well understood, particularly when
compared to land plants. The origin of land plants from green algae has
further led to initiation of the terrestrial ecosystem (Kenrick and Crane,
1997). The diversity in microalgae is huge, consisting of 64 taxa and
70,000 species (Guiry, 2012), ranging from 30,000 to one million in the
most diversified Bacillariophyceae group (Armbust, 2009). However,
only 2000 species has been explored so far in the Chlorophyceae family
(Chu, 2012), which indicates a lot of potential to identify new and
useful species exists in the microalgal world. More than 20 species from
marine, freshwater (lake, ponds) have been well studied (Chu, 2012).
Several studies have investigated microalgal diversity, for example 141
taxa of freshwater microalgae from high altitude of Western ghats has
been reported by Nasser and Suresh kumar. (2012). Many green algae
are found to be present in both hot and cold environments (Lewis and
Lewis, 2005; De Wever et al., 2009; Schmidt et al., 2011) as well as
high saline (Vinogradova and Darienko, 2008) and marine habitats
(Zechman et al., 2010). Applying rate limiting factors to enhance the
Chapter 1
23
biomass and metabolites within specific microalgae is necessary to
achieve commercially useful production of compounds of interest.
1.5 Autotrophic stress factors 1.5.1 Temperature Temperature is a stress factor that greatly influences the rate of growth, net
lipid productivity and FA profiles in a wide range of microalgal
species (Ho et al., 2014b). As microalgal growth and secondary
metabolic pathway vary with temperature (James et al., 2013), greater
knowledge of the biochemical response to temperature may yield useful
insights into developing efficient systems for biofuel production (Wei
and Huang, 2015). Normally, higher growth rate of microalgae is
achieved by increasing the temperature to its optimum level (González-
Fernández et al., 2012).
Studies on a large number of species have shown that both low and high
temperatures can boost lipid productivity (Converti et al., 2009; Xin et
al., 2011). Net lipid productivity decreased as temperatures increased from
25 °C to 35 °C in Monoraphidium sp. SB2 (Wu et al., 2013); from 15 °C
to 20 °C in Nannochloropsis oculata (Converti et al., 2009); LX1, from
20 °C to 30 °C in Scenedesmus sp. (Xin et al., 2011); and, from 14 °C to
30 °C in Nannochloropsis sp. (Hu et al., 2006) (Table 1.2). Both low and
high temperatures are preferred for attaining higher lipid profiles,
depending on the species: the levels of unsaturation in FAs increase under
low temperatures, whereas those of total saturated FAs increase at high
temperatures, as observed by Liu et al. (2005). The level of
unsaturation is high at low temperatures mainly because of the higher
concentration of dissolved oxygen (DO), which allows the oxygen-
Chapter 1
24
dependent enzymes, known as omega-3 desaturases, to function (Ward
and Singh, 2005).
Besides lipids, ratio of saturated FAs to unsaturated FAs is species
dependent but only partly because of innate differences in the ability of
the species to synthesize lipids (either DHA or other FAs) under low
temperatures. In species such as Nannochloropsis sp. and P.
tricornutum, high (31.7% and 24.2% respectively) content of EPA
C20:5 (omega-3) have been reported at low temperature stress (Hu et
al., 2006; Jiang and Gao, 2004). The influence of temperature on lipid
productivity is complex and requires further investigation. However, it
appears that these strains incorporate EPA into their membranes to
counteract the negative effects of decreased membrane fluidity at low
temperatures. Furthermore, an increased level of EPA in TAG lipids
protects the photosynthetic machinery of the cell against low temperatures
(Hoffman et al., 2010). Further, temperature is also known to affect
carbohydrate production in microalgae; for example, in Spirulina sp.
carbohydrate content increased up to 50% when temperature was
increased from 25 °C to 40 °C (Ogbonda et al., 2007). It seems that in
many microalgal species temperature changes from low to high increases
the lipids productivity. High temperature also favours biofuel properties.
We have also noted that temperature stress is strain dependent and all
strains do not thrive equally at high temperature. Therefore,
temperature is a crucial stress factor to be taken in to account for
optimising lipids and biomass productivity.
Chapter 1
25
1.5.2 Light
Light can bring significant changes in the chemical composition of
microalgae. It is essential for photosynthesis, and together with
photoperiod is a critical factor for microalgal growth (Wahidin et al.,
2013). Light intensities of 100–200 μE/m²/s are commonly used for
microalgal production (Zhao and Su, 2014). For example, increasing the
light intensity f r o m 200 μE/m²/s to 400 μE/m²/s increased t h e rate of
growth in microalgae (Radakovits et al., 2010). In Mychonastes
homosphaera, Chlorella vulgaris, Raphidocelis subcapitata and
Scenedesmus sp. (Tang et al., 2011; Goncalves et al., 2013 and Liu et
al., 2012), lipid productivity increased with increase in light intensity
from low to high as shown in Table 1.2. Khotimchenko and Yakovleva.
(2005) reported high content of PUFA under low light intensity
whereas high light intensity resulted in greater accumulation of
saturated FAs and monounsaturated FAs. Optimum light requirements
varies with the microalgal species, and several parameters need to be
considered in selecting the right combination when using artificial light in
view of the overall energy balance considerations. It may be feasible to
use high light intensities for enhanced production of lipids, biomass
as well as suitable fatty acid profile for improving biofuel potential.
Lipids productivity is therefore found to be influenced under high light
stress. A serious concern is that the cells experience photoinhibition at
high light. Thus cultivation of microalgal strains in a two stage system,
where in first phase cells are cultivated under low light and then shifted
to high light in second phase can overcome the limitation caused by high
light alone. This strategy may improve the overall biomass and lipid
Chapter 1
26
productivity and addresses appropriately the concern of photoinhibition.
1.5.2 Nitrogen sources Of all the macronutrients in the culture medium, nitrogen, which
accounts for 1%–10% of the total dry matter in microalgae (Wijffels
et al., 2010), is quantitatively the most important nutrient affecting
growth and lipid accumulation in various algae (Griffiths and Harrison,
2009). Nitrogen in different forms such as ammonium, nitrate, yeast,
peptones, and urea, when used as a nutrient supplement, changes the rate
of cell metabolism significantly (Chen and Chen, 2006) and plays a
significant role in lipid synthesis (Figure 1.2). Increased lipid
accumulation under nitrogen starvation conditions in several microalgae is
well documented. For example, under nitrogen starvation, lipid
production increased about twofold and onefold in Neochloris
oleoabundans and Nannochloropsis sp. F&M-M24 (Li et al., 2008b;
Rodolfi et al., 2009), respectively. Nitrogen deficiency also affects the
biosynthetic pathway of carbohydrates (Cheng and He, 2014). For
example, carbohydrate content increased fourfold in Tetraselmis
subcordiformis and by 29% in S. obliquus CNW-N under nitrogen
deficient conditions (Ji et al., 2011; Ho et al., 2012). Restricted supply of
nitrogen and phosphorus shifts lipid synthesis from membrane lipids to
neutral lipids (Juneja et al., 2013).
Moreover, lipid productivity increased with increasing nitrate
concentration in Auxenochlorella pyrenoidosa (Singh et al., 2011b) and
Scenedesmus sp. (Xin et al., 2010), but the exact opposite pattern was
observed in A. protothecoides (Shen et al., 2009) and C. vulgaris (Prîbyl
et al., 2012), in which lipid productivity was higher at lower
Chapter 1
27
concentrations of nitrate (Table 1.1). A serious concern about
cultivating microalgae under nitrogen limiting condition lowers biomass
and consequently decreases the overall lipid productivity.
Two nitrogen limitation strategies has been defined to boost the lipids
production in several microalgae. The first one is two stage nitrogen
depletion as shown in C. vulgaris AG10032 (Mujtaba et al., 2012) in
which microalgal cells are grown in nitrogen rich conditions to stimulate
biomass growth and then transferred in to nitrogen starved medium for
lipids production. The second one is one stage cultivation in which the
medium contains desired nitrogen concentration and then as culture
grows the nutrient starts depleting and finally starved conditions is
achieved (Yen et al., 2013). The lipid and biomass accumulation in
both of these strategies is species-dependent.
Thus, nitrate levels must be controlled throughout the growth in
culture to maintain the nutrient contents of a medium stable and thereby
prevent damage to the cell membrane, loss of biomass, and other
changes in cell composition. Thus, screening based on lipid productivity,
biomass production, and FAME profiles among strains grown under
common parameters can achieve the highest resource utilization
efficiency and maintain a positive material and energy balance under low
cost nitrogen starvation technique. It is clear that under nitrogen
limitation, the lipid production is increased in all microalgal species.
However, biomass limitation under nitrogen deficient conditions is major
concern, as lower biomass tends to reduce the net lipid productivity. Thus
cultivating microalgae in two phase system as described above can
overcome the limitation of biomass production.
Chapter 1
28
1.5.3.1 Cow urine
Cow urine was selected as nitrogen source because of low cost which can
be advantageous for commercial production. However, the use of cow
urine for enhancing lipid and biomass productivity is limited but it
can be beneficial for commercial exploitation of biodiesel due to the low
cost of this nitrogen source. Cattle urine is a rich source of nitrogen and
other micronutrients; total N ranging from 6.8 to 21.6 g/L, of which an
average of 69% is present in the form of urea (Guschina and Harwood,
2006). Chapter 4 in this thesis describes research on the cultivation of
Scenedesmus bijugus and Chlorella sp. using cow urine.
1.5.4 Phosphate Phosphorous makes up considerably less than 1% of total algal
biomass (approximately 0.03%–0.06%) and yet is an essential
component in the medium for sustainable growth and development of
microalgae (Hu, 2004; Hannon et al., 2010). Absence of phosphorus in a
medium results in repression of photosynthesis (Belotti et al., 2014) and
affects the growth of microalgae. However, phosphorus has a significant
impact on the growth and metabolism of microalgae (Fan et al., 2014). It
has been observed that limitation of phosphates resulted in
increased lipid accumulation in microalgal cells. For example, in
the freshwater microalgae Scenedesmus sp. LX1 (Xin et al., 2010) and
Botryococcus braunii (Venkatesan et al., 2013), net lipid productivity
increased under phosphorus-limited (stress) conditions. Another study,
by Markou et al. (2012b), showed that under phosphorus-limitation,
carbohydrate content increased from 11% to 67% in Arthrospira
Chapter 1
29
(Spirulina) platensis (Table 1.2). Thus, while designing the
strategies to achieve higher lipid and biomass under environmental
stresses combination of low nitrates and optimal phosphate
concentration may increase the lipid and biomass production in one
growth cycle. Developing such strategies in future can make the process
more cost effective.
1.5.5 Salinity Salinity is a stress factor that causes biochemical changes in the growth
of microalgae (Kan et al., 2012). Recently, use of salinity stress in
microalgae has received attention in terms of the effect on growth and
lipid production (Asulabh et al., 2012). The ability of microalgae to
survive in saline environment under the influence of several osmotic
stress can affect cell growth and lipid formation. Numerous studies
reported that using NaCl as a stress factor in growth medium
increased total lipid content (Rao et al., 2007; Salama et al., 2014).
Increase in salinity can also increase lipid productivity in Scenedesmus
obliquus (Kaewkannetra et al., 2012). Microalgal species can tolerate
salinity stress up to an extent. Effects of salinity on marine algae have
been examined by several researchers (Bartley et al., 2013; Martinez-
Roldan et al., 2014) however limited reports are available on freshwater
microalgae. Dunaliella sp. is one such species that can tolerate high
salinity stress and produce large quantities of lipid and biomass (Azachi et
al., 2002). For example, in D. tertiolecta ATCC 30929 under high salinity
concentration, increased lipid content up to 70% (Takagi et al., 2006). In
the freshwater alga Scenedesmus sp., NaCl is believed to stimulate higher
lipid production (Salama et al., 2013). However, excess salinity stress in
Chapter 1
30
cultivation medium inhibits photosynthesis which further reduces the
biomass and net lipid productivity.
As discussed above, salinity stress tends to remarkably affects the fatty
acid profile of microalgae. The changes in each fatty acid chain is
species specific. Thus cultivating microalgae in combination with salinity
and nitrate stress in lab scale can further enhance the production of
lipids and cell biomass i n a single life cycle. A correlation between
changes in the production of lipids and photosynthetic efficiency has not
been well studied across diverse microalgal strains.
Adjusting the concentration of NaCl and light intensity can also impact
lipid production, biomass yields and fatty acid ratios related to biofuel
utility, in a diverse range of microalgal species. The effects of NaCl in
combination with those of other stress factors therefore need to be
examined further.
1.6 Carotenoids produced by microalgae 1.6.1 Bioactive compound Microalgae are known to produce pigments and bioactive compounds.
The major pigments produced by microalgae include chlorophyll,
phycobilins and carotenoids (Abalde et al., 1995). Chlorophyll is
mainly found in algae, cyanobacteria and higher plants. Carotenoids
are divided in to primary and secondary carotenoids. Lutein is
considered a major carotenoid since it acts as a primary carotenoid
maintaining the membrane integrity of cells and protecting cells from
many forms of stress (Sanchez et al., 2008), whereas astaxanthin is
considered a secondary carotenoid, present in lipid bodies outside
Chapter 1
31
the chloroplast (Grünewald et al., 2001), and has potential applications
in human health. Astaxanthin and canthaxanthin are considered major
ketocarotenoids in algae, fungi, and bacteria (Abe et al., 2007) and
therefore of great interest for biotechnological applications and as feed
supplements in aquaculture (Yaakob et al., 2014).
In plants and algae, carotenoids are synthesized in the chloroplasts around
the nucleus and their accumulation is effected only when they are exposed
to some stress factors (Lemoine and Schoefs, 2010). Astaxanthin
accumulation in lipid bodies outside the chloroplast increases under
several stress conditions (Wang and Peng, 2008). Phycobiliproteins are
water soluble pigments found in cyanobacteria and eukaryotic algae
(MacColl, 1998). The above pigments are high-value compounds,
produced from Porphyridium cruentum, Synechococcus sp., and
Chlorella sp. (Rodrigues et al., 2014). Due to their colour, carotenoids
are used in dyes and as colouring agents (Yuan et al., 2011; Christaki et
al., 2013).
Astaxanthin, β-carotene, and lutein are the main promising microalgal
pigments with high market potential (Table 1.1). The above pigments
are vital for the survival of a cell because they form the basic and
functional components of photosynthesis in the thylakoid membrane
(Guedes et al., 2011). Other pigments such as xanthophylls, mostly
found in cyanobacteria, are well linked with chlorophyll-binding
polypeptides (Grossman et al., 1995); however, in a few green
microalgae, xanthophylls are localized, and synthesized within the
plastid, whereas in Haematococcus sp., astaxanthin accumulates in the
cytoplasm (Eonseon et al., 2003). Carotenogenesis pathways and
Chapter 1
32
relative enzymes have been studied in cyanobacteria (Takaichi and
Mochimaru, 2007). The enzymes and genes involved in carotenoid
synthesis are different in algae and cyanobacteria (Takaichi, 2011).
Haematococcus pluvialis, D. salina, and Chlorella sp. are well-studied
candidates for the production of commercially important carotenoids
(Gimpel et al., 2015) (Table 1.1). Furthermore, in photosynthetic
organisms these carotenoids play important roles in either absorbing
energy or protecting cells from photo-oxidative damage (Demmig-
Adams and Adams, 2002). Unfavourable environmental conditions such as
nutrient deficiency, intense irradiation, and excessive photosynthesis
lower the rate of electron transfer, and in turn, photo-oxidative damage
(Solovchenko et al., 2011) as shown in Figure 1.5. Although the
pigments are basically associated with exposure and duration of light
intensity. The production of metabolites under dark conditions is yet
to explored.
Figure 1.4 Carotenogenisis in Haematococcus (Boussiba., 2000)
Chapter 1
33
1.6.2 Strategies to enhance microalgal carotenoids production Secondary carotenoids synthesis is altered by changes in culture
conditions and their production can be enhanced by controlling several
stress factors (Lemoine and Schoefs, 2010) as shown in schematic
Figure 1.4. Carotenogenesis is enhanced by reactive oxygen species
(ROS), under stress conditions such as high light intensity, salt stress
(Kobayashi et al., 1993). Because of this, astaxanthin is believed to
protect the body from such free-radical-linked diseases as oral, colon,
and liver cancers (Guerin et al., 2003).
Several unfavourable environmental conditions such as nutrient
deficiency, intense irradiation, and excessive photosynthesis lower the
rate of electron transfer and, in turn, photo-oxidative damage
(Solovchenko et al., 2011). Primary carotenoids, such as lutein degrade
under stress and therefore their biomass content is decreased. However,
several primary carotenoids, such as β-carotene act as secondary
metabolite and therefore accumulate under stress conditions (Rabbani et
al., 1998). Carotenoids content and accumulation in high concentration
in total biomass varies under several stress factors. Combined effect of
several stress factors have enhanced the production of astaxanthin in
H.pluvialis (Ben-Amotz, 2004).
Two stage cultivation process (Aflalo et al., 2007), continuous process
(Zhang et al., 2009) have been opted by researchers for increasing the
astaxanthin production in microalgae. Under stress conditions, β-
carotene is accumulated in lipid bodies with in the chloroplast with
highest content of 12% dry cell weight (Del campo et al., 2007).
Similar to astaxanthin, β-carotene content in microalgal species
Chapter 1
34
increases when growth is arrested under several stress factors (Lamers
et al., 2012). Lutein production in Muriellopsis sp. and Scenedesmus
almeriensis is unaffected under stress conditions (Table 1.2) unlike
astaxanthin and β-carotene (Del campo et al., 2000). However, these
two species have been tested in large scale cultivation system for
commercial purpose (Fernández-Sevilla et al., 2010). Autotrophic and
mixotrophic cultivation has been studied in detailed below to develop
the method for enhanced production of metabolites.
1.6..2.1 Autotrophic growth factors: Temperature Temperature plays an important role in the accumulation of carotenoids
within microalgal cells. Higher temperatures result in increased
accumulation of carotenoids in microalgal species due to increased
photo-oxidative stress (Tripathi et al., 2002). Temperature affects the
synthesis of astaxanthin in H. pluvialis (Chekanov et al., 2014) and
Chromochloris zofingiensis (Del Campo et al., 2004) as depicted in Table
1.2. High temperature is rarely used for inducing astaxanthin production
because it is reported to reduce biomass yield drastically, thereby
leading to an overall reduction in astaxanthin production.
Temperatures above 28°C were found to decrease the overall
productivity of astaxanthin in C. zofingiensis (Del campo et al., 2004)
because ROS are assimilated in greater quantities during photosynthesis,
which causes the accumulation of astaxanthin (Kovacic, 1986). In
Dunaliella salina, high temperature results in accumulation of lutein
(García-González et al., 2005). In addition, in S. almeriensis the lutein
content and net biomass increased to 0.46% and 0.53 g/L, respectively,
Chapter 1
35
when temperatures reached up to 30 °C (Sánchez et al., 2008). A similar
trend was also observed in C. protothecoides (Shi et al., 2006) (Table
1.2). In contrast low temperature increased β-carotene and α-carotene
content in Dunaliella sp. (Gómez and González, 2005). However, overall
lutein content was affected when temperature was increased from 20 °C
to 33 °C in Muriellopsis and S. almeriensis (Del Campo et al., 2007) in
one growth cycle (Table 1.2).
Temperature is the only factor which enhanced the production of lutein.
Temperature effect on carotenoids production is not well known,
however it is observed high temperature tends to accumulate more
lutein content whereas it has no effect on astaxanthin and β-carotene
production. Autotrophic cultivation of microalgae under temperature
stress is species dependent. For enhanced production of carotenoids,
cultivating microalgal strains at high temperature in combination with
other stress factors such as light and salinity could be more effective.
1.6.2.2 Light Light availability is one of the most serious limiting factors for production
of several carotenoids, biomass (Cordero et al., 2011), and FAs (Ward and
Singh, 2005). Little is known about carotenoids metabolism as affected
by the quality of light (Fu et al., 2013). Light intensity as well as
photoperiod affect growth, biomass, and other metabolites of interest in
diverse microalgae species (Seyfabadi et al., 2011; Khoeyi et al.,
2012) . Increasing the light intensity resulted in a threefold increase in
astaxanthin in (Table 1.2) H. pluvialis (Del Campo et al., 2004). Besides
astaxanthin, β-carotene content was also increased in D. salina up to
3.1% of dry cell weight (Table 1.2) when light intensity was changed
Chapter 1
36
from 100 to 1000 μmol photons/m²/s (Lamers et al., 2010). Lutein
content is also affected under high light intensity; for example, in
Muriellopsis sp. lutein content peaked at 460 μmol photons/m²/s. High
light intensity alone helps in promoting the growth of microalgae but
reduced overall lutein content in Scenedesmus sp. (Xie et al., 2013). In
all the above species, greater exposure to light increases the concentration
of astaxanthin and lutein.
The production of lutein was studied in C. sorokiniana (Cordero et al.,
2011) and Scenedesmus sp. (Sánchez et al., 2008) because both the
species have a high growth rate and high lutein-accumulating power.
On the other hand, formation of primary β-carotene and SC is affected
by low as well as by high light intensity: although photosynthesis occurs
under both levels, under low light intensity it generates fewer oxygen
radicals, whereas under high light intensity, cells are unable to utilize all
the energy that is generated, and the surplus energy activates more
oxygen molecules, leading to the formation of SC such as astaxanthin
(El-Baz et al., 2002). Such an exception has not been reported in H.
pluvialis, in which cells continue to generate oxygen molecules during
photosynthesis. In summary, carotenogenesis in photosynthetic cells is
correlated with different developmental changes and depends upon
environmental conditions. It is observed that, high light favours the
production of astaxanthin and β-carotene. Cultivation of microalgae
under high light stress in combination with nitrates can further enhance
the production of biomass and carotenoids (like astaxanthin, lutein
and β-carotene).
Chapter 1
37
1.6.2.3 Nitrogen sources Nitrate as a stress factor influences not only lipid production but also
affects carotenoids accumulation in green algae. Several factors trigger
carotenoids synthesis and should be considered in developing
microalgal products. Limiting nitrogen in the growth medium increases
the production of SC but lowers overall biomass yield (Cordero et al.,
2011). Ammonium nitrate have less impact on microalgal growth and
carotenogenesis (Markou and Nerantzis, 2013). Moreover, total
astaxanthin content of 3% dry cell weight (Table 1.2) was obtained in
a nitrogen-deficient medium (Imamoglu et al., 2009). Absence or low
levels of nutrients including nitrogen stimulate rapid physiological
responses, which further trigger the secondary biosynthetic pathways
(Touchette and Burkholder, 2000). Lutein accumulation in microalgae
cells is highly dependent upon nitrogen concentration within the
medium (Del Campo et al., 2000). Under nitrogen-limiting conditions,
carotenoids (β-carotene, astaxanthin, and canthaxanthin) begin to
accumulate in aerial microalgae (e.g. Coelastrella sp. and many more),
and the colour of cells changes from green to red (Abe et al., 2007).
Chromochloris zofingiensis, however, is an exception, and accumulates
lutein and astaxanthin as the main compounds (Table 1.2). Besides
lutein and astaxanthin, β-carotene content increased up to 2.7% of dry
cell weight in D. salina (Lamers et al., 2012) under nitrogen-depleted
conditions (Table 1.2). In general, nitrogen deficiency has a greater
impact than excess nitrogen on carotenoids production, mainly that of
astaxanthin in H. pluvialis (Table 1.2), because a culture growing in a
nitrogen- rich medium requires carbon to assimilate the nitrogen. Low
Chapter 1
38
nitrogen, on the other hand, leads to greater competition for carbon,
which is required for the synthesis of both carotenoids and proteins
(Borowitzka et al., 1991).
Hence a combination of urea and other nitrogen sources achieved maximal
lutein production in Auxenochlorella protothecoides under heterotrophic
conditions (Shi et al., 2000). A two stage system where both nitrogen
enriched and deficient medium is used in presence of high light
intensity stimulate the growth of microalgae during first phase with
biomass enhancement and in second phase deficient medium enhances
carotenoids enrichment. These observations could be vital considerations
towards defining future cultivation strategies.
1.6.2.4 Salinity Salinity has diverse effects on species and varies greatly across habitats
(Table 1.2). The effect of salinity on growth and carotenoids formation is
complex. Several species of microalgae can tolerate a range of salinity
levels because of efficient osmoregulation, which involves continuous
synthesis of glycerol (Pick, 2002). For example, carotenoids production
differs among the four species of Dunaliella, namely D. salina, D.
bardawil (Gomez et al., 2003), D. tertiolecta, and D. viridis (Hadi et
al., 2008): D. tertiolecta produces greater quantities of carotenoids at
optimum salinity levels whereas D. viridis does so only at higher salinity
levels. However, salinity affects overall β-carotene content (Coesel et al.,
2008). Salt stress of 5.5 M increased carbohydrate content in D. salina (a
2.5-fold increase) (Mishra et al., 2008). Increase in salinity increases
nitrogen and carbon content, which forces the cell to produce glycerol
and amino acids to cope with the high salinity (Jiménez and Niell,
Chapter 1
39
1991b). In Chromochloris zofingiensis, lutein and astaxanthin accumulate
only during later stages of growth because both are synthesized from a
common precursor, lycopene, with primary carotenoids as derivatives
(Hirschberg et al., 1997). The astaxanthin content increased under
salinity stress in H.pluvialis and C. zofingiensis (Table 1.2). However, H.
pluvialis cannot tolerate salt concentrations above 10% w/v (Borowitzka,
1991), whereas concentrations higher than 2 mM of NaCl decreased
total lutein content (Cordero et al., 2011) in Chlorella sorokiniana.
Salinity stress can further stimulate the production of total carotenoids.
However anything in excess causes harm to the metabolic system of
microorganism. Thus we inferred that NaCl stress have positive impact
on increasing the astaxanthin and β-carotene production however it has
no impact on the lutein production. Cultivating microalgal strains in the
medium containing high nitrates and high salinity together can increase
the production of lutein, astaxanthin and β-carotene ,while using a two
stage cultivation process. Combination of several environmental
stresses in one culture medium could become a novel strategy for
enhancing the metabolites production during single growth cycle.
However the metabolic pathway functioning of different stress factors
in diverse microalgal strains is a complex phenomenon and has not been
well explored and analysed.
1.6.2.5 Iron Iron is needed for the growth of microalga such as Dunaliella. Of all the
micronutrients, iron was the best for accumulation of astaxanthin in
cysts of H. pluvialis (Kobayashi et al., 1993) because iron acts as a
chelating agent; can scavenge hydroxyl radicals in the Fenton reaction
Chapter 1
40
widely used in the enzyme system of animals, microbes, and plants
(Raven et al., 1999); and acts as limiting factor under hypersaline
conditions. The site for assimilation of iron is usually the plasma
membrane (Polle and Song, 2009). He et al. (2007) reported that
phosphorus, iron, and sulfur are important nutrients for enhancing
astaxanthin accumulation. However, supplementing the medium with
Fe²+-EDTA (Table 1.2) produced biomass that contained 3.1% dry
cell weight astaxanthin in H.pluvialis (Cai et al., 2009). The different
stress factors used with different microalgae for producing lipids and
carotenoids are listed in Table 1.2.
.
Chap
ter 1
41
Tabl
e 1.
2 Ef
fect
of s
tress
fact
ors o
n lip
id a
nd b
ioac
tive
prod
uctio
n.
M
icro
-alg
al s
peci
es
Ope
ratio
n st
rate
gy
Net
lipi
d pr
oduc
tivity
(g
/L)
Ast
axan
thin
N
otes
R
efer
ence
Chl
orel
la v
ulga
ris
Hig
h lig
ht in
tens
ity
2664
–932
4 27
.0
Gon
calv
es e
t al.
2013
25
°C
; N
, 0.
15 g
/L
14.7
Om
ega-
3, 6
5%
Con
verti
et
al. 2
009
N
, 0.
61–2
.5 g
/L
5.5–
3.5
Prib
yl e
t al.
2012
Glu
cose
(20
.0 g
/L)
38.0
K
ong
et a
l. 20
11
Nan
noch
loro
psis
sp.
14–3
0 °C
7.
2–4.
7
H
u et
al.
2006
N
anno
chlo
rops
is o
cula
ta
15–2
0 °C
14
.9–7
.9
Con
verti
et a
l. 20
09
Scen
edes
mus
sp.
20–3
0 °C
11
.2–7
.4
Xin
et a
l. 20
11
Mon
orap
hidi
um sp
. 25
–35
°C
18.7
–14.
7
W
u et
al.
2013
Scen
edes
mus
sp.
Ligh
t int
ensi
ty 5
0–25
0 μm
ol/m
²/s
0.7–
1.60
Li
u et
al.
2012
N
aCl,
Hig
h lig
ht a
nd
0.1
3 g/
L N
aNO
3 59
.0 m
g/L/
d
0.59
mg/
g O
leic
aci
d 37
.1%
Peng
et a
l. 20
12
Myc
hona
stes
hom
osph
aera
Li
ght i
nten
sity
740
0–
29 6
00 lu
x 17
.4–3
1.2
Tang
et a
l. 20
11
Raph
idoc
elis
sub
capi
tata
Li
ght i
nten
sity
266
4–93
24
9.0–
39.0
G
onca
lves
et a
l. 20
13
B. b
raun
ii H
igh
light
inte
nsity
46
.8
Rua
ngso
mbo
on 2
012
18
°C
37.1
K
alac
heva
et
al
.200
2
18 °C
38
.3
Sush
chik
et
al. 2
003
Chap
ter 1
42
Mic
ro-a
lgal
spe
cies
O
pera
tion
stra
tegy
N
et li
pid
prod
uctiv
ity
(g/L
) A
stax
anth
in
Not
es
Ref
eren
ce
0.
1%
NaC
l; B
G11
m
ediu
m
Lute
in 8
4.3%
R
anga
rao
et a
l. 20
10
Sa
linity
: NaC
l +
sodi
um
acet
ate
15
.14%
Lu
tein
68.
4%
Ran
ga
Rao
et
al.
2010
Scen
edes
mus
sp.
30 °C
7.
4
X
in e
t al.
2011
N
, 0.0
02- 0
.025
g/L
5.
1–16
.2
Xin
et a
l. 20
10
N
, 1.5
g/L
24
.0
Rod
olfi
et a
l. 20
09
Auxe
noch
lore
lla p
roto
thec
oide
s N,2
.4- 6
.0g/
L 5.
9-4.
7
Sh
en e
t al.
2009
U
rea
1.0
g/L
9.1
Kon
g et
al.
2011
G
luco
se, 3
0.0
g/L
234.
0
H
ered
ia-A
rroy
o et
al.
2010
C
rude
gly
cero
l 14
.6
Che
n et
al
.
35 °C
Lu
tein
4.6
%
Shi e
t al.
2006
20g/
L N
aCl
43.4
%
Cam
penn
i' et
al.
2013
Scen
edes
mus
alm
erie
nsis
Hig
h te
mpe
ratu
re
Lute
in 4
.5 %
D
el c
ampo
et a
l. 20
07
30
°C
Lute
in 0
.46
%
Sanc
hez
et
al. 2
008
Mur
iello
psis
sp.
Hig
h te
mpe
ratu
re
Lute
in 4
.3%
D
el
cam
po
et a
l.200
7
Hae
mat
ococ
cus
pluv
ialis
A
dditi
on o
f Fe
²+ -ED
TA
3.1
%
Cai
et a
l. 20
09
H
igh
light
9.
5 μg
/mL
Tota
l fat
ty a
cid
133.
2 μg
/mL
Zh
ekish
eva
et a
l. 20
05
N
def
icie
nt +
ligh
t 3.
0 %
Im
amog
lu e
t al.
2009
Chap
ter 1
43
Mic
ro-a
lgal
spe
cies
O
pera
tion
stra
tegy
N
et li
pid
prod
uctiv
ity
(g/L
)A
stax
anth
in
Not
es
Ref
eren
ce
N
def
icie
nt +
ligh
t 21
.8 m
g/g
Oro
sa e
t al.
2000
H
eter
otro
phy
0.5
%
Kob
ayas
hi e
t al.
1992
b
H
igh
light
inte
nsity
and
m
ixot
roph
ic c
ondi
tions
10
%
Dom
ingu
ez-
Boc
aneg
ra
et a
l. 20
04
H
igh
tem
pera
ture
5.
5 %
C
heka
nov
et a
l. 20
14
N
, 1.5
g/L
65
.0 p
g/ce
ll
B
ouss
iba
and
Von
shak
19
91
0.8%
NaC
l 80
.0 p
g/ce
ll C
arbo
hydr
ate
48%
Bou
ssib
a an
d V
onsh
ak
1991
28°C
; 25
530
lux
98 m
g/g
Gar
cia-
Mal
ea e
t al.
2009
Ph
osph
ate
star
vatio
n
Saha
et a
l. 20
13
Scen
edes
mus
obl
iquu
s 0-
50m
M N
aCl
18-2
8 %
Ole
ic
acid
28.
3-41
.1%
Sala
ma
et a
l. 20
13
Nitz
schi
a la
evis
Ure
a+ N
itrog
en
EPA
175
mg/
L/d
Wen
and
Che
n. 2
001
Nan
noch
loro
psis
sp.
14 °C
7.
2
PUFA
11.
7%
EPA
7.9
%
Hu
et a
l. 20
06
22
°C
EPA
25.
3%
Hu
et a
l. 20
06
15
°C
14.9
C
onve
rti e
t al
. 20
09
N
, 1.5
g/L
13
.5
Rod
olfi
et a
l. 20
09
N
, 0.0
12 g
/L
13.6
H
u et
al.
2006
H
igh
CO
2 9
%
Hu
and
Gao
. 200
3
Chr
omoc
hlor
is z
ofin
gien
sis
24–2
8 °C
-
12.3
mg/
L Lu
tein
21
mg/
L D
el c
ampo
et a
l. 20
04
N
, 2.5
g/L
-
18.0
mg/
L Lu
tein
25
mg/
L D
el c
ampo
et a
l. 20
04
Chap
ter 1
44
Mic
ro-a
lgal
spe
cies
O
pera
tion
stra
tegy
N
et li
pid
prod
uctiv
ity
(g/L
) A
stax
anth
in
Not
es
Ref
eren
ce
H
igh
light
inte
nsity
-
19.0
mg/
L Lu
tein
24.
7 m
g/L
Del
cam
po e
t al.
2004
Ze
ro
salin
ity +
ligh
t -
15.0
mg/
L Lu
tein
15.0
mg/
L D
el c
ampo
et a
l. 20
04
G
luco
se,
- 10
.0 m
g/L
Su
n et
al.
2008
50
.0 g
/L
G
luco
se, 5
0.0
g/L
- 10
.2 m
g/L
Fe
ng e
t al.
2005
300
mM
NaC
l
0.8
mg/
L
Oro
sa e
t al.
2001
Chl
orel
la zo
fingi
ensi
s N
def
icie
ncy
65.1
%
Feng
et a
l. 20
12
P
defic
ienc
y 47
.7 %
Fe
ng e
t al.
2012
H
igh
light
inte
nsity
1.5
%
D
el c
ampo
et a
l. 20
04
0.
2 M
NaC
l
4 %
Del
cam
po e
t al.
2004
Mix
otro
phy
12
.5 m
g/L
Ip
et a
l. 20
04
H
eter
otro
phy
32
.0 m
g/L
Ip
et a
l. 20
04
Dun
alie
lla sa
lina
Hig
h li
ght i
nten
sity
-
β-
caro
tene
3.1
%
Lam
ers e
t al.
2010
N
, 2
.5 m
M 1
6% N
aCl
37.7
%
El-B
aky
et
al.
N st
arva
tion
β-ca
rote
ne 2
.7 %
La
mer
s et a
l. 20
12
Dun
alie
lla te
rtiol
ecta
N
, 0.0
26 g
/L; N
H4C
l (0
.16
m)
PUFA
73%
C
hen
et a
l. 20
11
C.v
ulga
ris A
G10
032
Two-
stag
e ni
troge
n de
plet
ion
77.1
mg/
L/d
Muj
taba
et
al
. 201
2
N. g
adita
na
Two-
stag
e co
ntin
uous
cu
lture
+ ni
trate
dep
letio
n) 51
.0 m
g/L/
d
Sa
n Pe
dro
et a
l. 20
13
Chap
ter 1
45
Auxe
noch
lore
lla p
yren
oido
sa
Sem
i- co
ntin
uous
(lo
w-N
m
ediu
m re
plac
emen
t) 11
5.0
mg/
L/d
Han
et a
l. 20
13
N
, 0–0
.4 g
/L
1.4–
6.66
g/L
S
ingh
et a
l. 20
11b
Chl
amyd
omon
as sp
. JSC
4 Sa
linity
: NaC
l 22
3.2
mg/
L/d
Ho
et a
l. 20
14c
Arth
rosp
ira
(Spi
rulin
a)
plat
ensi
s Ph
osph
ate
limita
tion
Car
bohy
drat
e 67
%
C
arbo
hydr
ate
67%
M
arko
u et
al.
2012
b
Chapter 1
46
1.7 Multiple factors that influence the synthesis of lipids and
carotenoids
The microalga has gained much attention as a rich source of carotenoids
and natural factories with industrial application in food and cosmetics
(Raina and Hala, 2008). The combined production of lipids and
carotenoids can make the process commercially feasible with maximum
utilisation of biomass. Besides lipids having biofuel properties, various
high- value added products such as vitamins, polysaccharides,
phycobilins, antioxidant can also be extracted from same biomass which
could be used as commodities in several biotechnology industries (Mata
et al., 2010). For example, polysaccharides which are produced by several
microalgal species has potential use in health industry (Markou and
Nerantzis, 2013). The combined production of lipids and carotenoids
from the same microalgal biomass therefore seems promising (Table 1.2).
Members of Chlorella sp. well known for lipids production, being now
used in cosmetic industry for their protein extract (Stolz and
Obermayer, 2005). Besides PUFAs, several value added products i.e.
carotenoids, astaxanthin, phycocyanin, terpenes, indoles etc. can be
extracted from algae (Bellou et al., 2014). Furthermore, C. vulgaris
extract is used in antiaging creams and tissue regeneration (Spolaore et
al., 2006) and C-phycocyanin extract from A.patensis has been used
against chronic myeloid leukemia (Nishanth et al., 2010; Subhashini et al.,
2004). The above compounds possess antioxidant, anti-inflammatory and
antifungal properties (Cardozo et al., 2007; Sekar and Chandramohan,
2008). Data from the literature shows as depicted in Table 1.2 that lipid
Chapter 1
47
synthesis is dependent upon carotenoids accumulation (Zhekisheva et
al., 2005) when exposed to high light. In H. pluvialis, carotenoids
production is dependent upon TAGs production. However, astaxanthin
accumulation within these TAGs bodies involves the transfer of
intermediates from the early steps of the biosynthetic pathway in
chloroplasts (Grunewald et al., 2001). Daily consuming microalgae
derived astaxanthin may protect body from oxidative damages, since
astaxanthin has more fighting capacity (500 times) more than that of
vitamin E (Duffosse et al., 2005). In addition to astaxanthin
accumulation, Haematococcus pluvialis (Zhekisheva et al., 2005) and
Scenedesmus sp. (Peng et al., 2012) have the potential to produce
multiple biomolecules a t the same time (Table 1.2). However, a
combination of several stress factors may results in the enhanced
production of carotenoids and lipids from the same biomass. For
instance, in D.salina combined the use of light stress apparently
increased the production of β-carotene (Table 1.2) (Lamers et al., 2010).
Another study conducted by Peng et al. (2012) showed accumulation of
lipids and carotenoids under salinity and light stress. The emerging value
of a few carotenoids (especially lutein) in the prevention and treatment
of disorders, such as degenerative diseases, and of omega FAs (EPA
and DHA) as a general wellness product, has strengthened the
biorefinery model. Microalgal pigments are gaining interestdue to their
wide used including treatment of chronic diseases, arthritis and cataract
(Sies and Stahl, 2004). Despite the promising cultivation conditions for
production of multiple metabolites from microalgae, the process has not
Chapter 1
48
been yet commercialised due to lack of reactors and methods that can
produce large amount to supply to market. In some cases, because of
limited production of biomass under stress, the overall productivity of
compounds is decreased (Adams et al., 2013). However, such negative
effect could be countered by applying a combination of stress factors
along with the two-stage strategy employed in a given life cycle in
photoautotrophic, heterotrophic, or mixotrophic condition. High light,
nitrogen deficiency and high salinity could influence the production of
lipids and carotenoids within one life cycle if two stage strategies are
followed with attenuated physio-chemical factors employed. It is clear
that different strains of microalgae respond differently to several
environmental stress factors (Table 1.2). Based on the literature
reviewed it is understood that nitrogen deficiency will results in
accumulation of higher lipid content whereas salinity impacts positively
the fatty acid profile and carotenoids production in diverse microalgal
species.
Simultaneous production of lipids, biomass and carotenoids may be
achieved through two stage process where growth medium rich in
nitrogen and phosphates in first phase for attaining higher biomass and
in second phase post pre-stationary high salinity and high light intensity
can further enhance the production of net lipids and carotenoids. The
adoption of above strategies in closed systems could achieve
economically sustainable process. These strategies can increase the total
productivity of the metabolites and lipids in the same biomass obtained at
the end of growth cycle. The exact combination of different stress
Chapter 1
49
factors may vary with the species drawn from different climatic
conditions. To date this concept of simultaneous production of lipids and
carotenoids from one life cycle and two stage process by employing
several rate limiting factors have not been fully explored and needs
to be pursued emphatically in future.
1.8 Solvent compatibility studies for algal milking of live cells during
continuous fermentation.
Current methods on extraction of lipids (Cooney et al., 2009; Herrero et
al., 2006) and harvesting of algal cell biomass requires lot of energy
which makes the process economically not feasible in terms of energy
inputs (Chisti et al., 2007). Unlike the aforementioned methods, milking
of simultaneous production of lipids and pigments have been reported
(Hejazi et al., 2004). Milking of lipids from microalgae has several
advantages i.e microalgal lipids can be easily milked in to organic phase
without dewatering, algae can be maintained in a nitrogen-limiting
condition which can increase the lipid yield up to 60-70% of dry cell
mass (Zhang et al., 2011). Nevertheless, only two microalgal species
Dunaliella salina and Botryococcus braunii has been studied and tested to
produce metabolites in presence of organic solvents such as dodecane and
hexadecane (Hejazi et al., 2004, 2002). Milking process integrate
harvesting, dewatering together which makes the process more
economically attractive. Moreover, milking from diatoms is been gaining
interest (Ramachandra., 2009). Moreover, milking from diatoms is been
gaining interest (Ramachandra., 2009). Although milking seems to be so
Chapter 1
50
efficient process for extraction of metabolites however few problems need
to be addresses. Firstly the cells has to be viable and recycled in to two
stage system to produces metabolites in continuous mode (Mojaat et al.,
2008). Secondly, the lipid extractability and recovery should be high from
economic point of view.
Compatibility studies between solvent and algal strains are important
for effective extraction of various compounds from living cells using the
method known as ―algal milking (Hejazi and Wijffels, 2003) An
important consideration is the structure of cell since they must be
solvent tolerant and also solvent must be able to extract materials
through the cell wall (Hejazi and Wijffels, 2004). Testing the effect of
organic substances on the permeabilization of cell membranes of specific
species of interest is necessary in early stage experiments (Lerner et
al., 1978). The permeability of these membrane depend on the
hydrophobicity of the solutes (Lieb et al., 1986) or polar (hydroxyl
groups) and charged (anionic and cationic) molecule.
Polar organic solvents like ethanol, acetone and dimethyl sulfoxide can
increase oil extraction yields, particularly yields of polar lipids (Dombek
et al., 1984). Variation of temperature can also result in changes in the
fatty acid composition in extracted oil (Suutari et al., 1990). Leon and
co-workers (2001) showed that organic solvents are often toxic to
photoautotrophic unicellular microorganisms (Hejazi and Wijffel, 2003).
The toxicity of organic solvents depend on two major factors, the
dispersibilty of solvent in water and the compatibility of the solvent with
cells at high agitation speeds (Sikkema et al., 1995). Hejazi et al. (2002)
should that determining the toxicity of alkanes to the cell is a good
surrogate for overall solvent biocompatibility.
Chapter 1
51
The log P value must be greater than 5 for any solvent, because the
higher the log P value the less water-soluble the solvent and therefore
fewer molecules are required to cause toxic effect (Basseti et al., 1994)
(Leon et al., 2001). Solvent studies on Dunaliella salina showed that
this organism can remain active in aqueous/organic solvent mixtures
(Hejazi et al., 2003).
Leon et al. (2003) showed that the solvent decane, at log P of 6 for
more than 72 h, is suitable for β-carotene extraction from Dunaliella
salina because β-carotene is very hydrophobic. For categorizing solvents
in relations to biocompatibity, Bar et al. (1987) suggested that two types
of toxicity can appear in cultures. One is molecular toxicity, which
appears when a solvent is dissolved in an aqueous phase and the
other is phase toxicity, which occurs when a second aqueous phase
appears. When larger quantities of cell biomass were contacted with
hexane or other organic solvents the efficiency of recovery dropped due
to a poor contact, as a result of increased aggregation in hydrophobic
solvent. Recovery yields can be influenced by the physiological stage of
the extracted culture.
The mechanism of extraction of products from biocompatible solvents
has been investigated. Alteration in the cell membrane occurs after
exposure to biocompatible solvents (Sikkema et al., 1995 and Hejazi et
al., 2004) because these solvents get incorporated into cell membranes.
For example, in Dunaliella salina β-carotene accumulation occurs inside
chloroplast in the form of oil globules (Ben-Amtoz et al., 1995). In
Chapter 1
52
general to conduct any experiment one must select an organism on the
basis of cell wall structure, location and style of accumulation. For
example, Haematococcus has a rigid cell wall, and accumulation of
astaxanthin occurs through different mechanism than that of β-carotene in
D salina (Lee et al., 1994).
1.9 Conclusion
Recent advances in biotechnology have created opportunities for the
production of high value added metabolites with unique benefits in
various industrial sectors.
Microalgae under optimal conditions demonstrate significant potential
to produce multiple metabolites (Table 1.2), many of which can be
produced simultaneously under one growth cycle. Producing lipids and
bioactive compounds from the same organism is necessary to enable
cost effective production of these compounds and associated biofuel.
The cultivation conditions outlined are based on the experimental data
of known microalgae but in principle could also be applicable for the
developing strains by use of genetic engineering. Genetic manipulation of
promising organisms can further lead to higher commercial returns from
those which are already preselected on the basis of physiological
performances. The advantage of using culturable biological resources
instead of refining the finite stocks of crude oil is that the processes will
improve as technology improves, while crude oil stocks will decrease and
costs rise over the longer term. This study defines many favourable
stress factors those influence the production of lipids and carotenoids.
Microalgae, under optimal conditions, demonstrate a significant potential
to produce multiple metabolites, many of which can be produced
Chapter 1
53
simultaneously in one growth cycle.
Exploring biodiversity and selecting potential strains that produce the
desired multiple products with maximum efficiency can pave the way
to implement the biorefinery approach cost-effectively. Genetic
manipulation of promising organisms can lead to higher commercial
returns from the strains chosen on the basis of their physiological
performance. The biorefinery approach provides new insights into the
feasibility of cost effective and continuous production of lipids and
bioactive compounds from one growth cycle and possibly the two-stage
cultivation process under altered physico-chemical conditions. The
advantage of using culturable biological resources instead of refining the
finite stocks of crude oil is that the processes will only get better with time
and will lead to better economics and efficiency i n tune with the growing
human needs.
Microalgal biorefinery process is still in the early stage development and
more focus is needed about extraction and purification of multiple
products besides lipids and carotenoids from the same biomass.
Therefore, more and consistent research efforts are needed to understand
the trade-offs between the lipid and carotenoids production and
identifying most useful metabolic engineering strategies under
optimised conditions to develop better equipped biorefinery
approaches. Hopefully, this review will provide new insights in
development of different cultivation strategies for enhancement of
biomass rich in lipids and carotenoids using combination of stress
factors in diverse microalgal strains. Further milking out the metabolites
from same cycle using biocompatible solvents without killing algal cells.
Chapter 1
54
1.10Aims of the study The primary objective of this study was to isolate novel and
functionally efficient isolates from the diverse habitats with multiple
product potential. The research aims of my PhD dissertation were as
follows:
1. The first aim was to collect samples from the different geographical
regions of India and the World, isolate and purify unicellular cultures
and carryout identification on the basis of morphometric and molecular
analysis.
2. The second aim was to assess the innate functional abilities of selected
isolates as well as distinguish them with those having multiple product
potential with desirable biofuel characteristics. This is a major scientific
question and commercial need in the current time
3. The third aim was to evaluate the influence of different
environmental stress factors on further enhancement of multiple
products found in various selected cultures
4. The final aim was to attempt or explore non-destructive extraction of
various products using biocompatible solvents and study cell recovery.
This is crucial for any future exploitation of commercial feasibility.
The aims of this study by in large includes following research hypothesis
those were addressed during the course of presented work.
1. There are potential isolates in natural ecosystem with multiple products
potential
2. There is a possibility to further enhance the naturally existing multiple
products potential by physiological attenuations
3. Once achieved maxima of product enrichment non-destructive recovery
will make process economically feasible for commercial exploitation in
future
Chapter 2
55
Chapter 2: Isolation, identification and preservation of new strains of
microalgae from different geographical region of India and the World.
2.1 Introduction Microalgae, widely distributed, and with a longer evolutionary history
than terrestrial plants, show a rich diversity among their more than 200
000 species (Guiry and Guiry, 2014). Approximately 30 000 of these
species have been studied but so far not fully exploited (Mata et al.,
2010). This extensive biodiversity indicates the rich potential of
microalgae as agents to produce a range of substances useful for many
industrial sectors. In the past few years these organisms have been
extensively explored for biofuel (Lim et al., 2012) and bioactive
compound production (Pulz et al., 2004; Wijffels and Barbosa, 2010).
Freshwater microalgae exhibit a great diversity and specific species
distribution depends not only on the chemo physical environment but
also on the surroundings to which species can colonise (Andersen,
2005). In recent years, there has been interest in the isolation of unique
strains from tropical areas such as freshwater, backwater and terrestrial
habitats (Minhas et al., 2016). Such extremes as very high temperatures
and prolonged exposure to intense light found in the tropics have conferred
on the extremophiles some distinctive physiological properties (Gouveia et
al., 2009). In tropical and subtropical regions, microalgae productivity
is greater than in temperate zones, due to higher growth levels at
elevated temperature (Franz et al., 2012). Various altitude and climatic
conditions may result in distinct isolates with industrial potential
(Schittek et al., 2016).
Chapter 2
56
Microalgae are difficult to isolate since they are slow growing organism
compared to other microorganism such as bacteria, yeast and fungi.
Traditional techniques for the isolation of microalgae (Beijerinck and
Miquel, 1890) with microscopic characterisation and cultivation in a
range of culture medium for both freshwater and terrestrial algae were
used. The use of molecular approaches to identification has resulted in
the identification of increased microalgal diversity (Edgcomb et al.,
2002; Kawai et al., 2003; Stoeck et al., 2006; Lopez- Garcıa and
Moreira, 2008; Zhao et al., 2012). The use of molecular techniques and
phylogenetic methods provides new insights into the evolutionary history
of microalgae (Leliaert et al., 2012). The investigation of DNA
sequences in green plants was first reported in the mid 1980‘s (Hori and
Osawa, 1987), followed by 18S rDNA sequence analysis (Mishler et
al., 1992). Subsequent use of PCR and bioinformatics techniques in the
1990‘s has made it easier to analyse and compare large sequences in
databases. However, DNA sequence data is still lacking for many
genera and species. 18S rDNA sequencing has been the primary
source of data for analysing phylogenetic linkages among the green
algae (Pr¨oschold and Leliaert, 2007; Fawley, 2000). PCR has been
applied to species identification in environmental samples (Dyhrman et
al., 2006). Simple microscopic observation of microalgal can be
taxonomically misleading because of similar morphological
characteristics (Hu et al., 2008). Our investigation describe collection,
isolation and purification of microalgae from diverse habitats and
geographic locations with the aim of understanding microalgae diversity
from these locations.
Chapter 2
57
Comparing the use of direct plating on algae media and nutrient agar plate
for the isolation and purification of unialgal isolates, as well as the
preservation of purified isolates in alginate beads is dicussed in this
chapter. Strains were further subjected to morphometric analysis using
scanning electron microscopy (SEM) and identification using molecular
techniques.
2.2 Materials and Methods 2.2.1 Microalgal sample collection A total of two hundred samples were collected from aquatic and
terrestrial habitats from Asia, North America, and the Middle East
(Figure 2.1). The aquatic habitats included freshwater, polluted river
water, and backwater and the GPS coordinates for all the location were
recorded. Samples were collected in sterile glass bottles and sieved 3–4
times using a nylon mesh with pore size of 33 μm at various point
(González et al., 2013). The samples were placed in sterile 20mL
plastic bottles containing diluted 30% formalin. The plastic bottles
containing the samples were transported to the laboratory of The energy
and resources institute, New delhi and stored at 4°C for 4 days for further
use.
Chapter 2
58
Figure 2.1 Map of World and India depicting sampling locations
Chapter 2
59
2.2.1 Isolation and purification of microalga In order to isolate a large number of strains from field water samples,
the standard dilution plating method as described below was used to
separate large populations. Multiple media compositions were used to
isolates the colonies.
2.2.2.1 Direct plating method All the primary stock cultures from field samples were incubated up
to the late exponential phase in full-strength Bold‘s basal medium
(BBM), for freshwater collections (Andersen, 2005), in 100 mL
Erlenmeyer flasks at 25 °C with orbital shaking at 150 rpm and under
16 h of light (100 μE/m2/s) and 8 h of darkness. After two weeks, the
pre-cultured samples were isolated by serial dilution. Dilution up to 10-5
and 10-6 were performed for all the samples and these were then plated on
sterile BBM agar plates (1.6 % w/v). Growth colonies on the plates were
observed at 24 h intervals. Individual colonies were taken using a sterile
loop and grown in a 50mL Erlenmeyer flask containing 10mL fresh
medium until sufficient biomass was produced. Following the individual
colonies, each strain was labelled based on the sampling location and
specific medium requirement. Grown algae cultures were streaked using
sterile technique onto nutrient media plates. Isolated algae were
maintained by re-plating on new nutrient media plate depending on the
nature of each isolated strains. Different medium (BG11 (Waterbury and
Stainer, 1981), Basal (Herediyo-arroyo et al., 2010) and BBM were
optimised for all the strains collected from diverse habitats.
Chapter 2
60
Grown cultures were purified by passing the sample through glass fibre
prefilters (Millipore, USA) in a vacuum filtration unit with repeated
washing with milli-Q water and intermittent washing with mild
disinfectant chloramine T (0.2% w/v) (Sigma Aldrich, USA).
Axenicity of the isolates was confirmed by spread-plating 100uL
suspension of treated algal cells onto a small nutrient agar plate. Plates
were incubated at 30 °C for 72 h. All microalgal isolates were also
tested under microscope during cell counting to make sure that none had
been contaminated with other microalgae isolates during handling.
Unialgal colonies were further maintained by sub-culturing on nutrient
agar plates to obtain pure isolates.
2.2.3 Long term preservation of microalgae The active stage grown microalgal cells were harvested by
centrifugation at 4000 rpm for 10 min and washed three times with
sterile distilled water. The cell pellet was mixed thoroughly with sterile
solution of sodium alginate solution (2% w/v). Beads of about 0.35 cm
diameter were obtained by dropping the alginate cell mixture into a
solution of 0.07M CaCl2 (pH 7.8) with a sterile syringe needle. Beads were
left in CaCl2 for less than 20 min to harden and were then washed in
sterile distilled water. Beads were distributed amongst test tubes
containing 10 mL of distilled water and incubated at 4°C at low light of
25μE/m2/s for 6 months.
Chapter 2
61
2.2.4 Characterization of isolates by scanning electron microscopy (SEM)
For examining the cultures under a SEM, 2 mL of the fully grown
culture was centrifuged at 6000 rpm for 5 min. Primary fixation of the
algal cells was performed by incubating them in 2.5% glutaraldehyde
at room temperature in the dark for at least 4 h followed by
secondary fixation in 1% osmium tetroxide overnight at 4 °C. The
algal cells were dehydrated gradually in increasing percentages of
ethanol (10%, 30%, 50%, 70%, 90%, and 100%), critical point dried,
sputter coated with gold palladium using a polaron SC7640 auto/manual
high resolution sputter coater (Quorum Technologies, Lewes, East
Sussex, UK) and examined under a FEI Quanta 200 scanning electron
microscope (FEI, Hillsboro, Oregon, USA) at 10 kV.
2.2.5 Cultivation of microalgal strains For DNA isolation, isolates labelled by specific codes collected from
different habitats were used. The isolates collected from aquatic and
terrestrial habitats were transferred in to 20mL BBM medium in 150mL
Erlenmeyer flasks kept in an orbital shaker at 25 °C at 150 rpm under
photoperiod of 16h of light and 8h of darkness. The isolates were allowed
to grow for approximately 10 days until reaching the exponential stage,
since all the isolates were expected to reach the exponential phase by 10
days. Therefore at day 10 the culture was harvested and used further for
DNA extraction.
Chapter 2
62
2.2.6 DNA extraction from diverse isolates For molecular confirmation of isolates, genomic DNA from the
macerated biomass was extracted with a DNeasy plant mini kit (Qiagen
GmbH, Hilden, Germany) following the manufacturer‘s instructions.
The extracted DNA was either used for polymerase chain reaction
(PCR) or stored at -20°C for further use. The genomic DNA containing
18S rDNA gene was amplified using two universal eukaryotic primers,
namely (EUF: 5′ GTCAGAGGTGAAATTCTTGGATTTA-3′ as the
forward primer and EUR: 5′-AGGGCACGACGTAATCAACG-3′ as the
reverse primer), which are expected to amplify the ~700 bp region of
18S rDNA (Rasoul-Amini et al., 2009). The extracted DNA was analysed
by electrophoresis on 0.8% (w/v) agarose gel (Agarose, Sigma-Aldrich,
USA) to confirm the presence of an appropriate concentration of
products. The extracted DNA was either used for polymerase chain
reaction (PCR) or stored in -20°C for further use. The DNA
concentration was measured using a Nanodrop 1000 Spectrophotometer
(Thermo scientific, Wilmington, DE, USA).
2.2.7 PCR amplification, and 18S rDNA sequencing The PCR reaction was performed in a total volume of 25 μL
containing 2 μL of extracted DNA as template, 200 μM of each dNTPs,
1.5 mM MgCl2, 0.5 μM forward & reverse primer and 0.5 U Taq DNA
polymerase. The amplification was performed in a ABI Veriti thermo-
cycler gradient with the PCR program conditions as described by
Ghasemi et al. (2008). Similarly, the PCR reaction was performed
using designed algae-specific primers, namely (FP1:5′ -
GCCCGTAGTAAWWCTASAGCTAATAC -3′ as the forward primer
Chapter 2
63
and RP1:5′- AAAAACGGGCGGTGTGTACAAAGGGC -3′) as the
reverse primer with the following conditions for the PCR: 4 min initial
denaturation at 95°C; 35 cycles of 1 min denaturation at 95°C, 1 min
annealing at 56°C and 2.5 min elongation at 72°C; and a 10 min final
elongation. The sequences of purified products (PCR purification kit,
Qiagen, Germany) were determined using an ABI 310 automatic
DNA sequencer (Applied Biosystems, California, and USA). The 18S
rDNA sequences were compared with published sequences from the
NCBI database to determine the nearest homologous sequences and the
relative phylogenetic positions of individual isolates by ML tree analysis
using MEGA ver. 6. In the present study, mapping o f t h e the
existing universal primers from the literature as well as self-designed
primers was done and evaluated them for their appropriateness for
studying eukaryotic biodiversity
2.2.8 Construction of degenerate algal-specific 18S rDNA primers To confirm the results obtained using the universal primers, algae-
specific degenerate primers were designed based on multiple sequence
alignment of the 18S rDNA sequences using the software package
BioEdit and Clustal W. The ribosomal encoded 18S rDNA gene
sequences from a large data set corresponding to different genera of algae
were obtained from NCBI. The specificity of the degenerate primer sets
was chosen for amplifying a ~ 1.5–2.8 kb length, depending on the algal
isolates. The specificity of these designed primers was also tested in silico
using NCBI-BLAST to determine the non-specific binding. The products
of PCR (amplicons) were purified using a gel extraction kit (Qiagen,
Hilden, Germany) and were cloned using the pGEM-T (Promega, USA)
Chapter 2
64
vector system following the manufacturer‘s instructions. Colony-PCR was
performed using universal M13 forward and reverse primers to
confirm the positive clones. Two individual clones from each isolate
were sequenced by Xcelris Genomics Pvt. Ltd (Ahmedabad, India). The
NCBI-BLAST program was used to determine the sequences showing
maximum homology.
2.2.9. Phylogenetic analysis To ascertain the phylogeny of the isolated microalgae, their 18S rDNA
sequences were aligned with the sequences of different closely related
species that have been reported earlier. Multiple alignments were
performed using Bio-Edit Sequence Alignment Editor ver.7.2.5. The
algorithms available in MEGA ver. 6, namely Maximum Likelihood
(ML), Neighbour-Joining (NJ), and Maximum Parsimony (MP), were
employed for constructing phylogenetic relationships among the algal
isolates, and the reliability of the branches was assessed by bootstrapping
the data with 1000 replicates.
2.3 Results and Discussion 2.3.1 Isolation of microalgae Several isolation method such as serial dilution plating on agar, single
cell isolation, gravimetric separation and flow cytometry methods have
been established (Andersen, 2005; Sinigalliano et al., 2009). Previously,
agar plating, serial dilution, use of antibiotics and ultrasonication have
been used for the purification of large microalgae (Brahamsha, 1996;
Sensen et al., 1993). However in our hands serial dilution followed by
plating on agar was an easy and cost effective method for the
purification of diverse microalgae (Pringsheim, 1946).
Chapter 2
65
Two hundred samples from aquatic, backwater and terrestrial habitats
from Asia, North America, and the Middle East were collected (Table
2.1). The isolates include many unicellular green algae.
Chap
ter 2
66
Tabl
e 2.
1 C
olle
ctio
n si
tes o
f mic
roal
gal i
sola
tes f
rom
div
erse
hab
itats
.
Reg
ion
and
coun
try
of
colle
ctio
n um
ber
of
isol
ates
Sa
mpl
ing
mon
th a
nd
Hab
itat
Col
lect
ion
mon
th
Lat
itude
L
ongi
tude
Mid
dle
Eas
t, Q
atar
15
A
ugus
t 201
2 B
ackw
ater
D
ecem
ber
25° 1
7' 7
" N
51° 3
1' 5
1" E
Nor
th A
mer
ica,
St.
Loui
s , U
SA
8 N
ovem
ber 2
012
Fres
hwat
er
Sep
tem
ber
38° 4
7' 5
1" N
90
° 47'
8"
W
Asi
a, H
ussa
in S
agar
, H
yder
abad
, Ind
ia
8 A
ugus
t 201
2 Fr
eshw
ater
Ju
ly
7° 2
5' 2
5" N
78
° 28'
25.
7" E
Asi
a, R
ocky
bea
ch, G
oa,
Indi
a.
7 M
ay 2
012
Bac
kwat
er te
rres
trial
, ro
cky
Oct
ober
15
° 17'
57"
N
74° 7
' 26"
E
Nor
th A
mer
ica,
Tre
e ba
rk,S
t. L
ouis
, USA
6
Dec
embe
r 201
2 Fr
eshw
ater
terre
stria
l, ro
cky
Dec
embe
r 38
° 37'
37"
N
90° 1
1' 5
7" W
Asi
a, C
henn
ai, T
amil
25
June
201
2 B
ackw
ater
M
ay
11° 7
' 37"
N
78° 3
9' 2
4" E
Asi
a, V
ishak
hapa
tnam
30
Ju
ly 2
012
Fres
hwat
er
July
41
° 42'
44"
N
81° 1
5' 1
4" W
Asi
a, P
ango
ng la
ke,
6 A
pril
2012
Fr
eshw
ater
S
epte
mbe
r 33
° 44'
16"
N
79° 0
' 27"
E
Asi
a, M
anga
lore
, 20
Ju
ne 2
012
Bac
kwat
er
May
12
° 54'
50"
N
74° 5
1' 2
1" E
Asi
a, Y
amun
a r
iver
, In
dia
60
July
201
2 C
hem
ical
ly p
ollu
ted
July
28
° 36'
50"
N
77° 1
2' 3
2" E
Asi
a, Y
amun
a ri
ver,
D
elhi
, Ind
ia
15
June
201
2 C
hem
ical
ly p
ollu
ted
river
wat
er
July
28
° 2' 5
6" N
79
° 29'
23"
E
Chapter 2
67
The collected samples contained contaminants such as bacteria, yeast,
fungi and mixed culture of algae. The samples stored at 4°C were
centrifuged to remove 90% of suspended materials and other diatoms.
The mixed green algae at the bottom of the tubes were collected and
those species were used for mixed cultures in the liquid medium. After
culturing the mixed cultures in different media (BG11, BBM, Basal
medium), it was found that BBM with soil extract was the richest
medium reused at the first stage of the isolation of green and brackish
water microalgae, with a pH of 6.8, because this pH reduced the risk of
other algal inhabitants in the mix culture. After 2 weeks it was
observed that only green microalgae were growing in the medium
(Figure 2.2).
Figure 2.2 Mixed partial purified cultures. All the microphotograph of
the cultures were observed under 100X (Carl Zeiss microscope-
Primostar). a) Around 5-6 species. b) Around 3-4. c) Around 2 (green
algae).
2.3.2Chemical treatment: Purification of microalga Chloramine T acts as a biocide and inhibits the growth of bacteria that
can damage DNA via oxidation, thereby inhibiting microbes from
Chapter 2
68
replicating (Shetty and Gowda, 2004). Chloramine T was used at different
concentrations (0.5 -3 % w/v) to remove any bacterial or fungal growth.
At 1.5% (w/v) it was effective for eliminating the growth of bacteria
and other contaminants
Figure 2.3 Pure cultures on nutrient agar (left side) and algae agar plates
(Right side)
2.3.3 Streak-plating and Serial dilution
A process of serial dilution proposed by Andersen. (2005) for the
isolation of unialgal isolates was effective in our experiments. Solid
plating attempted several times with dilutions of up to 10-5 and 10-6
and was useful for the separation of unialgal colonies (Figure 2.4).
Finally, single colonies were picked and inoculated in fresh BBM
medium and used as unialgal isolates for all the strains (Dilution up to
105 and 106). The colonies were streaked and maintained on agar plate
at 4°C for further use (Figure 2.5).
G7 (Goa) G7 (Goa)
Chapter 2
69
Figure 2.4 Formation of algal colonies on agar plates. (a) Dilution at 10-5,
(b) Dilution at 10-6
Figure 2.5 Maintenance of unialgal isolates: Streak plate
2.3.4 Preservation of microalgae The alginate preservation method was studied in cyanobacteria in the
laboratory by Romo and Perez-martinez. (1997). In this process, isolates
were prepared mixed with sodium alginate (2% w/v) solution by
dropping the solution in to a 0.07 M solution of CaCl2 and leaving for
20 mins to form alginate beads. This method was well suited for
preservation of freshwater and backwater isolates from different habitats.
Ladakh Ladakh
P63 P63
Chapter 2
70
The algae beads were stored at 4°C for approximately 6 months. In our
study we were able to achieve a revival percentage of 90% while growing
beads in fresh sterile BBM medium. However, initially immobilised
algae alginate beads took longer a time to dissolve in nutrient rich
BBM media. Our study report that revival was possible while
preserving algae in the form of alginate beads for 6 months (Figure
2.6).
Figure 2.6 Preservation of (a) Algal beads in sterile distilled water, (b)
Dried alginate algal balls
(a)
(b)
Chapter 2
71
2.3.5 Identification of diverse microalgal isolates using SEM and
molecular biological techniques
Two hundred samples were collected from aquatic and terrestrial
habitats from Asia, North America and the Middle East. A total of 22
isolates were found to be taxonomically distinct, based on 18S rDNA
analysis. Of these, five belonged to Chlorellaceae, consisting of one
(Q12) backwater isolate, three other backwater isolates, and one
freshwater isolate. Another freshwater isolate belonged to
Eustigmatophyceae and the remaining strains belonged to
Scenedesmaceae. Scanning electron microscopy imaging have been
conducted in the 22 selected species which showed the lipids and
biofuel potential as depicted in Figure 2.7a-k. Phylogenetic analysis
using 18S rDNA sequences was performed to further characterise these
isolates.
The distingusible and unique morphometric observations from twenty
two isolates are shown in Figure 2.7a-k. SEM micrographs reveal that
the vegetative cells of Chlorella sp. (Q12) belonging to the family
Chlorellaceae within the class Trebouxiophyceae were spherical (Figure
2.7a) to somewhat ellipsoidal, 4.7 μm in diameter, with thick and rough
outer cell walls. The features were similar to those observed in
Chlorella strain R02/6 (Gartner et al., 2015). In the green and unicellular
Chlorella sorokiniana, the cell size was 5.54 μm in an isolate from the
USA and 5.36μm in another one from Husain Sagar (HS), India (Figure
2.7b and 2.7c), and the cells were spherical with thick and rough cell
walls.
Chap
ter 2
72
Sim
ilar
feat
ures
wer
e re
porte
d by
Im
ase
et a
l. (2
013)
for
the
sam
e sp
ecie
s. In
the
iso
late
fro
m p
ollu
ted
wat
er,
nam
ely
Para
chlo
rella
kess
elri
(Y
8), b
elon
ging
to f
amily
Eus
tigm
atop
hyce
ae w
ithin
the
cla
ss.
Figu
re 2
.7 S
cann
ing
elec
tron
mic
rogr
aphs
of
mic
roal
gal
isol
ates
sho
win
g so
me
of
cell
shap
es o
bser
ved:
(a)
Chl
orel
la s
p. (
Q12
), (b
)
(c) (g) (k)
(i)
(d) (h)
Chapter 2
73
Chlorella sorokiniana (HS), (c) Chlorella sorokiniana (USA), (d)
Parachlorella kesselri (Y8), (e) Auxenochlorella protothecoides (Goa),
(f) Coelastrella sp. (V3), (g) Coelastrella sp. (Tree), (h) Scenedesmus
bijugus (Ladakh), (i) Scenedesmus sp. (V9), (j) Scenedesmus sp. (V15),
and (k) Scenedesmus sp.(V11).
For Chrysophyceae, the apices of cells were interconnected and
formed a chain (Figure 2.7d), a unique feature. In Auxenochlorella
protothecoides, cells were 5.18
μm in diameter, with thin cell walls covered with an irregular
network of ribs. Smooth- and rough-walled cells were present in the
same culture at maturity (Figure 2.7e). Huss et al. (2002) noted similar
features in C. protothecoides. In Coelastrella sp., belonging to family
Scenedesmaceae within the class Chlorophyceae, cells of isolate V3
were 7.03 μm in diameter and those of the isolate from a tree were 5.07
μm in diameter (Figure 2.7f and 2.7g). In both the isolates, cells were
lemon shaped and their walls were covered with ridges and fine ribs, as
in Coelastrella sp. (Hegewald et al., 2010), and some of the ribs did not
end at the cell pole. Whether these ribs have any function is not known;
however, they are an important morphological feature and typical of the
genus Coelastrella (Hu et al., 2013). Other groups of the genus
Scenedesmus showed a different cell ornamentation. The cells in almost
all the Scenedesmus sp. were spherical to semispherical and lacked
spines. Cells of S. bijugus (from Ladakh) averaged 4.61 μm in
diameter and showed a flattened surface in contact with sister cells
(Figure 2.7h), as reported in Scenedesmus sp. (Lewis and Flechtner,
2004). Isolates V9 (5.01 μm) and V15 (4.02μm) of Scenedesmus sp.
Chapter 2
74
showed an unornamented wall surface and wall partitioning (Figure 2.7i
and 2.7j), whereas isolate V11 of the same species showed knobby cell
apices (Figure 2.7k), as observed in other species of Scenedesmus (Lewis
and Flechtner, 2004).
New taxanomic tools such as SEM provides a new inights in to speciation
of algae. SEM method provides lot of new information. By using SEM
surface morphology of cells can be revealed (Kownacki et al., 2015). SEM
helps us to identify the species and find a new morphological feature.
2.3.6 Molecular identification of microalgal isolates using 18S rDNA
sequencing
Molecular techniques were used for the identification of organisms to
species level (Fawley, 2000). Sequence data has improved the ability to
identify a species that is difficult to identify using morphological
techniques (Lewis and Court, 2004; Fletchner, 2008). This has led to
more accurate identification of diverse algal species useful for biofuel
production (Georgianna and Mayfield, 2012). Confirming the true
phylogeny of an organism can be difficult without genetic information and
important for understanding the relationship of new strains to existing
organisms.
In the present study, amplification of 18S rDNA sequence was carried
out using both universal and self-designed primers. The amplification of
18S rDNA using universal eukaryotic primers showed efficient
amplification of a unique single band varying in size from 341 bp to
669 bp (Table 2.2) in P155 (Scenedesmus sp.), obtained from
chemically polluted river water, and in Q12 (Chlorella sp.), obtained
Chapter 2
75
from a backwater.
For accurate identification of the isolates, new algae-specific degenerate
primers were designed based on the complete 18S rDNA sequences of
widely different species of algae that belonged to a few representative
species of Chlorophyta, Charophyta, Ochrophyta, Cryptophyta,
Haptophyta, Rhodophyta, and Bacillariophyta. The newly designed
primers aligned with and amplified more or less complete 18S rDNA
sequences ranging from 1.5 kb to 2.8 kb depending on the algal strain.
The identity index was 88% for the forward primer
(GCCCGTAGTAAWWCTASAGCTAATAC) and did not amplify up
in any other environmental cultures except algae, whereas that for the
reverse primer (AAAAACGGGCGGTGTGTACAAAGGGC) was 92%;
however, that primer also detected another class of higher-order
organisms. We tested the specificity of the primers using different
samples of algae from the environment, which were PCR-amplified,
cloned, and sequenced. In none of the clones, DNA sequences from
other microbes from higher-order organisms highlighted; these primer
combinations were useful as molecular tools for specific amplification of
18S rDNA gene of algal species alone. The primers used to amplify the
rDNA region successfully amplified DNA from all the twenty two
microalgae isolates.
The PCR products fell into two size categories, namely 1.491–1.938 kb
and 2.4–2.8 kb. The amplified and cloned 18S rDNA sequences using
degenerate algae- specific primers were sequenced to obtain detailed
information on their close relatives through GenBank accessions number
(Table 2.2). Although all the sequence were analysed together, due to the
Chapter 2
76
differences in size, no common sequences were found for pair-wise
analysis and for multiple sequence alignments. The phylogenetic
positions of the 18S rDNA sequences in the size category 1.491–1.938
kb were analysed separately, along with other close relatives of algal
species obtained through NCBI-BLAST (Figure 2.8a, b). Percentage
similarity values obtained after pair-wise alignment of the rDNA
sequences of the isolates in that size category are listed in Table 2.2.
The relative phylogenetic positions based on the designed primers
sequence were matched with the results of morphological studies and
served to further confirm the identification of the isolates. Other unique
intronic positions were observed in the isolates of Scenedesmus sp.
collected from diverse habitats. These sequences were compared using
the software Bio-edit to define the novel pattern of intronic that
aligned with known Scenedesmus sp. reported earlier (Figure 2.8c).
Although the use of universal primers allows direct comparison between
studies, in analysing environmental samples, such use places some
limitations on the interpretation of results. These limitations can be
overcome by obtaining whole sequences from 18S rRNA genes as
reference trees. Since the selection of primers has a significant impact
on the results obtained from environmentally diverse ecosystems, such
reference trees could be used for studying large algal populations. The
present study thus provided a designed primer pair more suitable as
a molecular marker in studying eukaryotic diversity or in
characterizing microalgae. However, the overall morphological features
and 18S rDNA sequences confirm the taxonomical positions of these
isolates to a great extent.
77
Ch
apte
r 2
Tabl
e 2.
2 M
olec
ular
det
erm
inat
ion
of m
icro
alga
l iso
late
s.
Mic
roal
gal i
sola
te
Des
igne
d pr
imer
am
plic
on p
rodu
ct
(kb)
Clo
sest
rel
ativ
e an
d G
enB
ank
acce
ssio
n no
. U
nive
rsal
pr
imer
am
plic
on p
rodu
ct
(bp)
Phyl
ogen
etic
rel
ativ
e po
sitio
n us
ing
univ
ersa
l am
plic
on
sequ
ence
s
Chlo
rella
sp. (
Q12
) 1.
53
Chl
orel
la sp
. ZB
-201
4 K
J734
869
(99%
) 66
9 Au
xeno
chlo
rella
sp. C
CA
P 21
1/61
Para
chlo
rella
kes
sleri
(Y8)
1.
77
Para
chlo
rella
kes
sleri
stra
in S
AG
21
1 -11
c K
M02
0114
(99
%)
661
Chl
orel
la sp
. B
UM
1109
8
Chlo
rella
soro
kini
ana
(USA
) 1.
53
Chl
orel
la so
roki
nian
a st
rain
UK
M
2 K
P262
476
(99%
) 35
0 M
icra
ctin
ium
sp. T
P-20
08a
CC
AP
271/
1 Ch
lore
lla so
roki
nian
a (H
S)
1.49
C
hlor
ella
soro
kini
ana
stra
in S
AG
211-
31 K
F673
387
337
Chl
orel
la sp
. ZB
-201
4
Auxe
noch
lore
lla
prot
othe
coid
es (G
oa)
1.53
Au
xeno
chlo
rella
pro
toth
ecoi
des
stra
in S
AG
211
-10a
KM
0201
50
643
Auxe
noch
lore
lla s
p.
Scen
edes
mus
sp. (
V8)
2.
06
Scen
edes
mus
sp. K
GU
-D00
2
AB
7438
27 (9
1%)
401
Eust
igm
atos
vis
cher
i CC
AP
860/
7
Scen
edes
mus
sp. (
Mn2
5)
2.81
Sc
ened
esm
us sp
. KG
U-D
002
AB
7438
27 (9
2%)
NI
NI
Coel
astre
lla sp
. (Tn
1)
2.81
C
oela
stre
lla st
riol
ata
var.
mul
tistr
iata
CC
ALA
309
NI
NI
78
Ch
apte
r 2
Mic
roal
gal i
sola
te
Des
igne
d pr
imer
am
plic
on p
rodu
ct
(kb)
Clo
sest
rel
ativ
e an
d G
enB
ank
acce
ssio
n no
. U
nive
rsal
pr
imer
am
plic
on p
rodu
ct
(bp)
Phyl
ogen
etic
rel
ativ
e po
sitio
n us
ing
univ
ersa
l am
plic
on
sequ
ence
s Co
elas
trella
sp. (
V3)
2.
81
Coe
lastr
ella
stri
olat
a va
r.
mul
tistr
iata
stra
in C
CA
LA;
NI
NI
Coel
astre
lla sp
. (P1
9)
1.93
C
oela
stre
lla st
riol
ata
var.
mul
tistr
iata
stra
in C
CA
LA;
342
Coe
last
rella
stri
olat
a C
AU
P H
3602
Co
elas
trella
sp. (
Tre
e)
2.40
C
oela
stre
lla te
rres
tris
stra
in
CC
ALA
476
JX51
3882
(92%
)
NI
NI
Scen
edes
mus
sp. (
Mn9
) 2.
81
Scen
edes
mus
sp. N
J-1
JX28
6515
(93%
)
353
Chl
orel
la e
mer
soni
i CC
AP
211/
8P
Scen
edes
mus
sp. (
P155
) 2.
4 Sc
ened
esm
us sp
. KG
U-D
002
AB
7438
27 (9
2%)
341
Chl
orel
la e
mer
soni
i CC
AP
211/
8P
Scen
edes
mus
sp. (
P58)
2.
81
Scen
edes
mus
sp. K
GU
-D00
2
AB
7438
27 (9
2%)
339
Coe
last
rella
sp. S
AG
247
1
Scen
edes
mus
sp. (
P115
) 2.
63
Scen
edes
mus
sp. N
J-1
JX28
6515
(93%
)
343
Coe
last
rella
stri
olat
a C
AU
P H
3602
Sc
ened
esm
us sp
. (V
11)
2.63
Sc
ened
esm
us sp
. KG
U-D
002
AB
7438
27 (9
2%)
658
Visc
heria
hel
vetic
a C
CA
LA 5
14
Scen
edes
mus
sp. (
V9)
2.
81
Scen
edes
mus
sp. K
GU
-D00
2
AB
7438
27 (9
2%)
656
Pote
riooc
hrom
onas
mal
ham
ensi
s D
S
79
Ch
apte
r 2
Mic
roal
gal i
sola
te
Des
igne
d pr
imer
am
plic
on p
rodu
ct
(kb)
Clo
sest
rel
ativ
e an
d G
enB
ank
acce
ssio
n no
. U
nive
rsal
pr
imer
am
plic
on p
rodu
ct
(bp)
Phyl
ogen
etic
rel
ativ
e po
sitio
n us
ing
univ
ersa
l am
plic
on
sequ
ence
s Sc
ened
esm
us sp
. (P1
52)
2.63
Sc
ened
esm
us sp
. KG
U-D
002
AB
7438
27 (8
9%)
343
Chl
orel
la e
mer
soni
i CC
AP
211/
8P
Scen
edes
mus
biju
gus
(Lad
akh)
1.55
Sc
ened
esm
us b
ijugu
s var
.
obtu
sius
culu
s vou
cher
IZTA
:
NI
NI
Scen
edes
mus
sp.
(V15
) 2.
81
Scen
edes
mac
eae
sp. T
ow 9
/21
P-
14w
AY
1976
39 (9
3%)
667
Scen
edes
mac
eae
sp. T
ow 9
/21
P-
14w
Co
elas
trella
sp.
(P63
) 2.
40
Coe
last
rella
sp. B
UM
1111
3;
KC
2184
97 (9
5%)
NI
NI
Visc
heria
stel
lata
(P11
) 1.
51
Visc
heria
stel
lata
stra
in S
AG
33.8
3 K
F848
919
(98%
)
338
Eust
igm
atos
vis
cher
i CC
AP
860/
7
NI:
not i
dent
ified
80
Chapter 2
Chapter 2
Coelastrella sp. (TN1)
Coelastrella sp. SAG 2471 |KM020087.1|
Coelastrella terrestris CCALA 476 |JX513882.1|
Coelastrella striolata var. multistriata CCALA 309 |JX513880.1|
Coelastrella sp. (V3 )
Coelastrella sp.(Tree)
Coelastrella multistriata var. corcontica |AB037082.1|
Coelastrella sp. (P63)
Coelastrella sp. (P19)
Schroederiella apiculata |AB037098.1|
Scenedesmus bijugus (Ladakh)
Scenedesmus bijugus var. obtusiusculus voucher IZTA:1830 |KJ808696.1|
Acutodesmus obliquus GB1g |AB917101.1|
Scenedesmus basiliensis strain ACKU 646-06 |KF898121.1|
Scenedesmus obliquus CCAP 276/7 |FR865737.1|
Schroederia setigera |AF277650.1|
Auxenochlorella protothecoides NIES-2164 |AB488569.1|
Auxenochlorella protothecoides SAG 211-7a |X56101.1|
Auxenochlorella protothecoides CCAP 211/8D |FR865686.1|
Auxenochlorella protothecoides SAG 211-10a |KM020150.1|
Auxenochlorella protothecoides NIES-2165|AB488570.1|
Auxenochlorella protothecoides (Goa)
Auxenochlorella sp. CCAP 211/61 |FN298932.1|
Poterioochromonas stipitata |AF123295.1|
Vischeria stellata SAG 33.83 |KF848919.1|
Vischeria Stellata (P11)
Vischeria helvetica KGU-Y001|AB731568.1|
Eustigmatos sp. MT-2012 |JX188078.1|
Eustigmatos vischeri UTEX 310 |FJ858973.1|
Chlorella sorokiniana LB65 |KJ616755.1|
Chlorella sorokiniana YeoJu4 |KF864471.1|
Chlorella sorokiniana strain Icheon4 |KF864476.1|
Chlorella sorokiniana (USA)
Chlorella sorokiniana (HS)
Chlorella sp. ZJU0208 |JX097060.1|
Chlorella sp. ZJU0205 |JX097055.1|
Chlorella sp. (Q12)
Chlorella chlorelloides CB 2008/110 |HQ111432.1|
Parachlorella kessleri (Y8)
Parachlorella kessleri SAG 211-11c |KM020114.1|
Parachlorella kessleri CCAP 211/11H |FR865655.1|
Parachlorella kessleri BAFC CA10|HM744739.2|
Parachlorella kessleri SAG 27.87 |KM020113.1|
Chlamydomonas reinhardtii CC-621 |JX888472.1|
Chlamydomonas reinhardtii CCAP 11/32CW15 |FR865576.1|
Rhodospirillum rubrum strain BRP1 |FJ853177.1|
2
99
98
87
87
5152
100
2
79
95
32
42
39
43
4
64
74
15
8
49
13
91
80
96
88
0.1
Chapter 2
Figure 2.8 Phylogenetic analysis of algal isolates by the Maximum
Likelihood method based on the Tamura-Nei model.
(a) The value at each branching point shows the ML (as a percentage,
based on 1000 replicates) that a given isolate belongs to the cluster in
question. GenBank accession no. of the sequences from earlier studies is
given, where available, after the isolate. All positions containing gaps
and missing data were eliminated. Evolutionary analyses were conducted
in MEGA6
(b) Phylogenetic analysis of Scenedesmus sp. isolates by the Maximum
Likelihood method based on the Tamura-Nei model, (c) 18S rDNA
sequence alignment of Scenedesmus sp. using Bio-edit. 18S rDNA
sequences from Scenedesmus sp. isolates were compared with those from
other species of Scenedesmus reported earlier and gathered from NCBI.
Broken lines indicate the intron/gap regions.
Scenedesmus sp. (P115)
Scenedesmus sp. (MN25)
Scenedesmus sp. (P155)
Scenedesmus sp. ISTGA1 |KC342184.1|
Scenedesmus sp. (V11)
Scenedesmus sp. (P58)
Scenedesmus sp. (V8)
Scenedesmus sp. (V9)
Scenedesmus sp. (MN9)
Scenedesmus sp. (P152)
Scenedesmus sp. KGU-D002 |AB743827.1|
Scenedesmus sp. CCMP 1625 |AF339735.1|
Scenedesmus sp. NJ-1 |JX286515.1|
Scenedesmus sp. II22 |FJ946899.1|
Scenedesmaceae sp. Tow 9/21 P-14w |AY197639.1|
Scenedesmus sp. (V15)
Chlorella sp. ZJU0205 |JX097055.1|
76100
100
100
100
96
51
99
99
98
76
69
42
46
0.01
Chapter 2
2.4 Conclusion Microalgae strains can be found in a range of extreme environments,
including in polluted water bodies and in salty lakes. In the present
study twenty isolates, selected from 200 collected, were isolated and
identified using designed primers (1.5-2.0 kb). Simple strategies to
isolates the unialgal colonies and to subsequent purify the diverse
isolates, free from any contamination of other microorganisms were used.
In addition to genetic identification, the morphometric characteristics of
these twenty isolates were observed under SEM, and these
characteristics were compared with related isolated previously reported
in the literature. To preserve the diverse isolates for further study
methodology were standardized for preservation of these green
microalgae by immobilising in alginate beads. The purification,
identification and preservation methods described in this chapter were
successful for obtaining and storing a range of diverse green microalgae.
Chapter 3
84
Chapter 3: Characterising new oil and carotenoid producing
microalgal isolates with biofuel potential.
3.1 Introduction
Photosynthetic microalgae have significant potential as energy producers
and are also promising sources of metabolites of importance in industrial
applications (Spolaore et al., 2006). Microalgal diversity constitutes
200,000 species of which only a small percentage have been studied
in detail (De Clerck et al., 2013). Such an untapped resource has
potential to produce chemicals for an array of sector and industries
(Kolympiris et al., 2014). Bioprospecting of microalgae has been
carried out from ecologically diverse habitats such as the deep seas
(Boeuf and Kornprobst, 2009) and polluted waters (Sterrenburg et al.,
2007). Such diverse habitats may harbour distinct isolates with unique
propertiesand multiple applications. During the past fewyears, microalgae
have been extensively explored for biofuel and for bioactive
compounds. In recent years, unique strains have been isolated from a
range of habitats in the tropics including both aquatic (lakes, streams,
and backwaters) and terrestrial habitats. Such extremes as very high
temperatures and prolonged exposure to intense light found in the tropics
have conferred on the extremophiles some distinctive physiological
properties (Gouveia et al., 2009). Since the early 1970s, bioprospecting
of microalgae (such as green algae, cyanobacteria, and diatoms) has been
attempted for various uses, and their ability to consume carbon
dioxide during photosynthesis and to act as a sustainable source of
biofuels has led to increasing attention in recent years (Jones and
Chapter 3
85
Mayfield, 2012). Both lipid and biomass content are equally important
for achieving high lipid productivity (Yen et al., 2013). Microalgal lipids
are divided in to two main categories, namely storage lipids (non-polar)
and structural lipids (polar) (Sharma et al., 2012). Because they are rich in
essential nutrients that are not synthesized by higher eukaryotic
organisms. Deriving multiple products such as lipids and high-value
products from the same biomass in one growth cycle can help make
the process economically feasible (Nobre et al., 2013). Biologically active
compounds derived from microalgae have attracted increasing industrial
attention (De Morais et al., 2015). The diverse gene pool of microalgae
still remains to be fully explored for the production of bioactive
compounds, particularly using a biorefinery approach.
Carotenoids are an important class of bioactive compounds produced by
microalgae, with large existing markets for the two carotenoids β-
carotene and astaxanthin. The market for other carotenoids, such as
lutein, are still smaller but developing. Lutein has multiple applications
and have been shown to help prevent age-related macular degeneration
and improve cardiovascular function. Lutein is also used as a food dye
and additive in aquaculture for the pigmentation of salmonids,
ornamental fish, and in the poultry industry (Lorenz and Cysewski,
2000). The market for these bioactive compounds continues to expand.
Thus, new microalgal species that can produce these or other useful
compounds is a continuing area of research. It is against this background
that the present study sought to compare twenty two microalgal isolates
for identifying the strains most suitable for biofuel and carotenoids
production.
Chapter 3
86
The selection criteria were the ability to produce (1) lipids with fatty
acid methyl ester (FAME) profiles suitable for biofuels and (2)
carotenoids which would potentially offset the cost of biofuel
production. Selected strains were subjected to lipid localisation studies
using fluorescent dyes. Fourier transform infrared spectroscopy (FTIR)
was also used to study the accumulation of total lipids within the
microalgal cells.
3.2 Materials and Methods 3.2.1 Chemicals used All chemicals, including fatty acid methyl ester standards (C19:0) used in
this study were of analytical grade and procured from Sigma-Aldrich
(St. Louis, Missouri, USA) and Merck Chemicals (Frankfurter Strabe,
Darmstadt, Germany). For carotenoid extraction and HPLC analysis
for quantification, the solvents acetone, hexane (Fischer scientific,
Waltham, MA, USA), methanol, acetonitrile and ethyl acetate (HPLC
grade from Fischer scientific, Waltham, MA, USA) were used. The
carotenoid standards astaxanthin and zeaxanthin were from obtained
from Cayman Chemical Co., whereas canthaxanthin, β-carotene and
lutein were obtained from Sigma Aldrich (St. Louis, Missouri, USA).
For lipid localisation studies, BODIPY 505/515 was obtained from
molecular probes (4,4-difluoro-1,3,5,7-tetramethyl-4- bora-3a,4a-diaza-s-
indacene; Invitrogen Molecular Probes, Carlsbad, CA, USA).
Chapter 3
87
3.2.2 Culturing of microalgal isolates The twenty two isolates were grown in 250 mL Erlenmeyer flasks
containing 100 mL BBM medium and incubated for nine days at 25
°C under 16 h of fluorescent white light (120 μE/m2/s) alternating with
8 h of darkness and with constant shaking (150 rpm). Cell suspensions
of equal strength of the isolates were also grown in experimental tubes
containing 70 mL of the medium in Multi-Cultivator (MC) 1000-OD
(Photon Systems Instruments, Drasov, Czech Republic). To ensure that
cell counts for the comparative growth experiments the same initially,
the number of cells was counted using a Neubauer haemocytometer
(Rohem Instruments, Nashik, Maharashtra, India). The initial optical
density (OD) of the cell suspension with a concentration of 3 × 106
cells/mL was recorded. All the tested isolates were maintained at an
illumination intensity of 120 μE/m2/s as a standard growth condition. The
source of illumination was placed 2.0 cm away from the external wall
of the vessel, the internal diameter of which was 15 mm at 25 °C.
Cultures were aerated with plain air at rate of 0.12 mL/min
throughout the experiment.
3.2.3 Screening of microalgal isolates All the cultures with an initial cell count of 3×106 cells /mL were
cultivated under photoautotrophic conditions in a MC 1000-OD at 25
°C under 16 h of light (120 μE/m2/s) alternating with 8 h of darkness.
The aeration rate was maintained at 0.12 mL/min. Microalgal growth was
monitored by measuring the daily changes in OD at 680 nm with an
OD viewer software attached to the cultivator.
Chapter 3
88
For studying growth kinetics, all isolates were cultivated in BBM media.
The growth rate of microalgal isolates was estimated by fitting the OD at
the exponential phase of each isolates to an early stage exponential
growth phase in Eq. (1) (Wang et al., 2010)
(1)
Where OD0 is the initial OD and ODt is the optical density
measured on day t.
The microalgal cells were cultured for about 9 days on average and then
cells were harvested by centrifuging (10 min at 6000 rpm at 4 °C)
followed by lyophilisation. The twenty two characterized isolates were
further evaluated for their efficiency in producing biomass (dry biomass
produced, expressed in milligrams per litre per day) and lipids (expressed
as a percentage of total biomass on dry weight basis). The pellets were
washed twice with sterile deionized water, lyophilized for 48 h, and
their dry weight was determined gravimetrically. Carotenoids (lutein
and astaxanthin) production, expressed in milligrams per litre per day,
was estimated using the following equations:
Lp = Bp × Lc and Ap= Bp × Ac (2) where Lp is lutein productivity, Ap is astaxanthin productivity, Bp
is biomass productivity, and Lc and Ac are lutein and astaxanthin contents,
respectively.
Chapter 3
89
3.2.4 Qualitative analysis of lipids using BODIPY 505/515 staining Cells of the twenty two isolates from 10-day-old cultures on average
were harvested by centrifuging for 10 min at 6000 rpm at 4 °C. A
lipophilic dye, BODIPY 505/515 (4,4-difluoro-1,3,5,7-tetramethyl-4-
bora-3a,4a-diaza-s-indacene (Invitrogen Molecular Probes, Carlsbad,
California), was dissolved in dimethyl sulfoxide (DMSO) at a stock
concentration of 1 mg/mL. The stock solution was stored in the dark at –
20 °C and was diluted to achieve a range of concentrations (1, 2, 3, and
5 μg/mL) in 1% DMSO. The cells were prewashed with Milli-Q
water, fixed with 3.7% paraformaldehyde, stained using each of the
above concentrations separately, and incubated for 20 min in the dark.
The concentration of 2 μg/mL proved the most suitable for staining
intracellular lipids. The fixed samples were then mounted on clean
glass slides, covered with coated coverslips, and examined using
confocal microscopy (an apochromat ×63 objective with oil
immersion). The signal from BODIPY was captured using a 488 argon
laser excitation line. For confocal microscopy, a Carl Zeiss microscope
(model LSM 710), running the image capture software LAS AF ver.
2.6.3.8137, was used. For BODIPY, the excitation came from a 458 nm
argon laser and the emission was between 493 nm and 525 nm.
Chapter 3
90
3.2.5 Spectral Monograms and Biomass composition using Attenuated
total reflectance (ATR-FTIR)
ATR-FTIR spectrum of dried microalgal biomass was recorded at room
temperature using a Nicolet 6700 (Thermo scientific, Waltham, MA,
USA). An FTIR instrument using a diamond iTR reflectance with
DTGA detector was used. The dried algal samples of all twenty two
isolates were pressed against a diamond cell prior to scanning.
Background was collected before each samples spectrum. A total of 62
scans were made for each sample spectrum. A region of 4000–400 cm-1
were used for scanning.
Data were exported using Omnic 8.0.342 Software (Thermo Scientific)
and processing including baseline correction and vector normalization
was performed using the OPUS 6.0 software suite (Bruker Optik
GmbH, Ettlingen, Germany) or unscramble X software.
3.2.6 Lipid extraction and preparation of FAMEs Total lipids were extracted from the freeze-dried biomass following
the method developed by Lewis et al. (2000) with some modifications: a
known quantity (10 mg) of dried cell biomass was extracted in
chloroform : methanol (2 : 1, v/v); A suspension was homogenized
using vortex (Spinix, Maharashtra, India) for 2 min followed by
centrifuging for 15 min at 10 000g. The extraction was repeated three
times until the biomass became colourless. A method described by
Christie and Han. (2010) was employed to study FAME profiles. For
preparation of FAME, 500 μL of toluene was added into dried
lipid samples
Chapter 3
91
followed by 10 μL of internal standard (50 mg of C19:0- Methyl
nonadecanoate), 400μL of freshly prepared hydrogen chloride in
methanol (prepared by adding 1 mL acetyl chloride drop wise to 10
mL of methanol on ice), 200μl of antioxidant butylated hydroxyl toluene
(BHT) and incubated at 50 °C overnight. Following day, 1mL of 5%
NaCl and 1mL of hexane was added to dried samples. Finally, the
solvent layer containing hexane were analysed using Gas
chromatography.
3.2.7 Gas chromatography The samples of FAMEs were analysed by gas chromatography (Agilent
122-2332 column, Santa Clara, California, USA) with mass
spectrometry (MS). Each sample (2 μL) was injected into a capillary
column (DB-23; 30 mm × 0.25 mm; film thickness, 0.25 μm). The
initial oven temperature of 70 °C was gradually increased by 10° C/min
until it reached 180 °C, which was maintained for 14 min. The oven
temperature was raised again at 4 °C/min to reach 220 °C, which was
maintained for 36 min. The detector temperature was set at 250 °C.
The injector was maintained at 250 °C and a sample volume of 1 μL
was injected. The carrier gas, namely helium, was released at 2.0
mL/min. A hexane injection was used as a blank. The sample injections
used C19:0 (Methyl nonadecanoate) as an internal standard to verify
the alignment of retention time with the calibration injection. The run
time for a single sample was 36 min. Fatty acids were quantified by
using the internal standard and identified by comparing the retention
time with that of a known standard mixture of FAs (Supelco FAME mix
Chapter 3
92
C4-C24 (Bellefonte, Pennsylvania, USA). Fatty acid peaks were further
analysed and integrated using a chemstation chromatography software
(Agilent Technologies, Germany).
3.2.8 Evaluation of microalgal oil properties for biofuel Some properties of the microalgal isolates were evaluated to screen them
for biofuel production. A high cetane number (CN) of a FA denotes
better combustion quality whereas the iodine value (IV) denotes the
degree of unsaturation of the microalgal oil. A high IV indicates low
oxidative stability of the fuel (Knothe, 2012). Hence, in the present study,
CN, saponification value (SV), and IV were used in evaluating the
FAME profiles using the following empirical Eq. (3–5) proposed by
Mandotra et al. (2014)
93
Chapter 3
(3)
(4) SV = (5) Where, F is the percentage of each FA, D is the number of double
bonds, and MW is the molecular weight of the FA. The standard biofuel
properties as laid down in ASTM D6751 and EN 14214 were considered
for evaluating the data.
3.2.9 Determination of microalgal carotenoids Total carotenoids were obtained using the method described by Garcia-
Plazaola et al. (2012). In general, 25 mg of freeze-dried biomass was
subjected to sonication at 40 kHz for 5 min at room temperature
(Branson, Danbury, Connecticut, USA) to break the cell walls, and
carotenoids and other compounds were extracted in acetone by
continuous agitation for 3 min. The mixture of microalgal cells was
then centrifuged at 4000 rpm for 15 min. The supernatant contained
pigments, and the extraction process was repeated two more times so that
the supernatant was colourless. All the extractions were carried out under
darkness or under dim light to avoid photo-oxidation of the carotenoids.
In the present study, we maintained all the isolates under uniform
conditions to enable an accurate comparison of lipids and carotenoids
productivities (expressed in milligrams per litre per day). The
carotenoids were analysed by injecting 10 μL of the carotenoid into an
HPLC system (Shimazdu Co., Japan) equipped with a photodiode array
94
Chapter 3
detector. The pigments were separated using a reverse-phase C18 column
coupled with a security guard column (Lune 3.0 mm C18 250 mm × 4.6
mm, 5 μm, Phenomenex, Torrance, California, USA). The binary mobile
phase consisted of solvent A (acetonitrile: methanol, 7: 1 v/v) and solvent
B (acetonitrile: methanol: water: ethyl acetate, 7 : 0.96 : 0.04 : 2 v/v),
and the following gradient method was used to separate the pigments,
as recommended by Garcia-Plazaola et al. (2012). The flow rate was
set at 1.0 mL/min, and pigment content was detected by measuring the
absorbance at 450 nm. The chromatographic peaks were analysed by
comparing the retention time against the standard. The standards for
lutein and canthaxanthin were bought from Sigma Chemical Co. (St.
Louis, Missouri, USA); for astaxanthin and zeaxanthin, from Cayman
Chemical Co.; and HPLC- grade acetonitrile, methanol, and ethyl
acetate, from Fisher Scientific, USA.
3.3 Results and Discussion 3.3.1 Lipid quantification using BODIPY 505/515
The conventional method of lipid quantification is labour intensive and
therefore lesscost effective. In recent decades, new and emerging
techniques based on fluorescence microscopy have shown promise in
the quantification of lipids (Mutanda et al., 2011). In particular, the fast
and reliable lipophilic fluorescence dye BODIPY 505/515 has proved
effective for rapid screening of microalgal isolates and localization of
lipid bodies using laser scanning confocal microscopy. In the present
study, all the isolates could be stained satisfactorily using this dye,
and the lipid globules showed up as bright green structures.
95
Chapter 3
The distribution of the green lipid globules was observed in all
cultures. However, the storage pattern of the lipid globules varied
among the isolates. In Chlorella sp., two or more large lipid globules were
found in C. sorokiniana (USA) (Figure 3.1 a), whereas Chlorella sp.
(Q12) always showed three small globules (Figure 3.1 b). On the other
hand, the globules in Parachlorella kesselri were not spherical (Figure
3.1 c). However, in two isolates of C. sorokiniana (USA and HS), lipids
appeared to accumulate by the merger of globules to form a new lipid
body within an existing lipid body in the cytoplasm.
Wong and Franz. (2013) reported that diverse isolates showing an identical
pattern of lipid storage might provide a new understanding of lipid
biosynthesis. However, in Coelastrella sp. (V3) and Coelastrella sp.
(Tree), a number of small, dispersed lipid droplets were seen throughout
the cell (Figure 3.1d and 3.1e). Moreover, the same Coelastrella sp.
(Tn1) from different habitats showed different patterns of lipid
distribution (Figure 3.1 f) and distinctly separate phylogeny from other
Coelastrella spp. On the other hand, clades of Scenedesmus sp. from
diverse habitats have shown different patterns of lipid localization within
a single clade. One of the Scenedesmus spp. (P155) (Figure 3.1g)
isolated from chemically polluted river water and Scenedesmus sp.
(Mn25) from a backwater (Figure 3.1h) showed large lipid bodies
merging with a fixed number of lipid bodies and forming new lipid
bodies within existing lipid bodies, whereas Scenedesmus sp. (P58 and
V11) from two different habitats (Figure 3.1 i and 3.1 j) displayed the
presence of lipid bodies on the cell membrane as well as distributed
96
Chapter 3
throughout the cell. These observations serve to confirm the
identification of species based on molecular data. However, in
Scenedesmus sp. (P152, V8, and Mn9), isolates that share a common
phylogeny may belong to distinct habitats, as shown by the varying
size of their lipid globules (Figure 3.1k, 3.1l, and 3.1m). Scenedesmus
sp. from a freshwater habitat (V15) showed a different morphology,
with spherical lipid globules widely scattered all over the cells (Figure
3.1n). This shows that the same isolates from a similar habitat may differ
in th pattern of lipid accumulation. We believe that variation in the
structure and composition of the cell wall results in two different
fluorescence patterns. In Scenedesmus bijugus (Ladakh), lipid globules
were observed on the cell membrane as well as in the cell matrix
(Figure3.1o). In Vischeria stellata (P11), which belonged to
Eustigmatophyceae, the lipid bodies were widely distributed within the
cells (Figure 3.1p). Thus the diverse isolates were equally diverse in their
pattern of lipid storage and distribution.
97
Chapter 3
Figure 3.1(a)-(p) Representative confocal laser scanning micrographs of
diverse microalgal isolates using BODIPY 505/515 under tested growth
conditions: a) Chlorella sorokiniana (USA), (b) Chlorella sp.(Q12), (c)
Parachlorella kessleri (Y8), (d) Coelastrella sp.(V 3),(e) Coelastrella
sp.(Tree), (f) Coelastrella sp. (Tn1), (g) Scenedesmus sp. (P155), (h)
Scenedesmus sp. (Mn25), (i) Scenedesmus sp.(P58), (j) Scenedesmus sp.
(V11 ), (k) Scenedesmus sp. (P152), (l) Scenedesmus sp.(V8), (m)
Scenedesmus sp.(Mn9), (n) Scenedesmus sp.(V15), and (o) Scenedesmus
bijugus (Ladakh). Scale bar =5μm.
3.3.2 ATR-FTIR analysis The ATR-FTIR spectra collected for all 22 isolates showed different peak
intensities under the same cultivation condition, showing variation in
biochemical composition, including lipid content. The obtained spectral
signature indicates different functional groups present within diverse
(a)
b
g g (b) (c) (d) (e)
(f) (g) (h) (i) (j)
(k) (l) (m) (n) (o)
98
Chapter 3
isolates with major change in the protein, lipid, phosphate and
polysaccharides content. Two of the main regions used for determining
total lipid content are the methylene region at 2800-3000cm-1 and the
ester bond stretches at 1740 cm-1 (Laurens and Wolfrum,2011). The ATR-
FTIR spectra from all 22 isolates are shown in Figure 3.2 (a)-(d). Major
variations were seen in the spectra range between 3000-2800 cm-1 and
1700–850 cm-1. Strong absorption bands at 1150-1000, 1530, 1650,
1745, 2855 and 3300 cm-1 were present in all twenty isolates
due to C–O–C stretching from polysaccharides, C=O peaks are the
Amide I and II from protein (Giordano et al., 2001), and C=O from
cellular lipid and fatty acid esters functionalities (Movasaghi et al.,
2008). The band at 2855 cm-1 was assigned to C-H stretching from
methylene lipid acyl chains, whereas the last band denotes string O-H
stretching arising from water molecules inside the cells. Smaller bands
at ~800-550 cm-1 are due to aromatic C-H bending. In order to analyse
and compare difference spectra of different isolates grown under the same
conditions, we performed secondary derivatization of ATR-FTIR spectra,
since this resolves the intersecting IR spectra bands (Susi and Byler, 1986).
We analysed secondary derivative spectra of cells at an average of 10
days. In Figure 3.2 (a)-(d) the spectra from all twenty two isolates are
dominated by lipid acyl groups (methyl and methylene groups) in the
range 3045-2796 cm-1. As shown in Figure 3.2 a1 and a2 the spectrum
is mainly characterised by four bands 2969 (νasym CH3), 2963 (ν antisym
CH2), and 2846 cm-1 (ν sym CH2). These spectral signatures were found
to vary among different isolates. Scenedesmus sp. (V11 and V8) in
freshwater habitats showed higher accumulation of methylene acyl lipid.
99
Chapter 3
A peak at 1745 cm-1 for lipid esters was lower than one for acyl lipids.
However the intensity of lipid esters varied among the isolates. Isolates
obtained from polluted water habitats which regularly received washing
deteregent (Figure 3.2 b1 and b2), Parachlorella kessleri and Coelastrella
sp., showed high oil accumulation, with relatively intense bands for
asymmetric stretching vibrations of CH2 of acyl chains (lipids),
symmetric (C-H) from methylene CH2 groups of lipids and C-O
stretching of total lipids. Isolates from backwater habitats (Figure 3.2 c1
and c2), Coelastrella sp. (Tn1) and Scenedesmus sp. (Mn25), showed
higher intensity of acyl lipid and C-) stretching of total lipids
compared to other isolates. The terrestrial isolates (Figure 3.2 d1 and
d2), Coelastrella sp. (Tree), showed accumulation of symmetric and
asymmetric acyl lipids to a greater extend.
Chap
ter 3
100
a1
Chap
ter 3
101
a2
Chap
ter 3
Cha
104
b1
Wav
enum
ber c
m-1
b2
Wav
enum
ber c
m-1
Chap
ter 3
c1
Wav
enum
ber c
m-1
c2
Wav
enum
ber c
m-1
Chap
ter 3
d1
Wav
enum
ber c
m-1
Chap
ter 3
Figu
re 3
.2 (
a)-(
d).
FTIR
mic
rosp
ectro
scop
ic s
pect
ra o
f di
ffer
ent
mic
roal
gal
spec
ies
from
div
erse
hab
itats
(a1
) A
TR-F
TIR
Fres
hwat
er
isol
ates
spe
ctra
(a2
) Se
cond
ary
deriv
ativ
e an
alys
is o
f fr
eshw
ater
isol
ates
inta
ct c
ells
(b1
) A
TR-F
TIR
pol
lute
d
wat
er is
olat
es s
pect
ra (
b2)
Seco
ndar
y de
rivat
ive
anal
ysis
of
Pollu
ted
river
wat
er i
sola
tes
inta
ct c
ells
(c1
) A
TR-F
TIR
Bac
kwat
er w
ater
iso
late
s sp
ectra
(c2
) Se
cond
ary
der
ivat
ive
anal
ysis
of
bac
kwat
er i
sola
tes
inta
ct c
ells
(d1
) A
TR-F
TIR
Bac
kwat
er a
nd f
resh
wat
er t
erre
stria
l is
olat
es s
pect
ra (
d2)
Se
cond
ary
deriv
ativ
e
anal
ysis
of
bac
kwat
er
and
fre
shw
ater
terr
estri
al i
sola
tes
inta
c c
ell
Wav
enum
ber c
m-1
d2
Chapter 3
108
3.3.3 Comparative analysis of photo-autotrophic growth, biomass and
lipid levels
Rapid growth is an important prerequisite to obtaining a high cell density
and intracellular lipids, under optimal culture conditions (Talebi et al.,
2013). Under identical conditions, the isolates varied a great deal in their
ability to produce biomass: the productivity ranged from 83.3 ± 8.5 mg
L–1 d–1 to 174.7 ± 6.7 mg L–1 d–1 (Figure 3.3). Microalgal growth as
assessed by OD in the isolates drawn from four different habitats,
namely freshwater, chemically polluted river water, backwater, and land,
is shown in (Fig. 3.4 a-d). The average specific growth ra t e was
estimated at the exponential growth phase (Table 3.1). The growth
varied among the isolates in the same medium (BBM) under similar
culture conditions. In the present study, lipid productivity and biomass
productivity – expressed as daily production, in milligrams, of biomass
or lipids produced per litre of the medium – were considered equally
important in selecting appropriate strains for biofuel production. Biomass
productivity is considered important in determining lipid productivity
for biodiesel production (Griffiths and Harrison, 2009).
Under experimental conditions, total lipid content of freshwater isolates
was highly variable (13.2 ± 1.8 - 36.5 ± 1.81 % of the dry weight)
whereas that of backwater isolates peaked at 36.5 ± 2.1 % and that of
land isolates was average 15.83 ± 1.65 (Table 3.1). Although it can be
difficult to translate the results of lab scale strain comparisons to large
scale production due to variability, these lab scale trials are a good
indication of ease of strain growth and general productivity.
Chapter 3
109
Reported average lipid content of green alga varies from 13% to
31% of dry weight (Griffiths and Harrison, 2009). Lipid productivity
varied between 10.0 ± 1.9mg L–1 d–1 to 43.8 ± 5.1mg L–1 d–1. Of all the
Scenedesmus isolates studied, one from chemically polluted river water
(P58) and one from freshwater (Ladakh), showed higher productivity
43.8 ± 5.1mg L–1 d–1 and 40.1 ± 3.3mg L–1 d–1 for lipids and 155.7 ± 14.3
mg L–1 d–1 and 174.7 ± 6.7mg L–1 d–1 for biomass, respectively (Table
3.1) – than that reported elsewhere, such as (1) Scenedesmus sp.
(Tripathi et al., 2015), and Scenedesmus SDEC-8 (Song et al., 2014)
under non-aerated conditions; and (2) less than that reported for
Scenedesmus sp. DM (Rodolfi et al., 2009). Chlorella sp. (Q12) showed
higher productivity (39.7 ± 3.8 mg L–1 d–1 for lipids and 169.8 ± 5.2 mg
L–1 d–1 for biomass) among previously reported Chlorella sp., including
one reported by (Cheirslip and Torpee, 2012), Chlorella sp. SDEC-6
(Song et al., 2014) and Chlorella sp. (Jena et al., 2012) as shown in
Table 3.2. The isolate from high altitudes, namely S. bijugus (Ladakh),
also produced more biomass (174.7 ± 6.7mg L–1 d–1) and lipids (40.1 ±
3.3mg L–1 d–1) than the figures reported for S. bijuga grown on
wastewater effluent diluted with BG11 medium (Shin et al., 2015). The
other group of isolates including Coelastrella showed high biomass and
lipid productivity. Coelastrella sp. (P63) from chemically polluted river
water produced 152.9 ± 13.3mg L–1 d–1 of biomass and 43.5 ± 6.2mg
L–1 d–1 of lipids—figures that were comparable to those reported
by others, such as Coelastrella M 60 (Karpagam et al., 2015) and
Coelastrella F 50 (Hu et al., 2013) (Table 3.2).
Chapter 3
110
The above results indicate that microalgal isolates from highly chemically
polluted as well as backwater habitats indicate the potential of these strains
to produce useful levels of lipids and biomass. Scenedesmus sp. (P58), S.
bijugus (from Ladakh), Coelastrella sp. (P63), and Chlorella sorokiniana
(USA) were the most promising lipid and biomass producers among all
the twenty two isolates (marked by an asterisk Figure. 3.3)
Figure 3.3 Biomass (mg/L/d) and lipid productivity (mg/L/d) of
twenty two microalgal isolates in 9 days old culture.
180 160 140 120 100 80 60 40 20
00
Biomass productivity Lipid productivity
60
50
40
30
20
10 00
Microalgal isolates
Chapter 3
111
Figure 3.4 Growth pattern of different microalgal species from diverse
habitats cultivated in Multicultivator MC OD-1000 for 10 days at 25 °C
under 16 h of light (120 μE/m2/s) alternating with 8 h of darkness. (a)
Freshwater isolates (b) Isolates from polluted river water (c) Isolates
from backwater (d) Isolates from terrestrial sites.
Chap
ter 3
112
Ta
ble
3.1
Biom
ass
prod
uctiv
ities
, lip
id c
onte
nt, l
ipid
pro
duct
iviti
es, l
utei
n co
nten
t, as
taxa
nthi
n an
d lu
tein
pro
duct
ivity
of
mic
roal
gal
isol
ates
Isol
ates
Sp
ecifi
c gr
owth
ra
te (
OD
,mμ)
B
iom
ass
prod
uctiv
ity
(mg
L-1d-1
) Li
pid
prod
uctiv
ity
(mg
L-1d-1
)
Lipi
d
cont
ent (
% d
ry w
t.)
Asta
xant
hin
prod
uctiv
ity
(mg
L-1d-1
)
Lute
in
cont
ent
(mg/
g)
Lute
in
prod
uctiv
ity
(mg
L-1d-1
)
Chl
orel
la sp
. (Q
12)
0.40
16
9.81
± 5
.22
39.7
2 ±
3.89
24
.97
± 2.
69
0.18
2.
26
0.36
Para
chlo
rella
kes
sler
i (Y
8)
0.39
15
9.49
± 7
.63
31.7
1 ±
4.34
21
.42
± 3.
19
0.07
0.
28
0.04
Chl
orel
la so
roki
nian
a (U
SA)
0.32
13
4.94
± 1
2.52
31
.94
± 1.
26
28.5
8 ±
1.33
n.
d.
0.68
0.
07
Chl
orel
la so
roki
nian
a (H
S)
0.27
10
2.26
± 9
.65
11.5
1 ±
2.61
13
.55
± 2.
77
0.06
5.
90
0.5
Auxe
noch
lore
lla
prot
othe
coid
es (G
oa)
0.53
16
7.11
± 1
2.22
31
.33
± 3.
64
24.9
7 ±
2.69
0.
06
0.76
0.
11
Scen
edes
mus
sp.
(Mn2
5)
0.30
10
3.23
± 8
.38
34.5
7 ±
2.64
36
.53
± 2.
13
n. d
. 0.
0 0.
00
Coe
lastr
ella
sp. (
Tn1)
0.
36
124.
51 ±
10.
18
20.4
1± 5
.32
13.1
5 ±
1.87
0.
08
0.93
0.
10
Coe
lastr
ella
sp. (
V 3
) 0.
25
112.
19 ±
14.
98
26.9
1 ±
6.11
21
.98
± 1.
72
0.07
6.
49
0.81
C
oela
strel
la sp
. (P1
9)
0.29
14
7.38
± 1
2.87
38
.96
± 4.
99
26.4
7 ±
1.41
0.
08
0.47
0.
05
Coe
lastr
ella
sp. (
Tree
) 0.
32
90.1
8 ±
18.7
9 10
.05
± 1.
98
15.8
3 ±
1.65
n.
d.
n. d
. n.
d.
Chap
ter 3
113
Isol
ates
G
row
th ra
te
(OD
) B
iom
ass
prod
uctiv
ity
(mg
L-1d-1
) Li
pid
prod
uctiv
ity
(mg
L-1d-1
)
Lipi
d
cont
ent (
% d
ry w
t.)
Asta
xant
hin
prod
uctiv
ity
(mg
L-1d-1
)
Lute
in
cont
ent
(mg/
g)
Lute
in
prod
uctiv
ity
(mg
L-1d-1
)
*Coe
lastr
ella
sp. (
P63)
0.
37
152.
93 ±
13.
35
43.5
5 ±
6.28
23
.94
± 2.
65
0.13
0.
64
0.08
Scen
edes
mus
sp. (
Mn9
) 0.
29
120.
06 ±
17.
89
22.8
6 ±
3.29
27
.22
± 1.
60
0.08
1.
33
0.12
Sc
ened
esm
us sp
. (P1
55)
0.38
14
7.64
± 1
1.95
33
.53
± 2.
55
26.3
4 ±
1.97
n.
d.
1.13
0.
14
*Sce
nede
smus
sp. (
P58)
0.
47
155.
71 ±
14.
34
43.8
1 ±
5.15
32
.35
± 2.
49
n. d
. 0.
67
0.08
Sc
ened
esm
us sp
. (P1
15)
0.35
10
6.76
± 1
3.69
25
.25
± 6.
32
23.6
1 ±
2.10
n.
d.
n. d
. n.
d.
Scen
edes
mus
sp. (
V11
) 0.
23
133.
32 ±
16.
71
22.1
7 ±
2.65
19
.86
± 2.
29
0.03
0.
26
0.03
Scen
edes
mus
sp. (
V9
) 0.
42
138.
44 ±
17.
30
29.9
0 ±
5.13
23
.13
± 2.
07
0.23
0.
48
0.05
Sc
ened
esm
us sp
. (P1
52)
0.40
14
4.08
± 5
.59
22.6
7 ±
4.75
13
.60
± 1.
96
0.09
1.
8 0.
24
Coe
lastr
ella
sp. (
V8)
0.
37
83.3
3 ±
8.51
37
.32
± 1.
95
36.5
2 ±
1.81
0.
05
1.31
0.
09
*Sce
nede
smus
bi
jugu
(d
kh)
0.31
17
4.77
± 6
.75
40.1
4 ±
3.31
24
.24
± 1.
76
0.27
2.
9 0.
47
Scen
edes
mus
sp.
(V15
) 0.
41
127.
71 ±
15.
63
26.6
6 ±
2.62
25
.15
± 2.
04
0.08
1.
00
0.10
Visc
heri
a St
ella
ta (P
11)
0.26
12
0.24
±16.
01
16.2
4±1.
73
17.1
5 ±
1.31
0.
08
1.50
0.
13
Chap
ter 3
114
Tabl
e 3.
2 C
ompa
rison
of
lipid
pro
duct
ivity
of
mic
roal
gal
isol
ates
with
oth
er p
ublis
hed
data
.
Mic
roal
gal i
sola
tes
Lipi
d pr
oduc
tivity
(mg
L-1d-1
) R
efer
ence
s
Chl
orel
la sp
.(Q12
) 39
.72
± 3.
89
This
stud
y 16
h C
hlor
ella
SD
EC-6
9.
6 So
ng e
t al.(
2014
)24h
Chl
orel
la sp
. 83
.3
Che
irslip
and
Tor
pee.
(201
2) 16
h Sc
ened
esm
us sp
.(P58
) 43
.81
± 5.
15
This
Stu
dy16
h
Scen
edes
mus
biju
gus (
Lada
kh)
40.1
4 ±
3.31
Th
is S
tudy
16hr
Scen
edes
mus
biju
ga
8.22
Sh
in e
t al.(
2015
)
Scen
edes
mus
sp.
21.5
Tr
ipat
hi e
t al.(
2015
) AG
,18h
r
Scen
edes
mus
sp.
SD
EC-8
13
.52
Song
et
al.(2
014)
no
aera
tion
,N
and
P
Coe
last
rella
sp.
(P63
) 43
.55
± 6.
28
This
stud
y 16
h
Coe
last
rella
sp. M
60
13.9
K
arpa
gam
et a
l.(20
15) A
G,1
2hr
16h
Cul
ture
gro
wn
with
16h
ligh
t and
air.
24
h C
ultu
re g
row
n w
ith 2
4h li
ght a
nd a
ir.
AG
,18h
Cul
ture
gro
wn
with
agi
tatio
n an
d 18
h lig
ht
AG
,12h
Cul
ture
gro
wn
with
agi
tatio
n an
d 12
h lig
ht
N a
nd P
lim
itatio
ns c
ultu
res s
uppl
emen
ted
with
low
nitr
ogen
and
pho
spho
rus c
once
ntra
tions
Chapter 3
115
As shown in Figure 3.5, the colour of the microalgal isolates in the
Multicultivator MC OD 1000 varied among the isolates. Tubes 1and 2
turned yellowish-green during the early stationary phase (13th day), and
were harvested on the same day. Figure 3.4 shows that the biomass
exhibited two different colours: Orange and green. One of the yellow
greenish isolates was examined in bright field microscopy at the 5th, 9th
and 13th day. At the 13th day the isolates showed the appearance of
carotenoids balls inside the cell.
Figure 3.5 Changes in morphology and colour of Scenedesmus sp. at
stages of cultivation
3.3.4 Fatty acid production of microalgal isolates Fatty acids are divided in to two major lipid classes i.e the phospholipids
and glycolipids. Both the above classes of lipids are important part of
membranes associated with chloroplast and endoplasmic reticulum
(Olmstead et al., 2013). The fatty acid richness may varies between
different lipid classes (Wang and Wang, 2011).
Harvesting at 13 day Liquid culture day 13
Chapter 3
116
Besides FA profile, quantify fatty acid within these lipid classes and major
lipid classes is a necessity for quantifying microalgae lipid content. Fatty
acids in polar lipids can be converted to FAMEs; moreover the large scale
conversion of polar lipids to biodiesel has not been yet conventional.
Hence, the neutral lipid composition of microalgae lipid is utmost
important for biofuel production (Olmstead et al., 2013). Besides lipid
productivity, FA profiles of the isolates were further examined and
compared because FA composition and the types of FAs
produced are considered important for the quality of biodiesel. The
quality depends mainly on the unsaturation ratio because unsaturated
fatty acids (UFA) enhance cold- flow properties whereas saturated FAs
maintain good oxidative stability (Wu and Miao, 2014). In the present
study, the FAs were predominantly esterified and the proportions of
major FAs were determined at the average of nine days by GC-MS.
Although the FAs were very similar, the level of each FA varied
significantly (Figure. 3.5). Compared to other isolates, Scenedesmus sp.
(V15) produced 23.4% more oleic acid; Scenedesmus sp. (Mn25)
produced 22.8 % more; Chlorella sorokiniana (USA), 21.7%; and S.
bijugus (from Ladakh, India), 22.9% (Fig. 3.6). In fact oleic acid is one of
the most desirable oil components for biofuel since it gives a good
balance between cold flow property and oxidative stability (Hoekman
et al., 2012). All the above four isolates showed a good biofuel
characteristics. In the present study, Parachlorella kessleri was the only
isolate that produced large amounts of C16:0 (up to 59.1 %).
Microalgae rich in MUFA (particularly, palmitoelic acid (16:1) and oleic
Chapter 3
117
acid (18:1) (Hoekman et al., 2012) and SFAs are good for biodiesel
production (Stansell et al., 2012).
Figure 3.6 Fatty acid compositions (wt %) of twenty two microalgal isolates under test conditions. In each bar, fatty acids are shown in descending order of the number of carbon atoms in each
3.3.5 Estimation of biofuel properties To ascertain the suitability of the isolates as producers of biofuel, a few
other properties of the lipids, namely CN, IV, and SV, were analysed
based on FAME profiling. According to the EN14214 standard, SV and
CN should be neither higher than 120 g I2/100 g not lower than 47 g
I2/100 g (Sun et al., 2014). Six isolates in the present study met the CN
norms specified in USA (ASTM D6751) and twelve, those in Europe
(EN 14214) (Figure 3.6). Four isolates failed to meet any of the above
specifications; however, they be important because they have a high
(>13%) proportion of C18:3 acids. In the present study, IV for
Chlorella sp. (between 12 g I2/100 g and 103 g I2/100 g) was consistent
with that reported for Chlorella sp. by Wu and Miao. (2014).
0102030405060708090
100
Fatt
y ac
id c
ompo
sitio
ns (w
t %)
Microalgal isolates
C22:0C20:1C20:0C18:3C18:2C18:1C18:0C16:3C16:2C17.0C16:1C16:0C14:0
ASTM EN specification
Chapter 3
118
Thus, many of the isolates among the twenty two were suitable for
biofuel production and the rest had other valuable properties (Table
3.3). The high content of SFAs and oleic acid in a few of the
microalgae isolated in the present study shows their potential for biofuel
production
Chap
ter 3
119
Ta
ble
3.3
Perc
enta
ge o
f SFA
, MU
FA, P
UFA
.SFA
and
est
imat
ed b
iodi
esel
pro
perti
es fr
om th
e FA
ME
prof
ile o
f tw
enty
two
mic
roal
gal
isol
ates
M
icro
alga
isol
ate
SFA
(%
) M
UFA
(%
) PU
FA
(%)
CN
SV
(mg
NaO
H/g
of
oil)
IV (g
/100
g of
oil)
H
eatin
g va
lue
(MJ/
kg)
Bio
dies
el st
anda
rd
spec
ifica
tion
Chl
orel
la sp
.(Q12
) 27
.0
8.39
54
.78
47.5
5 18
8.41
12
3.17
39
.80
AST
M D
6751
Para
chlo
rella
kes
sleri
(Y8)
70
.6
0.53
0
80.1
9 15
9.46
1.
46
42.8
6 EN
142
14
Chl
orel
la so
roki
nian
a (U
SA)
24.6
7 22
.32
37.0
4 53
.98
176.
08
103.
61
40.6
3 EN
142
14
Chl
orel
la so
roki
nian
a (H
S)
33.0
8 3.
5 47
.36
50.7
9 17
2.88
12
0.34
42
.34
AST
M D
6751
Auxe
noch
lore
lla
prot
othe
coid
es (G
oa)
24.1
5 10
.48
44.1
50
.90
172.
93
119.
78
40.5
0 A
STM
D67
51
Scen
edes
mus
sp.(M
n25)
30
.86
25.8
32
27.2
48
.74
197.
37
112.
03
42.6
5 A
STM
D67
51
Coe
last
rella
sp.(T
n1)
31.2
13
.49
47.4
68
.88
137.
77
75.7
0 39
.60
EN 1
4214
Coe
last
rella
sp.(V
3)
18.4
3 2.
94
68.3
45
.52
192.
50
129.
45
39.4
3 -
Coe
last
rella
sp.(P
19)
34.2
8 13
.04
47.7
39
.29
184.
35
162.
71
42.1
3 -
Coe
last
rella
sp.(T
ree)
23
.94
15.7
61
30.7
4 64
.10
86.4
0 14
6.10
39
.83
EN 1
4214
Coe
last
rella
sp.(P
63)
30.1
18
.03
44.7
49
.61
193.
40
110.
69
40.0
8 A
STM
D67
51
Scen
edes
mus
sp.(M
n9)
13.8
1.
75
50.3
47
.03
179.
70
131.
73
39.5
6 A
STM
D67
51
Scen
edes
mus
sp.(P
155)
19
.4
11.2
2 56
.4
55.5
2 14
0.08
13
2.16
41
.70
EN 1
4214
Chap
ter 3
120
Mic
roal
ga is
olat
e SF
A
(%)
MU
FA
(%)
PUFA
(%
) C
N
SV(m
g N
aOH
/g o
f oi
l)
IV (g
/100
g of
oil)
H
eatin
g va
lue
(MJ/
kg)
Bio
dies
el st
anda
rd
spec
ifica
tion
Scen
edes
mus
sp.(P
58)
35.6
19
.79
23.0
42
.36
183.
09
149.
96
39.6
7 -
Scen
edes
mus
sp.(P
115)
35
.3
21.7
3 24
.4
63.7
4 16
2.66
71
.60
41.6
0 EN
142
14
Scen
edes
mus
sp.(V
11)
35.3
7 21
.75
24.4
61
.26
168.
94
77.0
8 41
.31
EN 1
4214
Scen
edes
mus
sp.(V
9)
20.0
5.
88
57.8
6 42
.31
176.
13
155.
42
39.9
2 -
Scen
edes
mus
sp.(P
152)
31
.57
24.0
9 35
.228
51
.95
188.
72
103.
41
40.1
0 EN
142
14
Scen
edes
mus
sp.(V
8)
30.6
22
.09
34.0
54
.59
180.
24
97.7
0 40
.58
EN 1
4214
Scen
edes
mus
biju
gus
(Lad
akh)
33
.51
26.0
2 31
.49
53.3
7 18
8.89
96
.96
40.2
3 EN
142
14
Scen
edes
mus
sp.(V
15)
41.9
24
.46
31.3
55
.79
178.
55
93.6
4 40
.00
EN 1
4214
Vi
sche
ria st
ella
ta (P
11)
21.8
4 26
.51
7.3
82.5
9 13
0.53
24
.53
43.7
0 EN
142
14
C
N,
cet
ane
num
ber;
SV, s
apon
ifica
tion
val
ue; I
V, I
odin
e v
alue
; SFA
, sat
urat
ed f
atty
aci
d; M
UFA
, mon
ouns
atur
ated
fat
ty
acid
; PU
FA,
pol
yuns
atur
ated
fa
tty a
cids
.
Chapter 3
121
3.3.6 Standard curve preparation for carotenoid analysis
All standards were prepared in the concentration range of 1-5 μg/mL and
stored at - 20 °C for further use. The calibration curves were prepared
using the previously described HPLC program (Refer Chapter 3, section
3.2.9). All the carotenoids present in the samples were calculated from
these standard curves (Figure 3.7)
Lutein
y = 43064x - 40048 R² = 0.9995
5000000
4000000
3000000
2000000
1000000
00 50 100 150
Conc (μg/mL)
Astaxanthin y = 12202x - 33517 R² = 0.9884
1000000
800000
600000
400000
200000
0 50 Conc (μg/mL)
Abu
ndan
ce (m
AU
)
Chapter 3
122
Figure 3.7 Calibration curves for each carotenoid standard
3.3.7 Lutein and astaxanthin productivity It is important to select isolates that have the potential to produce
large quantities of the targeted products, lipids and carotenoids if their
use is to be commercially cost effective. The ability of the isolates to
produce lutein and astaxanthin was also assessed. The major carotenoid
present was lutein, followed by astaxanthin. The isolates were grown
autotrophically and without any known stress in MC OD-1000 and their
production of intracellular lutein, astaxanthin and biomass was
recorded at an average of nine days (Table 3.1). Chlorella sp. (Q12),
the most productive isolate in the present study, produced more
Zeaxanthin 700000 600000 500000 400000 300000 200000 100000
y = 6955.1x - 40750 R² = 0.9981
Conc (μg/mL)
Canthaxanthin700000 600000 500000 400000 300000 200000 100000
y = 6144.8x + 32607 R² = 0.9944
Conc (μg/mL)
Abu
ndan
ce (m
AU
) A
bund
ance
(mA
U)
Chapter 3
123
biomass, lutein, and astaxanthin than the figures reported for Chlorella
protothecoides (Araya et al., 2014). Scenedesmus bijugus (Ladakh), the
second most productive isolate, was comparable to Scenedesmus sp.
(Yen et al., 2011). Sanchez et al. (2008) claimed that Scenedesmus
almeriensis are the best candidates with high lutein content. Two other
isolates, namely Coelastrella sp. P63 and V3, were also highly
productive. Isolates (V3) showed the highest lutein content (up to
6.4 mg/g) and high lutein productivity (0.8 mg L–1 d–1). Some of the
isolates demonstrated the potential to produce carotenoids as well as
lipids, a potential that could be exploited for obtaining multiple products.
On the basis of the above results the ten best species were ranking by lipid
and lutein productivity, as depicted in Figure. 3.8, with the size of the
point representing biomass productivity.
Figure 3.8 Top ten isolates showing lipid and biomass productivity
against lutein productivity (mg/L/d).
Biomass productivity Lipid productivity Lutein productivity
190 170 150 130 110
90 70 50 30
0.9 0.8 0.7 0.6 0.5 0.4 0.3 0.2 0 1
Microalgal
Chapter 3
124
3.4 Conclusion
The results from described in this chapter confirm that some of the
isolates Scenedesmus bijugus, Coelastrella sp., Chlorella sp., and
Auxenochlorella protothecoides showed the potential to produce
significant quantities of multiple products. The designed primer sets
provided biodiversity information and helped identify new species that
would have been difficult to identify through microscopic observations
alone. Scenedesmus bijugus was the most productive among the four
selected microalgal isolates. Future studies are required to maximize
metabolite production toward scale-up.
125
Chapter 4
Chapter 4: Influence of rate limiting factors on lipid, carotenoids
and biofuel properties in four selected microalgal isolates
4.1 Introduction Microalgae are known to produce metabolites such as pigments (lutein,
zeaxanthin and astaxanthin), long chain polyunsaturated fatty acids
(LC-PUFA) (Pal et al., 2011) which can be used as food and feed
additives, in the nutraceutical industry and for biofuel production
(Priyadarshani and Rath, 2012). Lutein and astaxanthin are important
carotenoids since lutein act as a primary carotenoids maintaining the
membrane integrity of cells and protects cells against other stresses
(Sanchez et al., 2008) whereas astaxanthin is a secondary carotenoid
which is present in lipid bodies outside the chloroplast (Grünewald et al.,
2001) with potential application in human health. Increasing demand for
carotenoids and their application in heath has generated considerable
interest. Several strategies are employed for improving microalgal lipid,
carotenoids and biomass productivity, including alteration in medium
composition (salinity, nitrates) as well as physical parameters (such as
light, temperature and pH) (Mata et al., 2010). Biomass tends to
decrease under high stress conditions employed, while excess energy
generated during the growth cycle gets converted in to lipids or
carbohydrates (Pal et al., 2011). Therefore there is a need for finding a
balance in rate limiting factors so that desirable changes/improvements
and can targeted.
126
Chapter 4
Numerous studies have shown that the lipid and biomass productivity is
effected by varying the culture conditions (Asulabh et al., 2012) and
diverse stress conditions such as nitrogen availability (Lin and Lin,
2011), salinity (Rao et al., 2007) and light intensity (Lv et al., 2010).
Scenedesmaceae is one taxa which is fast growing and can survive under
a wide range of environmental stresses, and so has been worked with
extensively (Chinnasamy et al., 2010; Ho et al., 2010). This organism is
known to accumulate 8-28 % of lipids as a percentage of dry weight
(Nascimento et al., 2013; Pan et al., 2011). Several reports describe the
effects of salinity on marine algae (Bartley et al., 2013; Martinez-
Roldan et al., 2014). However, comparatively less results are available
on the influence of salinity on cultivation medium for lipid production
by freshwater microalgae. Different strains of Scenedesmus can tolerate a
wide ranges of salinity, light and nitrogen concentrations (Lin and Lin,
2011). Salinity showed a pronounced effect on both lipid content and
fatty acid profile. Increasing salinity ranging from 0-50mM increased
lipid content and increased oleic acid C18:1 levels in Scenedesmus
obliquus (Salama et al., 2013) and so high salinity is useful in optimising
the growth of this microalgae (Kaewkannetra et al., 2012). NaCl also
affects lipid levels and biofuel properties in Desmodesmus abundans (Xia
et al., 2014) and promoted the accumulation of saturated fatty acid in
Botryococcus braunii (Zhila et al., 2011). Dunaliella sp. is an example
of an organism that can tolerate high salinity stress (Azachi et al., 2002)
with an ability to still produce high levels of lipid and biomass.
127
Chapter 4
Studies conducted by Karpagam et al. (2015) observed that salinity at
2% resulted in high lipid productivity in Coelastrella M60 (Karpagam et
al., 2015). Salinity concentration also impacts the lutein content within
microalgal cells. High light and salinity together resulted in increased
pigment production in Coelastrella F50 (Hu et al., 2013), whereas in
Scenedesmus sp. (Aburai et al., 2015) the fatty acid content decreased
and carotenoids content was increased under these conditions. High light
alone promoted the growth of microalgae but reduced overall lutein
content in Scenedesmus sp. (Xie et al., 2013). Therefore manipulating
the culture conditions for improving carotenoids and lipid biofuel
characteristics is important in make the biorefinery process cost effective.
The effect of phosphate and nitrogen starvation conditions on lipid
production has been studied by many researchers (Muthuraj et al., 2013,
2014). Nitrogen and phosphorus significantly impact biomass and lipid
production in microalgae. Lpid content is increased under decreased
nitrates and phosphate concentration in Scenedesmus KMITL
(Ruangsomboon et al., 2013) and Scenedesmus obliquus (Mandal and
Mallick, 2009). Since nitrogen and phosphate are essential for
metabolic activity of microalgal cell and biosynthesis. A concentration of
phosphate above 2μM resulted in inhibition of cells and reduced lipid
content in freshwater Scenedesmus obliquus (Martınez et al., 1999). The
combination of high light (420μmol/m2/s) and nitrogen starvation
conditions resulted in increase in lipid productivity (Ho et al., 2012).
Nitrogen starvation alone resulted in low lipid and biomass productivity
in Coelastrella sp. (Karpagam et al., 2015).
128
Chapter 4
Nitrogen starvation conditions were also reported to influence
astaxanthin production in Chlorella sorokianana (Raman and
Mohamad, 2012). Temperature (20-25 °C) has no significant effect on
lipid production per say (Wan et al., 2012), as reported in many
microalgal strains, such as C. sorokiniana, C. vulgaris and Scenedesmus
sp. Light intensity and photoperiods together effect microalgal
metabolite concentration in diverse microalgal strains (Seyfabadi et al.,
2011; Wahidin et al., 2013) and is strain specific (George et al., 2014).
A low cost source of nitrogen is cow urine. Cow urine used as a
nitrogen source resulted in increased biomass and lipid production in
Chlorella pyrenoidosa (Sharma and Rai, 2015). Therefore we used cow
urine and compared microalgal lipid and biomass production with a
control nitrogen source. The biochemical composition of microalgae
varies depending upon growth stage as well as cultivation conditions
(Yeh and Chang, 2012). two stage cultivation of microalgae strains
was used to maximize biomass in the first stage and lipid production in
the second phase, by applying several stress conditions in nitrogen
deficient medium (Markou and Nerantzis, 2013; Venkata Mohan et al.,
2012; Mujtaba et al., 2012; Xia et al., 2013; Zhang et al., 2014) in a few
studies, such as in Scenedesmus obtusus (Xia et al., 2013), in
Desmodesmus abundans (Xia et al., 2014) and in Dunaliella tertiolecta
(Takagi et al., 2006). Thus, isolation of locally characterised diverse
microalgae strains with optimised culture conditions, particularly NaCl
level alterations, are important for the commercial exploitation of
metabolites of interest for food and fuel in photoautotrophic two stage
129
Chapter 4
cultivation system. In the present study, four isolates were selected from
bioprospecting studies (Chapter 3) due to their ability to produce high
lipid, biomass and carotenoids in one growth cycle. These four isolates
demonstrated wide adaptability to salinity, light, nitrogen, and phosphate,
and were considered for maximising the production of lipids,
carotenoids and biomass. Based on the above literature we designed the
experiment where Scenedesmus bijugus (from ladakh) was grown in
continuous mode with multiple stresses such as salinity, nitrate and
phosphate during one growth cycle. Coelastrella sp. (V3) was grown
in a two stage system where the impact of light, photoperiod and
salinity stress on lipid, biomass and carotenoid production was
studied. Chlorella sp. (Q12) was grown in a two stage system where the
cells were first grown in optimised media conditions and then in a second
phase the stress was induced to investigate the influence on
metabolites and lipids. The final isolate, Auxenochlorella
protothecoides (Goa), was grown in a two stage cultivation system. Few
studies on the use of a two stage system for improving lipid production
have been published. To our knowledge this is first time that isolates were
subjected to multiple stresses and media amendments to attempt to
maximise biomass, lipid and carotenoids within one growth cycle. Earlier
results from two-stage systems reported an influence on lipid and
biomass, whereas we have attempted to enrich lipids and carotenoids
without negatively affecting biomass.
130
Chapter 4
4.2 Materials and Methods 4.2.1 Chemicals used The solvents and chemicals used in this study were of HPLC and
analytical grades. The details of solvents and chemicals used are same
as reported in (Chapter 3, section 3.2.1). The other medium
components such as NaCl, NaNO3 (Fischer scientific Waltham, MA,
USA), K2HPO4, KH2PO4 (Sigma-Aldrich, St. Louis, MO, USA) and
distilled cow urine was used for biomass, lipid and carotenoids
production. Experiments were performed in triplicate.
4.2.2 Strain selection and inoculum preparation Two freshwater isolates Scenedesmus bijugus designated as from Ladakh
(GenBank accession number KJ808696), Coelastrella sp. designated as
V3 (GenBank accession number JX513880), a third backwater isolates of
Chlorella sp. designated as Q12 (GenBank accession number KJ734869)
and a fourth terrestrial microalgal isolates of Auxenochlorella
protothecoides designated as Goa (GenBank accession number
KM020150), were examined for their potential for producing high
biomass, lipids and carotenoids in bioprospecting studies (Refer chapter 3
conclusion). These isolates were further maintained in a 20 mL liquid
broth in BBM medium for Chlorella sp., Coelastrella sp. and
Scenedesmus bijugus whereas in basal medium for Auxenochlorella
protothecoides were subcultured after every 7 days. For inoculum
preparation seed cultures of Scenedesmus bijugus, Coelastrella sp. and
131
Chapter 4
Chlorella sp. were prepared by inoculating 2% of culture in 40mL full
strength BBM medium, whereas for Auxenochlorella protothecoides
inoculum was prepared in basal medium in programmable MC 1000-OD
(Photon Systems Instruments, Drasov, Czech Republic) under 120μE/m2/s
light intensity with 16h of light and 8h of darkness for 5 days at 25 °C
with pure plain air supply at 0.12 min/mL. The medium pH was adjusted
to 6.8 prior to autoclaving at 121 °C for 20 min. The number of cells
were counted using a neurabeuer haemocytometer (Rohem Instruments,
Nashik, Maharashtra, India). The initial cell count of 3x106 cells/mL was
used as inoculum in all the experiments throughout.
4.2.2.1 Experimental design for Scenedesmus bijugus The experiment was conducted using 8 different treatments in MC
100-OD. The effect of nitrogen deficiency, salts stress (NaCl),
phosphates, and cow urine in BBM medium was studied under
photoautotrophic cultivation condition with light intensity of
250μE/m2/s under continuous illumination of 24:0h light and dark cycles
at 25 °C. Temperature was kept constant throughout. The following
treatments were defined prior to set up of the experiments:
A. Standard BG 11 medium
B. Bolds basal medium (BBM) +25mM NaCl and 20 μM Phosphate
(K2HPO4 and KH2PO4)
C. Bolds basal medium (BBM) + 50mM NaCl and 20 μM Phosphate
(K2HPO4 and KH2PO4)
D. Bolds basal medium (BBM) + Nitrogen depletion + 25mM NaCl and 20
μM Phosphate (K2HPO4 and KH2PO4) and cow urine (12%)
Chapter 4
132
E. Bolds basal medium (BBM) + Nitrogen depletion+50mM NaCl and 20
μM Phosphate (K2HPO4 and KH2PO4) and cow urine (12%).
F. Bolds basal medium (BBM) + 25mM NaCl and 20 μM Phosphate (K2HPO4 and KH2PO4) and cow urine (12%).
G. Bolds basal medium (BBM) + 50mM NaCl and 20 μM Phosphate
(K2HPO4 and KH2PO4) and cow urine (12%).
H. Bolds basal medium (BBM) standard and cow urine (12%).
All the above treatments were performed in MC 1000-OD (Photon
Systems Instruments, Drasov, Czech Republic) with white LEDs
containing 8 panels of 120mL volume capacity with working volume of
70mL and were kept the same in each treatment. Pure plain air of
0.12min/mL was provided to 8 panel‘s containing 70mL medium at an
equal rate using an air compressor during 24 hrs to ensure effective
mixing of culture into medium.
Screening of different salinity stress was carried out by replacing sodium
chloride (NaCl) concentration in the original BBM production medium
(0.42mM) with 25mM and 50mM of NaCl. Further screening of
phosphates 20 μM Phosphate (K2HPO4 and KH2PO4) replacing with
original phosphates in medium (0.43 and 1.29 mM concentration of
(K2HPO4 and KH2PO4) under photoautotrophic condition was carried
out. Cow urine (12%) was additionally added to BBM medium as
nitrogen source. Further nitrogen in the form of NaNO3 was
eliminated from production medium in treatments D and E. A 13 day
experiment was conducted in batch cultivation.
Chapter 4
133
The cultures in all treatments were harvested at 7 day when the cells
reached pre-stationary phase in order to investigate effect of all 8
treatments on lipid, biomass and carotenoids in Scenedesmus bijugus.
20mL of each culture from all treatments were harvested into empty
falcon tubes whose empty weight was recorded and these samples were
freeze dried (Chapter 3 Section 3.2.3). The weight after freeze drying
was recorded and subtracted from the empty weight of tube in order to
calculate the total biomass achieved at end of each phase of the
experiments. For subsequent analysis the remaining 50mL of culture
was left to continue to grow under similar conditions until they
achieved their stationary phase. At day 13 all the treatments reached
stationary phase, were harvested, and freeze dried in a similar way to the
pre-stationary phase samples.
4.2.2.2 Experimental design for Coelastrella sp.: Two stage
cultivation of Coelastrella sp. in MC 1000-OD
Experiments were designed in autotrophic two stage cultivation. In this
experiment two different medium were selected one was standard
BBM and the second was Bold 3N medium (triple nitrogen). The initial
cell count of 3x106 cells/mL was used as inoculum for all treatments. To
ensure effective mixing the culture were supplied with aeration of
0.12min/mL in all the treatments. The four different treatments were as
follows:
In first treatment, Coelastrella sp.was grown in 70mL of BBM medium
with initial cell count of 3x106 cells/mL in MC 1000-OD under low
light (50μE/m2/s) with 12:12h light: dark photoperiod at 26 °C till
Chapter 4
134
culture reaches the pre-stationary phase. At 7 day when the culture
reached the pre-stationary phase, 20mL cell suspension were harvested,
freeze dried and biomass was recorded for further analysis. The 50mL
dewatered culture from first stage was used as inoculum for second
phase of the experiment and reinoculated in to 50mL of fresh BBM
medium, where the light intensity was increased from 50μE/m2/s -250
μE/m2/s under continuous illumination (24:0h) to generate natural stress.
In the second treatment, the first phase B3N medium with amendment of
1.5% NaCl was used. The rest light and photoperiods were the same
for the first treatment phases 1 and phase 2. The only exception was at
the pre-stationary phase where 50mL of dewatered culture from the first
stage was used as inoculum for the second phase of experiment and
reinoculated in to 50mL of fresh B3N medium with 1.5% NaCl.
In the third treatment, the first phase culture of Coelastrella was grown
in 70mL of BBM medium with initial cells count of 3x106 cells/mL in
MC 1000-OD under high light intensity (250μE/m2/s) with 24:0h light:
dark photoperiod at 26 °C till culture reaches pre-stationary phase. At 7
day when culture reached pre-stationary phase, 20mL cell suspension
were harvested, freeze dried and biomass was recorded for further
analysis. The 50mL dewatered culture from first stage was used as
inoculum for second phase of experiment and reinoculated in to 50mL of
fresh BBM medium at where the light intensity was increased from
250μE/m2/s - 50 μE/m2/s under continuous illumination (12:12h) to
generate natural stress.
In fourth treatment, the first phase B3N medium with amendment of 1.5%
NaCl was used. The rest light and photoperiods were the same as for the
phase 1 and phase 2. The only exception that at pre-stationary phase,
Chapter 4
135
50mL dewatered culture from first stage was used as inoculum for the
second phase of experiment and reinoculated in to 50mL of fresh B3N
medium with 1.5% NaCl.
4.2.2.3 Experimental design for Chlorella sp. The experiment on Chlorella sp.was designed and performed in two
stage cultivation mode in MC 1000-OD with working volume of 70mL
for all the treatments. The initial cell count of 3x106 cells/mL was used as
inoculum throughout the experiment. To ensure well mixing the culture
were supplied with aeration of 0.12min/mL in all the treatments. The
first phase was divided in to 4 different treatments as follows:
I. Bolds Basal medium (BBM)
J. Bolds Basal medium (BBM) as suggested by Seham et al. (2014)
K. Bolds Basal medium (BBM), cow urine (12%) and AMF spore
extracts (100μL)
L. Bolds Basal medium (BBM) as suggested by Seham et al. (2014), cow
urine (12%) and AMF spore extract (100μL)
The second phase was divided in to 4 different treatments as follows:
A BBM B BBM as suggested by Seham et al. (2014) amended with 6 mM
NaNO3 and 5% NaCl
C BBM, cow urine, AMF spore extracts, 6mM NaNO3 and 5% NaCl. D BBM as suggested by Seham et al. (2014), cow urine, AMF spore
extract, 6mM NaNO3 and 5% NaCl.
In the first phase, Chlorella sp. were grown autotrophically in 70mL of
medium in MC 1000-OD. The 70mL tube were supplemented with
standard BBM medium and BBM with vitamins in treatment A and B.
Chapter 4
136
The medium as suggested by Seham et al. (2014) additionally contains
vitamins rest all components are similar to standard BBM. Further
combination of cow urine and crushed AMF spore extract were added in
BBM and BBM with vitamins in treatment C and D. After 7 days of
cultivation 20mL culture was harvested and freeze dried for further use
(Section 3.2.3). The light intensity at first phase was kept at 100μE/m2/s
with photoperiod of 16:8h light and dark cycle at 26°C for initial 7 days
of cultivation period till the culture reaches pre-stationary phase. In the
second phase, 50mL of dewatered culture pellet from phase 1 was
suspended in all the treatments from B-D by replacing original NaCl
and NaNO3 concentration of 0.25% and 2.94mM in both medium,
respectively in to 50mL of respective fresh medium supplemented with
final concentration of 6mM NaNO3 and 5% NaCl at end of pre-stationary
phase. Treatment A was kept as a control and after 7 day 50mL
dewatered culture was added in to fresh BBM medium. The light
condition and photoperiod were shifted from 100 μE/m2/s to
250μE/m2/s of light intensity under continuous illumination (24:0h) at
26 °C in all the treatments and was maintained throughout the
experiment.
4.2.2.4 Experimental design for Auxenochlorella protothecoides
The experiment on Auxenochlorella protothecoides was designed and
performed in two stage cultivation systemin MC 1000-OD with working
volume 70mL for all the treatments. The initial cell count of 3x106
cells/mL was used as inoculum throughout the experiment. To ensure
effective mixing the culture were supplied with aeration of 0.12min/mL in
Chapter 4
137
all the treatments. The following treatments were selected prior to start of
experiment:
A. Basal medium as per Arroyo et al. (2010)
B. Standard basal medium
C. Basal medium as per Arroyo et al.(2010) and AMF spore extract All the above three treatments were grown at light intensity 100 μE/m2/s
under 16:8 h light: dark cycle at 25°C. 20mL of each culture from all
treatments were harvested in to blank falcon tube whose empty weight
was recorded and freeze dried (Chapter 3 section 3.2.3). The weight
after freeze drying was recorded and subtracted from empty weight of
tube in order to calculate the total biomass achieved at end of each phase
of experiments. The remaining 50mL of culture was left to continue to
grow under similar conditions until they achieved their stationary phase
for subsequent analysis. At 13 day all the treatments reached
stationary phase, were harvested, freeze dried in similar way to pre-
stationary phase.
4.2.3 Measurement of algal growth Algal growth was determined by measuring absorbance at 680nm. Data
on the growth kinetics of all the isolates – three replicates for each –
were obtained during their culture. The microalgal cells were harvested, at
pre-stationary and stationary phase by centrifuging (10 min at 6000
rpm at 4 °C) followed by lyophilisation. The four selected isolates
were further evaluated for their efficiency in producing biomass (dry
biomass produced, expressed in milligrams per litre per day) and lipids
(expressed as a percentage of total biomass on dry weight basis).
Chapter 4
138
The pellets were washed twice with sterile deionized water, lyophilized
at –85 °C for 48 h, and their dry weight was determined
gravimetrically. For studying growth kinetics, all four isolates with
different treatments were cultivated in suitable medium and prescribed
culture conditions as described above. All the cultures with an initial cell
count of 3× 106 cells/mL were cultivated in a MC 1000-OD at 25 °C.
The pure air at 0.12 mL/min was maintained throughout the
experiment. Microalgal growth was monitored by measuring the daily
changes in OD at 680 nm with an OD viewer software attached to the
cultivator. The growth rate of microalgal isolates was estimated by fitting
the OD at the exponential phase of each isolates to an exponential
function, as follows (Wang et al., 2010):
(1)
Where OD0 is the initial OD and ODt is the optical density measured on
day t.
4.2.4 Biomass productivity The biomass productivity (BP, mg/L/d) was calculated using Eq. (6)
(6)
Where B2 and B1 represents the dry weight biomass at the time T (days)
and at the start of the experiment.
4.2.5 Lipid extraction and preparation of FAME Lipid productivity was obtained using Eq. (7)
The extraction method and GC analysis is described in Chapter 2 (Section
3.2.6)
Chapter 4
139
4.2.6 Gas chromatography This method is described in Chapter 2 (Section 3.2.7) 4.2.7 Evaluation of microalgal oil properties for biofuel This method is described in Chapter 2 (Section 3.2.8) 4.2.8 Determination of microalgal carotenoids Carotenoids productivity was measured using Eq. (2)
where Lp is lutein productivity, Ap is astaxanthin productivity, Bp is
biomass productivity, and Lc and Ac are lutein and astaxanthin
contents, respectively. The extraction method and HPLC analysis is
described in Chapter 2 (Section 3.2.9).
Chapter 4
140
4.3 Results and discussion 4.3.1 Growth curves for Scenedesmus bijugus To achieve higher biomass and growth rate, Scenedesmus bijugus was
grown in a BBM medium containing multiple stress factors such as
salinity (NaCl) at 25 and 50mM, phosphates (20μM K2HPO4 +
KH2PO4) and 12% cow urine (CU) for 13 days. Maximum OD values
of 2.70 and 2.62 mu (Absorbance units), respectively were obtained for
Scenedesmus bijugus grown in BBM + CU and BG11 (Figure 4.1).
Stationary phase for Scenedesmus bijugus was first achieved after 8
days of incubation in BBM amendment with 25mM NaCl, phosphates
and then under nitrogen deficient conditions with 25mM NaCl,
phosphates and CU. This clearly shows that cow urine promoted more
growth even when the medium was nitrogen deficient. Where as in all
other treatments the stationary phase was achieved at about 9 days of
incubations except BG11 which was used as standard medium in this
study reached stationary phase at 10 day of incubation. The growth
curves are shown in (Figure 4.1). The highest specific growth rate (h-
1) of 0.018 and 0.017 was achieved when Scenedesmus bijugus cells
were grown in BBM with 50mM NaCl, constant phosphorus and cow
urine and secondly in nitrogen deficient BBM medium with 20mM NaCl,
constant phosphate and cow urine. Where as in standard BG11 medium
0.13 (h-1) of growth rate was achieved. Other studies of Scenedesmus
sp. have shown growth rates up to 0.018 (h-1) at 5g/L of NaCl. Further
increase in NaCl concentration from 10-30g/L significantly reduced the
growth rate.
Chapter 4
141
However in the same species when NaNO3 was increased from 0.26 to
11.6 g/L the specific growth of alga increased from 0.015 to 0.035 (h-1)
(Peng et al., 2012).
Chapter 4
142
Figure 4.1 Combined effect of phosphate, salinity and cow urine on the growth of Scenedesmus bijugus A: BBM +CU B: BBM+ 50mM NaCl and 20 μM (K2HPO4+KH2PO4)
C: BBM+ ND+50mM NaCl and 20 μM (K2HPO4+KH2PO4) +CU D: BBM+25mM NaCl and 20 μM
(K2HPO4+KH2PO4)
E: BBM+ND+25mM NaCl and 20 μM (K2HPO4+KH2PO4) +CU F: BBM+25mM NaCl and 20 μM
(K2HPO4+KH2PO4) +CU
G: BBM+50mM NaCl and 20 μM (K2HPO4+KH2PO4) +CU H: BG-11
3 2.5
2 1.5
A B C D E F GH
1
0.5
00 2 4 6
Time (days) 8 10 12
Abs
orba
nce@
680n
m
Chapter 4
143
However, the effect of 8 different treatments on Scenedesmus bijugus
resulted in biomass concentration ranging from 153.1 ± 2.6 to 249.6 ±
6.8 mg/L/d at stationary phase (Figure 4.2). The highest net biomass
was found to be at 249.6 ± .8 mg/L/d in BBM supplemented with 12 %
cow urine. The use of cow urine as a nitrogen source resulted in an
increase in net biomass productivity for Scenedesmus bijugus. Whereas
the nitrogen deficient BBM medium with 25mM NaCl and constant
phosphate, the lowest net biomass productivity was achieved.
Moreover, when nitrogen deficient BBM medium with a combination
of two different concentration of 25 and 50mM NaCl was used, then net
biomass productivity increased from 227.9 ± 5.7 to 242.1 ± 7.3
mg/L/d at 13 day. The biomass productivity of BBM with cow urine
was higher as compare to standard BG11 medium. Second highest
biomass productivity of 243.1 ± 8.5 mg/L/d was achieved in BBM
supplemented with 20mM and 50mM NaCl and cow urine at 13 day.
Other studies have also shown increases in biomass productivity using
NaCl (20g/L) under high light intensity in Scenedesmus sp. (Peng et al.,
2012) whereas, NaCl at 5g/L increased net biomass in M. dybowskii
LB50 (Yang et al., 2015). In contrast one study on Scenedesmus
obliquus found that high net biomass (0.65g/L) was obtained using BBM
with 20mM NaCl rather than at 50mM which was much higher than
observed for BBM without NaCl (0.5 g/L) under low light conditions
(Salama et al., 2013), but this appears to be strain specific. In treatments
without cow urine, biomass was decreased due to the presence of high
salinity. However since phosphates were also amended in to the
medium there was not much significant difference seen in both the
Chapter 4
144
treatments as compare to BBM+CU. Where as in nitrogen rich BBM
with 50mM NaCl + 20μM phosphates + CU and nitrogen deficient
(ND) BBM supplemented with 50mM NaCl + 20μM phosphates + CU,
the biomass decreased from 198.7 ±7.2 to 167.0 ±4.4 mg/L/d. On other
hand in nitrogen rich BBM with 25mM NaCl + 20μM phosphates +
CU and ND BBM with 25 mM NaCl + 20μM phosphates + CU the
biomass again decreased in ND conditions from 243.1 ± 8.5 to 153.1±
2.6. The reason being under ND conditions the biomass is decreased
therefore growth is reduced. However a previous report suggested when
only nitrogen deficient condition was used with CO2 bubbling the
biomass productivity reached 840 mg/L/d in scenedesmus obliquus (Ho
et al., 2012) which, was observed in this study. The studies conducted
demonstrate that the growth of microalgae is suppressed either by using
excess sodium chloride or by insufficient concentration (Ruangsomboon,
2012). However, the presence of salt tolerant enzymes may also
function to increase the ability of microalgae to survive under different
range of salinitied, leading to increase in biomass growth of microalgae
at high salt level (Talukdar et al., 2012). Moreover, the biomass yield of
Scenedesmus sp. was decreased when eliminating NaCl from the
medium or in presence of NaCl after 12 days of incubation (El Sheek et
al., 2012). Thus addition of 25mM NaCl, 20μM phosphates in BBM
with cow urine has a significant impact on increased biomass
productivity.
No studies have yet been published on lipid, biomass and carotenoids
production using cow urine as a nitrogen source to increase biomass in
combination with other stress conditions. Only a few studies have
Chapter 4
145
reported the use of salinity to enhance lipid production in freshwater
microalgae Scenedesmus sp. (Kaewkannetra et al., 2012;
Ruangsomboon et al., 2012; Salama et al., 2013). Since NaCl and cow
urine are readily available and low cost sources of nutrients, we selected
them for lipid and carotenoids inducer in our study. The mechanism
behind lipid production due to salinity is yet not clear. From the
perspective of biorefinery process development, the productivity of end
product of microalgal isolates is important for cost effective process
development. Therefore, it would be best to achieve higher productivity
of end products at the same time in one growth cycle. In this work, we
found that cow urine as a nitrogen source can lead to increased net
biomass yield when other stress conditions are also implemented to the
medium. We were successful in achieving higher net productivity of
end products in one growth cycle in the Scenedesmus bijugus strain
isolated from high altitudes in India. However it is necessary to evaluate
for how long the microalgae should be grown under stress conditions.
In this work we harvested the isolates at pre-stationary and stationary
phase in order to obtain optimal lipid, biomass and carotenoids
production.
4.3.2 Different stress induction during pre-stationary and stationary phase 4.3.2.1 The biomass, lipid production of Scenedesmus bijugus in
response to different stress conditions at pre-stationary phase
We investigated two different concentration of NaCl with a
combination of other stress factors to maximise and achieve an optimised
yield of the products of interest. This study reveals that lipid productivity
Chapter 4
146
in Scenedesmus bijugus was dependent upon NaCl and cow urine
concentration in the medium. Lipid productivity was increased by adding
NaCl at 25mM, constant phosphates and 12% cow urine. The net lipid
productivity was evaluated first at 7 day (pre-stationary phase) and then at
13 day (stationary phase) of cultivation period. At 7 day highest lipid
content of 37.7 ± 0.4% was achieved at 25mM NaCl with cow urine,
constant phosphates as compare to standard BG 11 medium (Figure 4.2).
The values for lipid content were higher than achieved in Desmodesmus
sp. under high salinity or nitrogen starvation conditions (Pan et al.,
2011). A similar observation was also recorded with lipid content of 34.0
% in Scenedesmus obliquus at 25mM NaCl (Salama et al., 2013). The
lipid content in all treatments ranged from 12.5 - 37.7% at the pre-
stationary phase. These lipid content were higher than reported from the
genus of Scenedesmus by Gouveia and Oliveira. (2009), Mandal and
Mallick. (2009) and Ho et al. (2010). The effect of cow urine, salinity
and phosphates on lipid productivity are illustrated in Figure 4.2. As can
be seen in Figure 4.2 while cow urine (12%) was added to the
medium, significant difference between biomass productivity was
observed. The biomass and lipid productivities were higher when cow
urine was used as a nitrogen source. Addition of NaCl has a significant
impact on lipid productivity in Scenedesmus bijugus. The maximum
lipid productivity of 98.7 ± 1.0 and 81.8 ± 1.5mg/L/d were obtained in
BBM with 50mM NaCl, constant phosphates and ND BBM
supplemented with 25mM NaCl, constant phosphates (Figure 4.2)..
Moreover the lipid productivity in BBM with 50mM NaCl, cow urine
and constant phosphates were 40% less than in medium deficient of
Chapter 4
147
nitrogen and in rest all conditions were same. Since salinity alone in the
absence of nitrogen decreases lipid productivity. This result were constant
with Pal et al. (2011). Many reports suggest that lowering the sodium
nitrate concentration in the medium has a positive effect on increasing
lipid productivity. However total biomass productivity is reduced (Lv et
al., 2010; Peng et al., 2012). Higher lipid productivity was achieved in
cultures amended with 50mM NaCl concentration after 7 day of
incubation. Increasing salt concentration form 25 to 50 mM increased
lipid productivity from 81.8 ± 1.5 to 98.7 ± 1.0 mg/L/d (Figure 4.2).
Nevertheless, lipid productivity was higher in medium supplemented
with 50mM NaCl, BBM with cow urine and 50mM NaCl, nitrogen
deficient with cow urine. However salinity alone has also made a
significant impact on lipid productivity at a concentration of 50mM with
and without addition of nitrogen. The result highlights the use of
salinity, phosphates and cow urine as major influences on lipid and
biomass productivity at pre-stationary phase.
1: BG11 2: BBM+CU 3: BBM+ 50mM NaCl and 20 μM (K2HPO4+KH2PO4) 4: BBM+ 50mM NaCl and 20 μM (K2HPO4+KH2PO4) + CU 5: BBM+ND+25mM NaCl and 20 μM (K2HPO4+KH2PO4) + CU 6: BBM+ND+50mM NaCl and 20 μM (K2HPO4+KH2PO4) + CU 7: BBM+ 25mM NaCl and 20 μM (K2HPO4+KH2PO4) 8: BBM+ 25mM NaCl and 20 μM (K2HPO4+KH2PO4) + CU
148
Chapter 4
Figure 4.2 Biomass, lipid productivity (mg/L/d) and lipid content (% dry wt.) of Scenedesmus bijugus from same experiment
4.3.2.2 The biomass, lipid production of Scenedesmus bijugus in
response to different stress conditions at the stationary phase
To further evaluate effect of salinity stress (25 and 50mM), cow urine
(12%), nitrogen deficient conditions (zero nitrogen) and constant
phosphates (20μM), on the lipid productivity and lipid content in
Scenedesmus bijugus, lipid and biomass productivity was analysed after
13 days of cultivation. As shown in Figure 4.2, after 13 days of
cultivation lipid productivity (107.0 ± 1.22 ±0.3 mg/L/d and 107.3
mg/L/d) was much higher in (1) BBM amended with 25mM NaCl
and constant phosphates and (2) BBM grown with 50mM NaCl, cow
urine and constant phosphate, with a high lipid content of 31%, as
compare to standard BG11 medium. Whereas in BBM supplemented
with CU, high net lipid productivity of 117.00 ± 1.29mg/L/d with low
lipid content (27.2 ± 0.6%) was obtained. On other hand in BBM with
50mM and 20mM NaCl with constant phosphates, net lipid and total
lipids (%) was highest at 25mM than BBM with 50mM. which
0.005.0010.0015.0020.0025.0030.0035.0040.0045.00
0.0050.00
100.00150.00200.00250.00300.00350.00400.00450.00500.00
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16
Lip
id c
onte
nt (%
dry
wt.)
Bio
mas
s an
d lip
id p
rodu
ctiv
ity
(mg/
L/d
)
Treatments
Biomass producivity Lipid productivity Lipid content
7d 13d
149
Chapter 4
indicates that this may be the optimal NaCl concentration (25mM
NaCl) for growing scenedesmus bijugus. At pre-stationary phase at
50mM NaCl net lipid productivity was higher, however at the stationary
phase the net lipid yield decreased due to generation of high osmotic
stress. Moreover in BBM +25mM NaCl + 20μM phosphates + CU
and nitrogen deficient BBM + 25mM NaCl + 20μM phosphates + CU
less biomass was produced in ND conditions and net lipid productivity
was decreased, although total lipid content remained higher than with
BBM + 25mM NaCl + 20μM phosphates, as seen in Figure 4.2.
Therefore it is important to know that at what culture stage or
conditions one has to harvest the microalgal cells in order to extract the
highest yield of products at the same given time for efficient biorefinery
development.
In this study 25mM NaCl was determined to be optimal for enhancement
of lipid in Scenedesmus bijugus. Cow urine and phosphates resulted in
increased biomass growth whereas salinity resulted in increased lipid
production at certain limits before lipid start to decline.
4.3.2.3 Total fatty acid composition and biofuel properties of
Scenedesmus bijugus at pre-stationary and stationary phase
Besides lipid productivity, determination of the fatty acid profiles,
using FAMES analysis, for better quality of biodiesel from microalgal
isolates is vital. Cells growing in MC 1000-OD were examined for their
fatty acid composition under high salt, nitrogen and nitrogen deficient
conditions at the end of pre-stationary and stationary phase. It has been
reported that nutrient sources in the medium (salinity, high light and
nitrogen) affected the fatty acid profiles (Salama et al., 2013; Xia et al.,
150
Chapter 4
2013; Yang et al., 2015; Ho et al., 2012) in several microalgal strains
such as Chlamydomonas mexicana, Scenedesmus obliquus, Desmodesmus
abundans and Monoraphidium dybowskii LB50. High NaCl act a
stimulant for lipid enhancement in Scenedesmus sp. (Kaewkannetra et
al., 2012). It is generally known that under nitrogen-deficient
conditions, total fatty acid content is increased with decreased
biomass productivity (Pruvost et al., 2011; Rodolfi et al., 2009). In our
findings we were able to achieve higher biomass with high lipid
productivity using multiple stress factors.
Saturated fatty acids (SFA) C16:0 and monounsaturated fatty acid
(MUFA) C18:1 were the major fatty acid produced at two optimum
concentration of NaCl and nitrogen deficient medium (Table 4.1). This
results is consistent with that reported in Nannochloropsis sp.
(Solovchenko et al., 2014). Decrease in polyunsaturated fatty acid
(PUFA) under high NaCl at pre-stationary and stationary was previously
observed in freshwater Chlamydomonas mexicana and S. obtusus XJ-15
(Salama et al., 2013; Xia et al., 2013). Our findings were that the most
abundant fatty acid in pre-stationary phase were palmitic acid (C16:0),
stearic acid (C18:0), oleic acid (C18:1), linoleic acid (C18:2) and
linolenic acid (C18:3) (Figure 4.3). Traces of arachidic acid (C20:0) in
all treatments at pre-stationary and stationary phase were present,
whereas behenic acid (C22:0) 0.089 % was only present at stationary
phase in BBM medium supplemented with cow urine. Arachidic (C22:0)
which is a saturated fatty acid mostly present in fish and peanut oil (1.1–
1.7%) (Yeesang and Cheirslip, 2011), was also found in our isolates of
Scenedesmus bijugus (0.06– 0.15%). Oleic acid (C18:1) increased as a
proportion of total fatty acid (TFA) relative to palmitic (C16:0) and
151
Chapter 4
stearic acid (C18:0) in nutrient deficient medium supplemented with (1)
25mM NaCl and cow urine at pre-stationary phase; (2) 50mM NaCl
with cow urine at stationary phase (Figure 4.3) and is consistent with
results of Desmodesmus abundans producing 36.12% of oleic acid under
salinity stress (Xia et al., 2014) but contrasts with finding of Yang et al.
(Yang et al.,2015) in M. dybowskii who observed that oleic acid was
less in medium containing 20g/L NaCl and in two stage cultivation
process in S.obtusus (Xia et al., 2013)
Figure 4.3 FAME profile of Scenedesmus bijugus at 7 day of
cultivation under different tested conditions
-5
5
15
25
35
45
Com
posi
tion
of fa
tty
acid
(% d
ry w
t.)
Fatty acid
BG11BBM+CU BBM+ 50mM NaCl and 20 μM K2HPO4BBM+50mM NaCl and 20 μM K2HPO4+CU
BBM+ND+25mM NaCl and 20 μM K2HPO4+CU BBM+ ND+50mM NaCl and 20 μM K2HPO4 +CU BBM+25mM NaCl and 20 μM K2HPO4BBM+25 mM NaCl and 20 μM K2HPO4+CU
152
Chapter 4
Figure 4.4 FAME profile Scenedesmus bijugus at 13 day of cultivation
under different tested conditions
The major fatty acid elevated in all the treatment other than oleic acid
were palmitic (24.5 - 46.8% of TFA), stearic (4.99 - 21.7% of TFA),
linoleic acid (4.1 - 23.1% of TFA) and linolenic acid (2.8 - 10.5 of
TFA) at the pre-stationary phase. Whereas at the stationary phase
palmitic (24.5 - 39.2% of TFA), stearic (6.2 - 21.2% of TFA),
linoleic acid (4.8 – 10.8 % of TFA) and linolenic acid (2.7 – 12.9 of
TFA) were that major fatty acids (Figure 4.4). Siaut et al. (2011)
reported that nitrogen starvation effected the fatty acid profile. For all the
treatments we observed that nutrient deficient medium supplemented
with 50mM with cow urine produced a high percentage of oleic acid
(35.4%) at the stationary phase. In the stationary phase,
monounsaturated fatty acid (MUFA) increased significantly, whereas
linoleic acid decreased eventually. However, the level of C 18:1
increased significantly from 8.36 % from pre-stationary phase to 30.03%
at stationary phase in (1) nutrient deficient medium amended with 50mM
0
10
20
30
40
50
Com
posi
tion
of fa
tty
acifd
(% d
ry w
t.)
Fatty acid
BG11 BBM+ 50mM NaCl Land 20 μM K2HPO4 BBM+25mM NaCl and 20 μM K2HPO4BBM+ND+25mM NaCl and 20 μM K2HPO4+CU BBM+ ND+50mM NaCl and 20 μM K2HPO4 +CUBBM+CUBBM+50mM NaCl and 20 μM K2HPO4+CU
153
Chapter 4
NaCl and cow urine. These results shows that combination of salinity
and nitrogen deficient conditions can enhance the production of FA,
particularly oleic acid (C18:1), in Scenedesmus bijugus. However in
medium supplemented with 25 and 50mM NaCl with cow urine made no
significant increase both at log and stationary phase. The overall
variation in FA profile of the same isolates under different treatments
varies with the level of salinity and different physiological states (Rao
et al., 2007).
Cow urine appears to have no role in changing the fatty acid composition.
However optimum concentration of salinity and nitrogen deficiency
significantly changed the FA profile of this isolate. The above finding
indicate that this strain can be grown in all the treatments and provide a
desirable FA profile, particularly oleic acid (C18:1) and palmitic acid
(C16:0), for biofuel production using low cost nutrient medium (BBM)
and NaCl, cow urine as compare to BG11. Irradiance and salinity levels
have a positive effect in changing the structural and functional variation
within cells, such as alteration in fatty acid compositions (Kalita et al.,
2011). Moreover, the effect of salt stress on fatty acid composition of
freshwater microalgal species have not been well studied (Takagi et al.,
2006; Kacka and Do¨nmez, 2008). With increases in salinity, the
concentration of SFA decreases while UFA increases (Fujii et al., 2001).
Thus, it is important to evaluate the fatty acid composition of
microalgae cells at different growth phases. Biofuel quality parameters
such as cetane number and saponification value are linked with fatty acid
profile (Knothe, 2009). In desmodesmus abundans, biofuel
characteristics were improved further by the addition of NaCl at
optimum concentration by Xia et al. (2014). NaCl is readily available
154
Chapter 4
and an effective andcheap nutrient (Venkata Mohan and Devi, 2014; Yang
et al., 2014) and so was utilised as one of stress factors in our studies.
Our findings report that the fatty acid profile of all lipids not just TAGs
obtained in this study possess biofuel characteristics. The fatty acid
profile and the biodiesel properties were calculated using the empirical
formula as listed in Eq. (3-5) in chapter 3. The CN and IV (g I2 100 g-1)
were 70.08 and 55.1 from Scenedesmus bijugus in medium
supplemented with 25mM NaCl and cow urine which was higher than
standard medium at pre-stationary phase (Table 4.1). However medium
supplemented with 25 mM NaCl, CN and IV were 59.6 and 77.3, which
met the standard specification to European standard EN 14214.CN and
IV, were even high in medium with only salts (25 and 50mM NaCl) from
both pre-stationary phase and stationary phase. In all the treatments, IV
value showed better biofuel properties. The IV obtained in our isolates
Scenedesmus bijugus was higher than obtained in Scenedesmus obtusus
(Xia et al., 2013; Yang et al., 2015). In this study 25mM NaCl alone or
with cow urine resulted in high CN and IV value as shown in Table 4.1.
It can be inferred from this study that medium supplemented with NaCl
(25mM) and medium with only cow urine significantly effect the oleic
acid compositionin BBM at pre-stationary phase. Thus a combination of
NaCl and cow urine was the best low cost nutrients source for the
enhancement of lipids as well as improving the biofuel properties in
Scenedesmus bijugus. Our results also show that this isolate can be
grown in medium amended with high salinity, which makes this isolate
more suitable for biofuel production in a one stage process. Taken
together, growing this isolates in one stage process was more effective
155
Chapter 4
in terms of achieving high biomass with higher net lipid and desired
biofuel properties.
Chap
ter 4
156
Tabl
e 4.
1 Pe
rcen
tage
of S
FA, M
UFA
, PU
FA a
nd e
stim
ated
cet
ane
num
ber f
rom
the
FAM
E pr
ofile
of S
cene
desm
us b
ijugu
s at
7th a
nd 1
3th d
ay
Scen
edes
mus
biju
gus (
Lad
akh)
SF
A
MU
FA
PUFA
C
N
BG
11
68.8
10
09.
75
68.9
B
BM
+CU
40
.46
25
.44
61.5
B
BM
+ 50
mM
NaC
l and
20
μM
(K2H
PO4+
KH
2PO
4)
34.
6 31
.06
21.
03
59.
2
BB
M+
50m
M N
aCl a
nd 2
0 μM
(K
2HPO
4+K
H2P
O4)
+ C
U
39.
15
30.0
21.
2 5
5.7
BB
M+N
D+2
5mM
NaC
l an
d 2
0 μ
M
(K2H
PO4+
KH
2PO
4) +
CU
3
4.49
32
.9
22.
3 5
7.6
BB
M+N
D+5
0mM
NaC
l an
d 20
μM
(K
2HPO
4+K
H2P
O4)
+ C
U
33.
09
24.7
2
8.02
58
.0
BB
M+2
5mM
N
aCl
and
20
μM
(K2H
PO4+
KH
2PO
4)
36.
1 24
.63
17.
93
65.
6
BB
M+2
5mM
N
aCl
and
20
μM
(K2H
PO4+
KH
2PO
4) +
CU
36
.01
20.4
4 18
.09
70.8
Chap
ter 4
157
Scen
edes
mus
biju
gus (
Lad
akh)
SF
A
MU
FA
PUFA
C
N
BG
11
61.0
2 12
.02
10.0
8 70
.15
BB
M+5
0mM
N
aCl
and
20
μM
(K2H
PO4+
KH
2PO
4)
35.2
9 3
7.34
2
4.8
52.
92
BB
M+2
5mM
N
aCl
and
20
μM
(K2H
PO4+
KH
2PO
4)
34.7
6 3
1.29
2
1.07
5
9.26
BB
M+N
D+2
5mM
NaC
l an
d 2
0 μ
M
(K2H
PO4+
KH
2PO
4) +
CU
39
.20
30.
01
24.
9 5
5.73
BB
M+N
D+5
0mM
NaC
l an
d 20
μM
(K
2HPO
4+K
H2P
O4)
+ C
U
34.
4 3
1.9
22.
86
57.
64
BB
M+C
U
33.0
8 24
.72
27.1
58
.02
BB
M+5
0mM
N
aCl
and
20
μM
(K2H
PO4+
KH
2PO
4)+C
U
33.0
6 1
8.97
3
1.02
5
7.9
Chapter 4
158
4.3.2.4 Changes in carotenoid profile in Scenedesmus bijugus in
response to several stress factors
Microalgae also produces secondary metabolites in order to cope with
high level of salinity and further maintain osmotic balance within cells
(Hart et al., 1991). Salinity also induces changes in lipid especially
secondary carotenoids (astaxanthin) in several species of microalgae
(Imamoglu et al., 2009; Rao et al., 2007; Dominguez- Bocanegra et al.,
2004). Carotenoids production can be enhanced under different stress
conditions as they act as lipid soluble membrane antioxidants (Burja et
al., 2006; Campenni et al., 2013). There are many species which are
known to produce lutein, for example Chlorella zofingiensis (Del Campo
et al., 2004), Chlorella sorokiniana (Cordero et al., 2011), and
Scenedesmus obliquus (Chan et al., 2013). Alteration in cultivation
condition may improve lutein production under photoautotrophic
condition (Cordero et al., 2011; Del Campo et al., 2004). In tropical and
subtropical regions the cultivation cost for microalgal lutein production is
much lower because of longer exposure to sunlight as well ambient
temperature conditions prevailing throughout the year (Franz et al.,
2012). Both lutein and astaxanthin are useful carotenoids, with the
former acting as a primary source to protect cells from of stresses or
damages (Sanchez et al., 2008), and the latter having high antioxidant
capacity and application in human health (Dufosse et al., 2005). The
primary purpose of using NaCl, cow urine and BBM in the present
study is to decrease the cost of fermentation. The use of biorefinery
approach to produce multiple useful compounds reduces the cost of
Chapter 4
159
production and harvesting and eventually leads to a cost effective process
development. A reports suggests that Scenedesmus growing in BG11
account for 80 % more nitrogen than BBM (Pancha et al., 2014).
Strategies such as use of different combination of nitrogen deficient
medium, high and low NaCl concentration and cow urine could result in
high yield of lipids, biomass and carotenoids in one cultivation cycle by
reducing overall cost economics. As can be seen in Figure 4.5, the
carotenoid profile of this isolates comprised mainly lutein and
astaxanthin. Lutein was present in the free form whereas astaxanthin was
present as monoesters. The maximal productivity of lutein (0.48 mg/L/d)
and astaxanthin (0.22 mg/L/d) were recorded at (50mM NaCl + CU) at
the pre-stationary phase. The opposite trend was seen in medium
supplemented with (25mM NaCl + CU) where net astaxanthin
productivity (0.49) was more than lutein productivity (0.21), indicating
that the stress factors (NaCl and cow urine) both enhanced the production
of astaxanthin and lutein. The medium with low and high salinity
without cow urine yielded low lutein productivity as compared to
astaxanthin production because of less nitrogen availability. As we can
see above, in same treatment with cow urine amendment in to medium
yielded high lutein shows that cow urine was serving as a rich nitrogen
source in medium thereby enhancing lutein productivity. The above
results were consistent with Peng et al. (2012) in Scenedesmus sp. SP-01.
The lutein content decreased at 25mM NaCl concentration which is
consistent with the results reported by Cordero et al. (2011) in Chlorella
sorokiniana. In the above treatments, the effect was seen due to
combination of cow urine and NaCl. The low NaCl (25mM) + CU
Chapter 4
160
promoted increased production of astaxanthin as compared to lutein. On
other hand 50mM NaCl + CU increased lutein production as compared to
astaxanthin. Another study showed availability of nitrogen affects
carotenoid production in some of microalgae strains (Del campo et al.,
2007). Increased lutein accumulation in microalgal cell was observed
under increased nitrogen concentration (Fernandez-Sevilla et al., 2010).
Both lutein and astaxanthin productivity declined further as the culture
reached the stationary phase, as shown in Figure 4.5. The decline in
lutein productivity may be due to limitation of nutrients in the medium.
Astaxanthin was absent in the pre-stationary phase in nitrogen deficient
medium + NaCl initially, but subsequently it increased from 0-
0.08mg/L/d in both nutrient deficient conditions supplemented one with
25mM and other with 50mM NaCl + cow urine (Figure 4.5). The
above results show that the accumulation of astaxanthin occur when
stress is generated in medium (Del campo et al., 2004) but it was
opposite for lutein, since lutein production is limited under low nitrogen
concentration because of long exposure to high light intensity (Ho et al.,
2014). The ability of Scenedesmus bijugus to adapt and grow on
different salinity, cow urine and nitrogen deficient condition could be
of useful in biotechnology industries. Strain improvement strategies
using Scenedesmus bijugus might exceed the current production of net
lipid and biomass productivity making it a potential candidate for large
scale production.
Chapter 4
161
Figure 4.5 Lutein and astaxanthin productivity (mg/L/d) against biomass
productivity (mg/L/d) in Scenedesmus bijugus under different treatments.
4.3.3 Effect of light and photoperiods on growth of Coelastrella sp. Both light intensity and photoperiod effects the growth, biomass and
other metabolites of interest in diverse microalgae (Khoeyi et al., 2012;
Seyfabadi et al., 2011) such as Dunaliella sp. (Janssen, 2002);
Botryococcus braunii (Ruangsomboons, 2012) and Scenedesmus obliquus
(Mata et al., 2012a). Besides the photoperiod and light intensity,
duration of light exposure are key factors controlling the productivity
of microalgal culture (Qin, 2005; Krzeminska et al., 2014). One study
found that same species react differently to different light intensity and
photoperiods (Danesi et al., 2004). Another study reported that
increasing the light from low to high can improve the growth of
microalgae (Xue et al., 2011). This study showed that altering the
photoperiod along with light intensity and suitable medium in a two stage
cultivation process could replace the current strategies showing individual
effect of photoperiods and light intensities for a range of species.Initially
we examined the growth parameters for Coelastrella sp. cultured using
multicultivator 1000-OD in 70mL final volume in two different medium
0.6
0.5
0.4
0.3
0.2
0.1
0
450.00 400.00 350.00 300.00 250.00 200.0
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 Treatments
Lutein productivity Astaxanthin productivity Biomass
C
Chapter 4
162
(BBM) and (B3N amended with 1.5% NaCl) under high and low light
intensity (50 and 250 μmol/m2/s) and two different photoperiods (12:12h
and 24:0h L:D) at 26 °C. The culture were bubbled with air at a rate of
0.12mL/min. A two stage cultivation process was developed for
Coelastrella sp. to achieve higher biomass, lipids and carotenoids. The
different condition for growing Coelastrella were defined prior (as seen
in chapter 4 material and methods). At irradiance of 250 μmol/m2/s,
minimum growth rate (0.24 μ) was observed under 24:0h (Light: Dark),
exponentially phase continued for about 7 days and then start to decline
further in both the medium. At the irradiance of 50 μmol/m2/s,
maximum growth rate was achieved (0.56 μ) under 12:12 h L:D,
exponential phase was just for 3 days in BBM (Figure 4.6). However no
logarithm phase prevailed under 12:12 h (L:D) which was consistent with
strain of Nannochloropsis sp. cultivated at 50 μmol/m2/s (Wahidin et al.,
2013).
Figure 4.6 Growth pattern of Coelastrella sp. under different treatments. Error bar denotes mean ± SD.
.000
.500
1.000
1.500
2.000
2.500
3.000
3.500
0 2 4 6 8 10 12 14
Abs
orba
nce@
680n
m
Cultivation time (days)
B3N 12:12 (First phase) toB3N 24:0 (Second phase)
BBM 12:12 (First phase) toBBM24:0 (Second phase)
B3N 24:0 (First phase) toB3N 12:12 (Second phase)
BBM 24:0 (First phase) toBBM 12:12 (Second phase)
Growth phase I Growth phase II I
Chapter 4
163
However, in the second stage of cultivation, both medium cultivated at
12:12h (L:D) and low light intensity was harvested at 7 day. 20 mL was
harvested for further lipid and carotenoids analysis, with the remaining
50mL cell pellet being resupened in the same fresh medium with 250
μmol/m2/s light intensity under 24:0h (L:D) photoperiod. Material
grown from both medium at high light intensity with 24:0 h (L:D)
20mL was harvested for subsequent analysis and the remaining cell pellet
was resuspended in the same fresh medium with low light intensity
under 12:12h (L:D). BBM cultured at low light under 12:12h (L: D) at 7
days when shifted to high light under 24:0h (L: D), showed an initial
increase in optical density (OD) (Figure 4.6) and soon attained stationary
phase. On other hand BBM cultivated at high light under 24:0h (L:D) at 7
day when shifted to low light under 12:12 h (L:D), stationary phase was
achieved much earlier due to long time exposure of cells to high light at 7
days (Figure 4.6). In medium B3N supplemented with 1.5% NaCl at low
light under 12:12 h (L:D) at 7 day when shifted to high light 24:0h (L:D),
under such condition the OD value increased and reached highest of all
the conditions (Figure 4.6), which may be due to the presence of salinity
that result in overall increase in biomass growth of microalgae (Talukdar
et al., 2012). But in the same medium at high light under 24:0 h (L:D),
when shifted to low light 12:12 h (L: D), the cells grew for the first two
days and eventually declined after 9 days. We find that the long time
exposure of cells to high light under continuous illumination may cause
photo inhibition, which was consistent with resulted observed by Babu
and Binnal. (2015). Thus selection of appropriate light intensity and
photoperiods is important for growing isolates under suitable condition to
Chapter 4
164
produce high biomass and lipid levels. It is difficult to translate the results
of lab scale strain comparisons to large scale production due to variability.
Results indicate which combination is most suitable under two stage
cultivation for producing higher lipid and other metabolites without
losing the cell biomass.
4.3.4 Effect of light and photoperiod on the biomass and lipid production of Coelastrella sp. Light intensity and photoperiod has an effect on the biomass yield of
Coelastrella sp. at stage I and stage II cultivation as shown in Figure
4.7. Both light intensity and photoperiod are important factors for
determining optimal algae growth conditions (Parmar et al., 2011). High
biomass was attained when the light intensity and photoperiod were
shifted from low to high light intensities and 12:12 to 24:0h (L: D) in
both medium (Figure 4.7) which was consistent with finding of
Dumrattana and Tansakul. (2006) which report the best growth of
Botryococcus braunii was attained under continuous illumination.
However, a slight increase was also observed when the light intensity
and photoperiod were changed from high to low and 24:0 to 12:12h
(L: D) in B3N medium (Figure 4.7). A report suggests that increasing
the photoperiod to 12:12h (L:D) regimes resulted in decrease in growth
rate (George et al., 2014), which was opposite to what was observed in
our study.
Chapter 4
165
LL: Low light (50 μmol/m2/s); HL: High light (250 μmol/m2/s); B3N: Bolds triple nitrogen medium; BBM: Bolds basal medium
Figure 4.7 Biomass, lipid productivity and lipid content of Coelastrella
sp. under different tested treatments in two stage cultivation process.
In our findings a longer duration of light under continuous illumination in
the second phase brings about an increase in biomass, which was
consistent with finding of Khoeyi et al. (2012) in Chlorella vulgaris.
Another study suggested a shift of light intensity and photoperiod from
low to high or 12:12 to 24:0h (L:D) cycle results in a large increase in
biomass in Tetraselmis chui (Meseck et al., 2005), which is opposite to
what was observed by Wahidin et al. (2013) who report high biomass and
growth rate under 12 or 18 h light periods. Under high light
intensity,biomass productivity is greater in most species of microalgae,
however, different species respond differently to various light regimes
and photoperiods (Phatarpekar et al., 2002; Danesi et al., 2004;
Richmond, 2004). In this study, maximum biomass productivity of 277.8
± 5.9 mg/L/d was achieved when light intensity and photoperiod were
changed from 12:12h to 24:0 h (L: D); from low light to high light in
B3N medium supplemented with 1.5% NaCl. On other hand second
maximum biomass productivity of 220.8 ± 13.0 mg/L/d was achieved in
Chapter 4
166
BBM when light intensity and photoperiod were changed from 12:12 to
24:0h (L: D); from low to high light. The present study showed a 1.5
fold enhanced biomass productivity of Coelastrella sp. in second stage of
cultivation in comparison to 12:12 h (L: D) cycle. This result is
comparable to that obtained by Krzeminska et al. (2014) who reported a
total biomass concentration of 155.0 mg/L/d under continuous
illumination which was more than that reported by Ruangsomboon et al.
(2012) in Botryococcus braunii and in Scenedesmus obliquus (Ho et al.,
2012).
Strategies designed toward increasing biomass productivity and
metabolite levels were successful. We were able to achieve higher
biomass productivity in both media tested. The result showed that
Coelastrella sp. was able to produce higher biomass under both light
regimes and photoperiod in two different media in a two stage
cultivation process. Besides the higher biomass productivity, different
light regimes and photoperiods has a positive effect on net lipid
productivity. This studies showed that different photoperiods and light
intensities resulting in significant changes in the lipid profile of
microalgae (Harwood, 1998). Figure 4.7 shows the total lipid (% dry wt.)
and net lipid productivity expressed as milligram per litre per day
for Coelastrella sp. cultured at light intensity of 50 and 250 μmol/m2/s
under different photoperiod (12:12 and 24:0 h L:D) in two different
medium in two stage cultivation process. The maximum net lipid
productivity of 77.83 ± 0.29 mg/L/d was obtained when the culture were
exposed to high light under 24:0h (L: D) from low light with 12:12 h (L:
D) (from stage I to stage II) in B3N medium due to the presence of high
Chapter 4
167
nitrogen in the medium which subsequently increased biomass content
and second highest lipid productivity of 54.53 ± 0.71 mg/L/d was
obtained in BBM when the culture were shifted from high to low light
(from 24:0h to 12:12 h L:D) at the second stage (Figure 4.7). Our results
showed higher biomass and lipid productivity when compared to
Coelastrella sp. under 1 . 5 % salinity stress and high light
(400μmol/m2/s), which showed only 29 mg/L/d net lipid yield (Hu et al.,
2013). Another study by Weldy and Huesemann. (2007) showed under
higher light intensity that lipid production was enhanced in Dunaliella
salina. To our knowledge, this is the first study demonstrating the
effects of two different light intensities and photoperiods on lipid,
biomass and carotenoid production in Coelastrella sp. High lipid yields
obtained under higher light intensities may be due to storage of energy in
the form of lipid due to excessive stress (Asada, 1994). Non-increasing
lipid productivity in BBM under higher light intensities under continuous
illumination in second stage may be due to photo oxidative
mechanism where under high light chlorophyll molecules form an
unstable structure when exposed to high light intensity. The excited
oxygen reacts with fatty acid thus forming lipid peroxides resulting in
low FA production (Carvalho et al., 2011). On other hand, light plays an
important role in stimulating the fatty acid profile and growth in
microalgae (Khotimchenko and Yakovleva, 2005). However, several
external growth factors also regulate the lipid composition particularly
(fatty acid profile) in several organism. Figure 4.8 shows the fatty acid
profile of Coelastrella sp. under different light intensity and photoperiods
detected by GC. In stage I cultivated in two medium BBM (High light and
Chapter 4
168
low light) and B3N (High light and low light) under 12:12 and 24:0
(L:D) cycle, chains of C16:0, C18:0, C18:1, C18:2 and C18:3 were
most abundant fatty acids present (Figure 4.8). 11-eicoseonic acid
(C20:0) was present in all the treatments. Whereas in B3N medium at
low light intensity under 12:12h (L: D), 11 eicosenoic (C20:0),
docosenoic (C22:0) as well as tetracosanoic acids (C24:0) were present at
stage I (Figure 4.8).
Chapter 4
169
LL: Low light; HL: High light; B3N: Bolds triple nitrogen medium; BBM: Bolds basal medium
Figure 4.8 Fatty acid composition of Coelastrella sp. in two stage cultivation process
under different tested conditions
The use of different photoperiods and light intensity alter the fatty acid profile of
microalgae (Harwood, 1998). At stage I the culture growing in two different medium at
low light intensity with 12:12h (L:D) photoperiod, showed varied fatty acid profile. The
cultures growing in BBM medium showed a high percentage of two long chain fatty
acid palmitic (C16:0) fatty acids in stage I, whereas in B3N with 1.5% NaCl,
Coelastrella sp. produced more of C 18:3. On other hand the same isolate at high
light intensity under continuous illumination yield a high percentage of C18:1 fatty
acid but less C18:3 in both medium in stage I. However the high amount of C18:1
and C16:0 were present in BBM medium at stage I cultivation.
Moreover, when the cells were harvested and shifted from stage I at 7th day and re- inoculated the cell pellet in to fresh medium in second stage, the fatty acid composition
C16:0
C18:2
C16:1
C18:3
C16:2
C20:0
C16:3 C17:0
C22:0
C18:0 C18:1
C22:1 (w9) C24:0
30
25
20
15
10
5
0
Stage I Stage II
BB (12:12h)
LL
BB (24:0h) HL
B3N B3N BB (24:0h) HL
BB B3N B3N(12:12h) (24:0h) HL
LL (12:12h) (24:0h) HL (12:12h)
LL LL
Chapter 4
170
of same isolates behaved differently. It was interesting to note that the same isolate
when shifted from low, (12:12h) to high light, (24:0h), the proportion of C18:2 and C16:3
increased in both the medium at stage II achieving higher percentage of above fatty
acids in BBM medium (Figure 4.8). Moreover when the culture was shifted from high
light, (24:0h) to low light, (12:12h), not much variation was observed in the fatty acid
profile in both medium. The percentage of C18:1 was unaffected, however the
percentage of C16:0 was reduced in BBM medium. Thus shifting of isolates from low to
high light regime showed significant variation in terms of altering the FA profiling in
Coelastrella sp. The isolates growing in B3N medium at low light intensity at stage I
cultivation produced high percentage of C18:3 (19.5%) content but when shifted to
high light (24:0h) the percentage of C18:3 (15.7%) decreased in same medium. The
effect of four treatment at stage I cultivation showed that high light intensity with
continuation illumination, C18:2 fatty acid percentage increased in both medium in
B3N and BBM (Figure 4.8). High percentage of oleic acid (26.8%) was present in BBM
under 24:0h (L: D). Changes in oleic acid were also observed by Karpagam et al. (2015)
in Coelastrella sp. at high light intensity. On other hand at stage I high percentage
C16:0 and C18:0 were present in BBM at low light under 12:12h (L: D) cycle. Our
results on FA showed that the two stage cultivation of Coelastrella sp. in BBM
yielded high percentage of FA profile at stage II cultivation. The major fatty acid
effected was C18:2 (linoleic acid) under continuous illumination in stage I (Figure
4.8).
In our finding the high amount of SFA and PUFA were observed in BBM under
continuous illumination as compared to 12:12 h (L: D) at both stage I and stage II
steps of cultivation (Table 4.2). Moreover a high PUFA content was observed in
Chapter 4
171
Coelastrella sp. (Table 4.2), which may make this strain applicable to the production of
this nutritional ingredient. Moreover it does not happens with every species that the
light intensity and photoperiod impact the FA profile, suggesting light intensity did not
significantly affect the regulation of genes and activity of enzymes involved in the
biosynthesis of Desmodesmus sp. (Ho et al., 2014).
Besides the percentage of FA, lipid profile needs to be evaluated to compare the oil
with suitability as a biodiesel, based on the European and America biofuel standard
specifications. In the present study, we investigated the biofuel properties of
Coelastrella sp. The estimated CN and IV value for Coelastrella sp. was higher
(58.3 and 85.12 Iodine/100 g) when cultivated in BBM at low light intensity under
12:12h (L:D cycle) in stage I. Whereas in same treatment when shifted from low
light (12:12h) to high light (24:0h) in stage two cultivation, the CN and IV were
higher than obtained in stage I (55.8 and 97.4 Iodine/100 g) (Table 4.2) in this
treatment due to higher unsaturation. The CN value obtained in stage II cultivation was
higher than the CN reported for Coelastrella sp. by Karpagam et al. (2015). The
SV (175.1 KOH g-1 oil) for BBM at low light intensity under 12:12h (L:D cycle) was consistent with Coelastrella sp. (Karpagam et al., 2015). The CN value for this
isolates matched the biofuel standard specification which was not the case in
bioprospecting studies carried out in Chapter 3 (Figure 3.3). In the present study, we
were able to obtain an optimum strategy that could enhance the net biomass and lipid
productivity with desired biofuel characteristics in a two stage cultivation process.
C
hapt
er 4
172
T
able
4.2
SFA
, MU
FA, P
UFA
and
des
ired
biof
uel c
hara
cter
istic
s of
Coe
lastr
ella
sp.
in
two
stag
e cu
ltiva
tion
proc
ess
Tre
atm
ents
C
N
SV (K
OH
g-
1 oi
l) IV
(I2/
100g
) SF
A
MU
FA
PUFA
Stag
e I
B
B (1
2:12
h) L
L 58
.3
175.
19
85.1
2 42
.62
9.07
29
.63
BB
(24:
0h) H
L 46
.6
196.
04
122.
29
30.6
3 12
.76
50.2
5
B3N
(12:
12h)
LL
46
.1
181.
36
134.
49
26.6
10
.3
50.8
5
B3N
(24:
0h) H
L
47.7
18
2.00
12
6.66
20
.55
10.3
50
.87
Stag
e II
BB
(24:
0h) H
L 55
.8
173.
55
97.4
3 30
.01
10.5
54
.98
BB
(12:
12h)
LL
46.5
17
9.16
13
4.12
21
.11
9.53
55
B3N
(24:
0h) H
L
41.0
8 19
6.56
14
6.56
22
.97
11.8
60
B3N
(12:
12h)
LL
46
.3
188.
8 12
8.30
31
.01
6.8
52.3
5
CN
: Cet
ane
num
ber;
SFA
: sat
urat
ed fa
tty ac
id; M
UFA
: mon
ouns
atur
ated
fatty
aci
d; P
UFA
: Pol
yuns
atur
ated
fatty
aci
d; S
V: S
apon
ifica
tion
valu
e ;
IV: I
odin
e va
lue
Chapter 4
173
4.3.5Analyses of carotenoids from Coelastrella sp. The carotenoids from microalgal cells at stage I and stage II of
cultivation was analysed using HPLC. Algal cells accumulated
astaxanthin and lutein as major carotenoids at low and high light
intensity under two different photoperiods (12:12 and 24:0h L:D) in a
two stage cultivation process. The freshwater isolate Coelastrella
showed significant variation in carotenoids profile under four different
tested conditions. In stage I cultivation the Coelastrella sp. produced 0.46
mg/L/d net lutein in BBM at low light intensity under 12:12h (L:D)
(Figure 4.9). However in the same conditions, when shifted to high light
under continuous illumination, yield not change in stage II indicating that
exposure to high light did not impact growth.
Figure 4.9 Lutein and astaxanthin productivity of Coelastrella sp. in two
stage cultivation
1: BB (12:12h) LL; 2: BB (24:0h) HL; 3: B3N (12:12h) LL; 4: B3N (24:0h) HL; 5: BB (24:0h) HL; 6: BB
(12:12h) LL; 7: B3N (24:0h) HL; 8: B3N (12:12h) LL
2.5Stage II
2.00 1.50 1.00 0.50 0.00
1.60 1.40 1.20 1.00 0.80
1 2 3 4 5 6 7 8 Treatments
Lutein Astaxanthin
Stage I
Chapter 4
174
When the algal cells was cultured from low light (12:12h L:D) to high
light under continuous illumination the cells produced highest lutein
productivity of 1.45mg/L/d with 0.11 mg/L/d of astaxanthin (Figure
4.9). The astaxanthin was not detected at stage I in three treatments
except for BBM cultivated at high light under continuous illumination.
The net lutein (1.0 mg/L/d) and astaxanthin (0.17 mg/L/d) was highest
under BBM when shifted from 24:0 to 12:12h (L:D) cycle from high
light to low light. It was shown that the content of lutein and astaxanthin
were markedly increased in stage II cultivation. The algal cells cultivated
in B3N supplemented with NaCl from low to high light and BBM high
to low light enhanced the lutein and astaxanthin productivity in the two
stage process as shown in Figure 4.9. The high light intensity under
continuous illumination produced higher lutein productivity due to
presence of triple nitrogen in the isolate of Coelastrella sp. as suggested by
(Xie et al., 2013) and higher astaxanthin due to thre presence of salinity
in B3N at stage II because carotenogenesis was induced due to the
presence of NaCl. On the other hand algal cell cultivated in BBM when
shifted from 24:0 to 12:12h (L: D) cycle from high light to low light
lutein productivity was increased from 0.68 to 1.0mg/L/d (Figure 4.8).
This was consistent with study of Solovchenko et al. (2008) and Del
campo et al. (2004), suggesting increased lutein content under low light.
The absence of astaxanthin in the first stage in all three treatments was not
surprising since accumulation of astaxanthin happens as the cell growth
occurs and is expected to accumulate in cells at the exponential phase
(Del campo et al., 2004). This microalgae can sustain both low and high
Chapter 4
175
intensity under two different photoperiods.
Changes in photoperiod and light intensity have different effects on
biomass, lipids, FAME profile and carotenoids production in Coelastrella
sp. The results obtained in this work show that Coelastrella sp. is a
promising candidate for biofuel and carotenoids productions.
The percentage of astaxanthin and lutein can further be enhanced by
applying more specific rate limiting factors for enriching the production of
lutein and astaxanthin specifically. The varied productivity investigated in
present study showed that the same species responds differently to
different light intensity and photoperiods. The freshwater isolate
Coelastrella sp., examined for the first time in this study in a two stage
cultivation process was dependent on light and photoperiod conditions,
and seems to be a promising isolate for algal biotechnology. To the best
of our knowledge this species has not been studied much for biofuel
production and we investigated new strategies to enhance production of
lipid and carotenoids without losing cell biomass.
4.3.6 Cultivation of Chlorella sp. using two stage cultivation strategy Extensive literature on microalgae report that Chlorella, Scenedesmus,
Neochloris, Dunaliella, and Nannochloropsis genera are candidate strain
for the production of lipid rich biomass under stress conditions
(Gouveia and Oliveira, 2009; Rodolfi et al., 2009). The Chlorella sp. is
very well known for production of carotenoids and emulsifiers that are
used for the alimentary industry (Chisti, 2007; Ahmed et al., 2014),
omega 3 and omega 6 fatty acids as nutraceuticals (Adarme-Vega et al.,
2014) and the treatment of cardiovascular disease (Mozaffarian, 2005).
Chapter 4
176
Significant variation has been observed on growth and other
composition of microalgae under several stress conditions (such as
depletion or starvation of particular nutrient) (Metzger and Largeau,
2005; Qin, 2005; Takagi et al., 2000). Nitrogen is one of the most critical
factors influencing lipid production in microalgae (Sharma et al., 2012).
This research was conducted to discuss the combined influence of
nitrogen and salinity stress on growth, lipid and carotenoids production
in a two stage cultivation process. Many studies report cultivation of
microalgae in a two stage cultivation where in first phase the algae
were grown in nitrogen sufficient condition and in the second phase
stress is generated to increase the lipid production (Li et al., 2008;
Schnek et al., 2008). In many Chlorella strains under low nitrogen
deficiency the lipid content accumulate up to 40 % (dry cell wt.) in
C. vulgaris, and 23% C. pyrenoidosa (Ilman et al., 2000), moreover the
biomass is reduced under such conditions. On the other hand Huan et al.
(2010) mentioned that nitrogen sufficient medium also has a positive
effect on microalgal growth. Whereas salinity imposes stress in some
microalgae to induce lipids (Talebi et al., 2013). Thus combination of
both these factors may enhance biomass and lipid productivity at the same
time. The aim of the present study was to culture Chlorella sp. under a
two stage cultivation process in order to produced biomass rich in lipid
and carotenoids without loss of cell biomass. In the present study we
adopted a strategy where we used combination of nitrogen, salinity and
cow urine together in two different medium using two different light
and photoperiod conditions. Sodium nitrate is known to be a good
nitrogen source for growth and lipid of microalgae than other
Chapter 4
177
preferred nitrogen sources (Li et al., 2008). Our main focus was to
retain cell biomass levels in the second stage of cultivation. Initially at
stage I Chlorella cells were grown in nutrient sufficient condition at 100
μmol/m2/s under 16:8 h (L:D) and in the second stage the light and
photoperiod were shifted to 250 μmol/m2/s under continuous
illumination and cells were provided with optimum concentration of
nitrogen (sodium nitrate), salinity (sodium chloride) and cow urine. In
order to grow the Chlorella sp. for producing high biomass, and high
lipid and carotenoid, two different medium were tested with different
stress condition in two stage cultivation process. A commonly used
medium for growing Chlorella sp. was selected based on the literature
review. The Chlorella sp. cultivation under stage I showed similarities in
biomass production in both the medium. At the 6th day an amount (70
mL) was withdrawn from the MC 1000-OD and replaced with an equal
amount of fresh medium in second stage. As expected Chlorella sp.
reached the highest OD (3.0) in BBM (QA) and BBM with vitamins
(QD) in the second stage (Figure 4.10). We can see from Figure 4.10
that there was not much significant variation among the growth of
Chlorella in two different medium. As for Chlorella sp. there was a lag
phase just for one day initially. However, on the 2nd day after
inoculation the exponential phase started reaching OD of 2.90 in BBM
supplemented with 6mM NaNO3, 5% NaCl and cow urine (QC) (Figure
4.10). There was no significant difference in growth observed in the
control and the combined effect of salinity and nitrate except for BBM
supplemented with cow urine (12%) and AMF spore extracts (100μL),
Chapter 4
178
under which the growth of Chlorella cells was decreased at 6th days
(Figure 4.10).
Figure 4.10 Growth kinetics of Chlorella sp. in two stage cultivation process.
Start-up phase QA: Bolds Basal medium (BBM); QB: BBM with Vitamins; QC: BBM with cow urine (12%) and AMF spore extracts (100μL); QD: BBM with vitamins, cow urine (12%) and AMF spore extract (100μL) at light intensity of 100 μmol/m2/s with 16:8 h (L:D) at 26ᵒC Starvation phase QA: BBM; QB: BBM with 6 mM NaNO3 and 5% NaCl; QC: BBM with cow urine (12%) and AMF spore extracts (100μL) + 6mM NaNO3 and 5 % NaCl; QD: BBM with vitamins, cow urine (12%) and AMF spore extract (100μL) + 6mm NaNO3 and 5 % NaCl at light intensity of 250 μmol/m2/s with 24:0 h (L:D) at 26ᵒC. However in second stage, the replacement of cells in fresh medium with
nitrogen and salinity stress increased the growth of Chlorella sp. in
almost all treatments at high light intensity under continuous
illumination. Chlorella cells continued to grow exponentially up to an
average of 9 days in all the treatments. Similar kinds of results were
obtained in Chlorella salina when exposed to ranges of salinity (Talebi
et al., 2013). However the growth was inhibited in Chlorella emersonii
and Scenedesmus apoliensis (Demetriou et al., 2007) at high salinity
(Setter and Greenway, 1983). Chlorella sp. from backwater habitats
.00
.50
1.00
1.50
2.00
2.50
3.00
3.50
0 2 4 6 8 10 12
Abs
orba
nce@
680n
m
Incubation days
QA
QB
QC
QD
Start-up phase Starvation phase
Chapter 4
179
showed good exponential growth with combined effect of NaCl and
NaNO3 due to the presence of high amounts of sodium ions. The salinity
did not inhibit the growth of Chlorella sp.in the second phase. Since
Chlorella sp. is a promising strain for biofuel production, therefore such
ability of adaptation to high saline conditions with high nitrogen
concentration could be promising.
4.3.7 Effect of salinity, nitrogen and cow urine on biomass and lipid
production
In the present investigation, Chlorella sp. was evaluated in terms of
response to nitrogen and salinity stress in particular for increased
the production of lipid, biomass and carotenoids. Li et al. (2008)
suggested that high lipid content with high biomass is not achievable
under stress condition but this is not the case in our species. We
were able to achieve higher biomass productivities simultaneously with
higher lipid productivity. During the start-up phase not much significant
difference was observed in biomass growth of Chlorella sp. However
the lipid productivity showed variation among tested treatments. On the
contrary, during stress phase the biomass and lipid both increased
significantly under all condition except control without any stress
(Figure 4.11). Chlorella sp. was grown in two different media with a
combination of salinity and nitrate developed in stage II in order to
produce high biomass rich in lipids and carotenoids. In the starvation
phase, using BBM supplemented with 6mM NaNO3, 5% NaCl and
12% cow urine (QC), lipid productivity was highest, increasing from
14.2 ± 0.7 to 80.3 ± 2.4 mg/L/d. Whereas in the second medium BBM
Chapter 4
180
with vitamins amended with 6mM NaNO3, 5% NaCl (QB), lipid
productivity was at the second highest, increasing from 11.7 ± 0.9 to 74.3
± 1.2 mg/L/d, which was higher than observed by Illman et al. (2002)
in Chlorella sp. However, BBM with vitamins amended with 6mM
NaNo3, 5% NaCl and cow urine the lipid productivity increased from
11.0 ± 0.2 to 45.0 ± 0.7 mg/L/d but was not as high as for the above
treatment (Figure 4.11). In the control medium without NaCl, NaNO3
and cow urine, lipid productivity (36.0 ± 0.6 mg/L/d) was at lowest
(Figure 4.11). The increased in lipid production was because of higher
biomass and optimum salinity concentration used in present study. Similar
results were reported by Battah et al. (2012) where overproduction of
lipid was due to increase of salinity in Chlorella vulgaris.
Figure 4.11 Lipid content, biomass and lipid productivity of Chlorella sp. in two stage cultivation process
Start-up phase QA: Bolds Basal medium (BBM); QB: BBM with Vitamins; QC: BBM with cow urine (12%) and AMF spore extracts (100μL); QD: BBM with vitamins, cow urine (12%) and AMF spore extract (100μL) Starvation phase QA: BBM; QB: BBM with vitamins 6 mM NaNO3 and 5% NaCl; QC: Bolds BBM with cow urine (12%) and AMF spore extracts (100μL) + 6mM NaNO3 and 5 % NaCl; QD: BBM with vitamins, cow urine (12%) and AMF spore extract (100μL) + 6mm NaNO3 and 5 % NaCl.
250.00 Starvation phase
200.00
150.00
100.00
50.00
0.00
45.00
40.00
35.00
30.00
25.00
20.00
15.00
10.00
5.00
0.00 1 2 3 4 5 6 7 8
Treatments Biomass productivity Lipid productivity Lipid content
Chapter 4
181
A summary of result obtained through the two stage cultivation process
are summarised in Figure 4.11. Results indicated that biomass remained
unaffected under all the tested conditions in the two stage cultivation
process. There was a large increase in biomass in treatment QC and QD
from the stage I to stage II cultivation (Figure 4.11). The highest results
were achieved in the QC treatment, with biomass and lipid productivity
of 227.6 ± 5.6 and 80.3 ± 2.4 mg/L/d, respectively. The two stage
cultivation process was also performed on Chlorella vulgaris under
nitrogen and phosphate deficiency in a photobioreactor (Vaičiulytė et
al., 2014). However, our motive was to achieve higher production of all
metabolite while retaining higher cell biomass.
In this study, there was an inverse relationship observed between lipid
and nitrogen concentration. Chlorella sp. were able to produce high
lipids under nitrogen rich conditions along with salinity stress. Since a
combination of salinity and nitrate was used the balance was maintained
and we were able to achieve biomass rich in lipids.
Our strategies toward developing a two stage cultivation process for
higher biomass and lipid yield was successful. The combined effect of
salinity, nitrate and high light intensity under continuous illumination
provided varied results through various treatments for lipid and biomass
productivity.
4.3.7.1 FAME analysis
Biofuel characteristics are affected by the composition of fatty acids
(Knothe, 2008; Abou- Shanab et al., 2011). In the renewable oil
industry suitable algal candidates produce desired quality and quantity of
oil under a particular growth condition (such as temperature, light
Chapter 4
182
intensity and several stress conditions). The fatty acid profiling of the
microalgae affects the quality of the biodiesel. The length of the carbon
chain and the saturation of the fatty acids affect the biodiesel properties
which includes cetane number, cold-flow properties and oxidative
stability (Francisco et al., 2010). The fatty acids that enhance the
biodiesel properties include C14:0, C16:0, C16:1, C18:0, C18:1, C18:2
and C18:3 (Schenk et al., 2008). The most common fatty acids in
biodiesel are palmitic (C16:0), stearic (C18:0), oleic (C18:1), linoleic
(C18:2) and linolenic acid (C18:3) (Hempel et al., 2012). The fatty acid
methyl esters of Chlorella sp. were characterized by GC. In the present
study we found that cultivation of Chlorella sp. in a two stage cultivation
using two different medium has impacted the FAME profile (Figure
4.12). The unsaturated fatty acid increased, mainly C18:1, C18:2 and
C18:3 (alpha-linoleic acid – ALA), in both stages. The unsaturated
fatty acid EPA and DHA were not produced under these conditions.
C16:0,C18:1,C18:2 ad C18:3 were major fatty acid present in Chlorella
sp. previously reported by Tang et al. (2011) in Chlorella minutissima
(Petkov and Garcia, 2007). In start-up phase, Chlorella sp. produced
C16:0, C16:1, C16:2, C16:3, C18:0, C18:1, C18:2 and C18:3 fatty acids
(Figure 4.12). In the control treatment the percentage of oleic acid
(C18:1) was highest (16.5%). 11-eicosenoic (C20:0), docosenoic (C22:0)
as well as tetracosanoic acids (C24:0) were found in traces in all the
treatments. In the second phase the percentage of C18:2 was increased,
however the percentage of linoleic acid (C18:3) was reduced. Docosenoic
and tetracosanoic methyl esters were found in traces in all the treatments
in the starvation phase, except in the control. Under control conditions
Chapter 4
183
not much difference was observed in the fatty acid profile.
Figure 4.12 FAME profile of Chlorella sp. in two stage cultivation process under tested conditions. Start-up phase QA: Bolds Basal medium (BBM); QB: BBM with Vitamins; QC: BBM with cow urine (12%)
and AMF spore extracts (100μL); QD: BBM with vitamins, cow urine (12%) and AMF spore extract (100μL)
Starvation phase QA: BBM; QB: BBM with vitamins 6 mM NaNO3 and 5% NaCl; QC: Bolds BBM with
cow urine (12%) and AMF spore extracts (100μL) + 6mM NaNO3 and 5 % NaCl; QD: BBM with vitamins,
cow urine (12%) and AMF spore extract (100μL) + 6mm NaNO3 and 5 % NaCl.
The relative PUFA content ranged from 47.4 to 52.6% and 50.0 to 64.3%
in stage I and stage II cultivation, respectively (Table 4.3). Palmitic
acid was highest in the control in the starvation phase, while linoleic
acid ranged from 7.7 to 16.2 and 19.4 to 20.9 % in the start-up and
starvation phase (Figure 4.12). In the present study, in the second phase,
nitrogen availability and salinity stress has a significant impact on
increasing linoleic (C18:2) and alpha-Linolenic acid (C18:3), which is
consistent with other reports of Chlorella sp. (Liu et al., 2011a, 2011b;
Ratha et al., 2013) with high ALA content.
0
5
10
15
20
25
30
Q12 A Q12 B Q12 C Q12 D Q12 A Q12 B Q12 C Q12 D
Com
posi
tion
of fa
tty
acid
(%dr
y w
t.)
Treatments
C14:0 C16:0 C16:1 C16:2 C16:3 C17:0 C18:0 C18:1 C18:2 C18:3 C20:0 C22:0 C24:0
Start-up phase Starvation phase
Chap
ter 4
184
Tabl
e 4.
3 SF
A, M
UFA
, PU
FA a
nd b
iofu
el p
rope
rties
of
Chl
orel
la s
p. u
nder
teste
d co
nditi
ons
Tre
atm
ents
C
N
IV (I
2/100
g)
SV (K
OH
g-1
oil)
SFA
M
UFA
PU
FA
Star
t-up
phas
e
QA
46
.1
131.
18
185.
79
26.6
3 12
.8
49.9
4 Q
B
49.9
12
3.12
17
4.31
26
.70
11.3
45
.15
QC
49
.9
132.
06
163.
49
22.6
0 7.
98
47.4
1 Q
D
47.8
13
2.54
17
3.90
24
.32
9.85
48
.96
Star
vatio
n ph
ase
QA
45
.6
126.
68
196.
15
31.1
5 12
.68
50.0
2
QB
42
.3
142.
61
194.
28
24.2
7 12
.51
55.8
6 Q
C
36.4
16
3.39
20
2.58
22
.47
10.7
6 63
.26
QD
35
.6
165.
70
204.
72
21.3
11
.88
64.3
CN
: Cet
ane
num
ber;
IV: I
odin
e va
lue;
Sap
onifi
catio
n va
lue;
SFA
: Sat
urat
ed f
atty
aci
d; M
UFA
: Mon
ouns
atur
ated
fat
ty a
cid;
PU
FA: P
olyu
nsat
urat
ed fa
tty a
cids
.
185
Chapter 4
The combined effect of sodium nitrate and salinity increased
unsaturation of fatty acid levels, particularly for polyunsaturated fatty
acid, in Chlorella sp. Our finding reports that FAME profile of QB, QC
and QD meets the European biofuel standard specification in the start-up
phase. However, it does not meet the specification in the starvation phase
(second phase) due to increased unsaturation. Chlorella sp. has been
shown to produce omega fatty acid (Ratha et al. (2013)) and we founds
the highest percentage of C18:2 and C18:3 of 19.49 and 21.8 %,
respectively, were attained in BBM supplemented with 6mM NaNO3
and 5% NaCl with addition of 12 % cow urine. High nitrogen
concentration in medium in the starvation phase resulted in more
unsaturation and reduced biofuel quality but increased overall lipid
yield. A novel aspect of this study was that the Chlorella sp.
demonstrated a high percentage of ALA of 24.2 % in the start-up phase
and 21.8% in the second stage in BBM supplemented with 6mM NaNo3
and 5% NaCl with addition of 12 % cow urine (QC) (Figure 4.12),
which is high, but not as high as in flaxseed oil which contains about
57% ALA (Davis and Kris-Etherton, 2003). Our strategies toward
cultivation of Chlorella sp.in a two stage cultivation process was
successful. We were able to enhance the production of lipids and
biomass in the second phase along with minor variation in their fatty
acid profile. The result showed that the Chlorella sp.is well suited to be
used as food supplement in nutraceutical industry or as omega 3 fatty
acid. Other researchers have shown that cultivation of microalgal
under nitrogen deprivation can enhance the lipid production in a two
stage cultivation (Li et al., 2008; Rodolfi et al., 2009). However, it is not
186
Chapter 4
a case with all the species .The present isolates Chlorella sp. collected
from local habitat is able to yield higher lipid and biomass and alpha
linoleic acid (ALA) under combined influence of rich nitrogen BBM
medium and salinity stress. Thus, one must design strain specific strategies
that it result in high yields of all products of interest using low cost
nutrients, particularly in the very cost sensitive biofuel industry.
4.3.7.2 Carotenoids analysis Nitrogen availability and salinity impacts carotenoid production in several
microalgal strains (El-baky et al., 2004, Del campo et al., 2007). El baky et
al. (2004) studied the combined effect of nitrogen limitation and salinity
on Dunaliella salina. Some strains are also a good source of lutein (Wu et
al., 2009). Chlorella zofingensis and Chlorella protothecoides are
excellent lutein producer (Del campo et al., 2004; Shi et al., 2000).
Previous research indicates that light related parameters are promising in
terms of increasing lutein production in microalgae (Sanchez- Saavedra
and Voltolina, 2002; Solovchenko et al., 2008). Based on prior literature
we designed strategies for improving the production of metabolites along
with biomass. Thus we considered the combined effect of salinity and
sodium nitrate stress along with cow urine in Chlorella sp. in a two stage
cultivation process. The present study evaluated the feasibility of using a
Chlorella sp. to produce lutein and astaxanthin in the same growth cycle.
In the start-up phase the productivity of astaxanthin and lutein was
very low as compared to in the second phase. Under control conditions
astaxanthin productivity was not significantly affected. However,
variation in lutein production in control conditions was due to high light
intensity under continuous illumination. By employing the combined
187
Chapter 4
effect of cow urine, nitrogen rich condition with salinity under continuous
illumination we significantly enhanced the production of lutein from
0.23 to 0.74 mg/L/d (threefold enhancement) and astaxanthin from 0.12 to
0.l3 mg/L/d (Figure 4.13) in QC treatment, which is more than
reported in Chlorella sorokiniana by Raman and Mohamad. (2012).
The yield of astaxanthin reached 6.8mg/g in Chlorella zofingensis at
low salinity (Orosa et al., 2001) and that of lutein reached 4.97mg/g in
Desmodesmus under nitrogen rich condition (Xie et al., 2013). The
increased production of lutein was due to high light and nitrogen
enriched conditions whereas that of astaxanthin was because of salinity
stress under continuous illumination. Previous reports suggest that
nitrogen limiting condition have a negative effect on lutein production.
Thus nitrogen rich medium can enhance the production of lutein in
some microalgal strains (Borowitzka et al., 1991; Del Campo et al.,
2000; Fernandez-Sevilla et al., 2010). Another study by Cordero et al.
(2011) reports that increasing sodium nitrate concentration increased lutein
productivity whereas NaCl concentration higher than 2mM inhibited lutein
production in Chlorella zofingensis (Cordero et al., 2011). Thus it is
important that culture harvest be carried out the optimum time to
ensure higher productivity of lutein (Ho et al., 2014).
188
Chapter 4
Figure 4.13 Lutein and astaxanthin productivity of Chlorella sp. in
two stage cultivation process
Start-up phase QA: Bolds Basal medium (BBM); QB: BBM with Vitamins; QC: BBM with cow urine (12%) and AMF spore extracts (100μL); QD: BBM with vitamins, cow urine (12%) and AMF spore extract (100μL) Starvation phase QA: BBM; QB: BBM with vitamins 6 mM NaNO3 and 5% NaCl; QC: BBM with cow urine (12%) and AMF spore extracts (100μL) + 6mM NaNO3 and 5 % NaCl; QD: BBM with vitamins, cow urine (12%) and AMF spore extract (100μL) + 6mm NaNO3 and and 5 % NaCl Light is an important parameter for improving astaxanthin production in
microalgae (Imamgolu et al., 2009). The quantity of light (continuous
illumination) is more important than the quality of light for enhanced
astaxanthin production (Fabregas et al., 2001). Astaxanthin accumulation
in microalgal cells is mainly effected by high light intensity, salinity,
nitrogen limitation, together with other stress factors (Boussiba and
Vonshak 1991; Harker et al., 1996; Kobayashi et al., 1997; Fábregas et al.,
1998). The present work demonstrated the potential of Chlorella sp. for the
production of astaxanthin and lutein using a two stage cultivation process.
Chlorella sp. has shown that nitrogen rich conditions with high light can
increase the production of lutein, while salinity stress under continuous
0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
0.8
0.9
QA QB QC QD QA QB QC QDLut
ein
and
Ast
axan
thin
pro
duct
iviv
ty
(mg/
L/d
)
Treatments
Lutein productivity Astaxanthin productivity
Start-up phase Starvation phase
189
Chapter 4
illumination can enhance astaxanthin productivity.
4.3.8 Photoautotrophic growth of microalgae in Auxenochlorella
(Chlorella) protothecoides collected from terrestrial habitats.
Cells of Chlorella protothecoides were grown under photoautotrophic
conditions in two different medium. One was standard basal medium
and the other was basal medium as suggested by Heredia-Arroyo et al.
(2010). Three different variants were selected for growing the cells of
C.protothecoides; 1) Basal medium as per Heredia- Arroyo et al. (2010);
2) Standard basal medium and 3) Basal medium by Heredia-Arroyo et
al. (2010) co-cultured with arbuscular mycorrhizal fungi (AMF) spore
extract at constant light intensity of 100 μmol/m2/s under continuous
illumination at 25 °C. The crushed spores of Glomus intraradices were
used prior to co-culturing with microalgal cells at initial cell count of
3x106 cells/mL. In the growth analysis of C.protothecoides, no lag phase
prevailed in all the treatments. However in basal medium as per
Heredia-Arroyo et al. (2010), exponential phase continued until the 7th
day, whereas in standard basal medium the exponential phase lasted for
first 5 days (Figure 4.14). On other hand, a longer exponential phase (till
9th day) was observed in the basal medium, as per Heredia-Arroyo et al.
(2010), co-cultured with AMF spore extract. Co-cultivation of
microalgae and AMF spore extract (GC) under photoautotrophic
conditions increased biomass up to 180.47 ± 2.14 mg/L/d by the 6 day.
The culture in the same medium (GA) without AMF spore extract
achieved similar biomass productivity of 181.68 ± 7.24 mg/L/d,
followed by standard basal medium with net biomass of 129.30 ± 9.17
mg/L/d at 6th day of cultivation (Figure 4.15).
190
Chapter 4
GA: Basal medium as per Arroyo et al. (2010); GB: Standard Basal medium; GC: Basal medium as per
Arroyo et al. (2010) with AMF spore extracts at 100μmol/m2/s with 16:8 h (L: D) at 25 °C
Figure 4.14: Growth curves for Auxenochlorella protothecoides under
photoautotrophic conditions
3.0 Late log phase Stationary phase
2.5
2.0
1.5 GA GB
1.0
GC
.5
.0 0 2 4 6 8 10 12 14
Incubation days
191
Chapter 4
Figure 4.15 Total lipid content, biomass and lipid productivity of
Auxenochlorella protothecoides under photoautotrophic conditions at 6
and 12 day
1,4: GA: Basal medium as per Arroyo et al. (2010); 2,5: GB: Standard basal medium; 3,6: GC: Basal medium
as per Arroyo et al. (2010) with AMF spore extracts at 100μmol/m2/s with 16:8 h (L: D) at 25 °C
At 12 day of cultivation biomass decreased in GA and GD except for
the second treatment (GB) (Figure 4.15). Our finding indicates that a
symbiotic association did occurred between microalgae and AMF under
photoautotrophic conditions, as suggested by Gultom et al. (2014), in
Chlorella vulgaris under mixotrophic and heterotrophic condition using
fungal spores. This process of co-cultivation of microalgae with fungal
spores is innovative since the lipid constitute the major carbon (45 to
95 %) component of AMF spores and vesicles (Cox et al., 1975;
Jabaji-Hare, 1988).
To further investigate the potential of metabolites along with lipids in
C.protothecoides we examined the lipid productivity. The microalgae co-
cultivated with AMF spores showed higher accumulation of net lipid of
Biomass productivity Lipid productivity Lipid content
200.0180.0160.0140.0120.0100.080.060.040.020.00.0
Stationary phase
40.0
35.0
30.0
25.0
20.0
15.0
10.0
5.0
0.0 1 2 3 4 5 6
Treatments
192
Chapter 4
65.3 ± 1.0 mg/L/d and 51.3 ± 0.7 at 6 and 12 day as compared to
without any co-cultivation with AMF spores in the same medium
(Figure 4.15). The standard basal medium also showed increased lipid
productivity from 152 in day 7 to 497 mg/L in day 12, which was fivefold
more than reported for C.protothecoides under a two stage cultivation
process (Cheng et al., 2013; Araya et al., 2014), but was less than
reported for Nannochloropsis sp. (Chisti, 2007). The highest lipid
production in was in co- cultivated treatment and is attributed to the role
of AMF spores in increasing total lipids. High lipid content of 36.3 ±
0.1 and 34.3 ± 0.1 % was achieved in basal medium as per Heredia-
Arroyo et al. (2010) and the same medium microalgae co- cultured with
AMF spores (Figure 4.15), which was 4 times more than achieved
under autotrophic conditions in C.protothecoides by Santos et al. (2011).
In addition to lipid productivity, the microalgae biodiesel qualities were
estimated from its fatty acid profile. The same profile of fatty ester
methyl esters were detected in extracts obtained from co-cultured
treatments and standard medium. C16:0, C18:1, C18:2 and C18:3 were
predominant fatty acids present in the oil from all three treatments
(Figure 4.16). However, the exact percentage of each fatty acid varied
according to medium and growth conditions. For example, algal cells
grown in standard basal medium (GB) accumulated more of C18:1,
C18:2 and C18:3 at 12 days, whereas more C16:0 was present after 6
days. In co-culture treatment C18:2, C18:3 and C16:3 were the main
FAs, while in the same medium without co-culture C16:0 and C18:2
were the major fatty acid present (Figure 4.16). The microalgae co-
culture with AMF spores (GC) increased unsaturation of fatty acids in
193
Chapter 4
C.protothecoides. High oleic content of (15.5%) was obtained in
standard basal medium (GB). Generally high oleic acid content is
obtained in species under heterotrophic conditions as suggested by
various researchers (Xiong et al., 2008; Cheng et al., 2009). Another
study suggested that triacylglycerides (TAGs) constitute the major
component of spore extract (Jabaji-Hare, 1988; Gaspar et al., 1994), which
is consistent with the present study.
Figure 4.16 FAME profile of Auxenochlorella protothecoides at 6 and 12day The fatty acid profile obtained in the present study showed
accumulation of relatively high levels of Linoleic acid (C18:2) (Figure
4.16) which is consistent with the report by Cheng et al. (2013) in
Chlorella protothecoides. In fact, C.protothecoides in all the three
treatments meets the standard specification (Miao and Wu, 2006) based
on their FAME profile (Table 4.4) at 7 and 12 days.
C14:0 C16:0 C16:1 C16:2 C16:3 C17:0 C18:0 C18:1 C18:2 C18:3 C20:0 C22:0 C24:0
30
25
20
15
10
5
0 Goa A Goa B Goa C Goa A Goa B Goa C
Treatments
2
2
1
1
194
Chapter 4
Table 4.4: SFA, MUFA, PUFA and biofuel characteristics of
Auxenochlorella protothecoides
Treatments CN SFA MUFA PUFA
6 day Goa A 60.9 36.9 12.48 30.44
Goa B 67.6 36.7 15.9 21.62 Goa D 48.9 26.3 13.50 46.82
12 day Goa A 51.76 34.18 16.8 40.16 Goa B 48.79 30.14 20.64 43.02 Goa D 46.20 28.09 13.22 50.01
GA: Basal medium as per Arroyo et al. (2010); GB: Standard basal medium; GC: Basal medium as per Arroyo et al. (2010) with AMF spore extracts at 100μmol/m2/s with 16:8 h (L: D) at 25 °C
Our studies were primarily targeted to lutein and astaxanthin as target
carotenoid products. Lutein is a primary carotenoids (Gouveia et al.,
1996) and treatments impact its level during the growth phases. The
highest lutein productivity was achieved in standard basal medium
(17mg/L at 7 day). As cells reaches 12 day, lutein productivity decreased
in all the treatments since nutrients were utilised by the cells as the
growth prevailed. High lutein productivity of 2.0 mg/L/d was achieved
in standard basal autotrophic medium (GB) (Figure 4.17) which was a
2.5 fold enhancement compared to that reported by Araya et al. (2014)
in C.protothecoides, C.vulgaris and C.zofingiensis. The highest
astaxanthin accumulation was observed in GB at 6 day (Figure 4.17).
Microalgae co-cultured with AMF spores did not showed significant
variation in astaxanthin production at 6 and 12 day. Whereas at 6 day,
astaxanthin was not detected in basal medium (Heredia-Arroyo et al.,
2010), although it increased up to 0.06 mg/L/d at 12 day (Figure 4.17).
The second highest levels of astaxanthin and lutein production was
195
Chapter 4
achieved in cultured algae with AMF spores (Figure 4.17). However
more studies are required to understand the interactions between the
lipid and lutein accumulation. Co-cultured algae with AMF spores has a
significant effect on increasing biomass, overall lipid and carotenoids
productivity at 6 day in C.protothecoid
Figure 4.17 Lutein and astaxanthin productivity of Auxenochlorella
protothecoides at 6 and 12 day.
This is the first report of co-culturing microalgae with AMF spore extract
in autotrophic medium for C. protothecoides. Biofuel production from
C. protothecoides under autotrophic conditions resulted in lower lipid
productivity compared to heterotrophic growth condition. The analysis
of valuable co-products such as lutein and astaxanthin along lipids can
offset the cost of biofuel production under autotrophic conditions.
4.4 Conclusion Four novel strains selected from bioprospecting studies were subjected
to different stress conditions to produce high biomass rich in lipids and
carotenoids. Scenedesmus bijugus and Chlorella sp. grown in continuous
Lutein productivity Astaxanthin productivity
2
1
0
0.2 0.18 0.16 0.14 0.12 0.1 0.08 0.06 0.04 0.02 0
6Treatments
6
86
196
Chapter 4
mode and two stage system was able to grow well in salinity and nitrate
stress yielding high net productivity of lipids and carotenoids. When
compared to standard medium the or control, the fatty acid profile
obtained from growth in salinity and nitrate stress varied particularly
increased oleic acid in Scenedesmus bijugus and linolenic acid in
Chlorella sp. respectively. Scenedesmus bijugus has the potential to be
used in biofuel sector and neutraceutical industry. Whereas Chlorella sp.
has the potential to yield more of linolenic acid (C18:3)fatty acids. The
limitation of Chlorella in this regard is the relative absence of long chain
omega-3s (EPA and DHA). The main carotenoids in both the above
strains were lutein and astaxanthin. The stress conditions used in above
two strains improve their production, making them potentially useful
lipid and pigment producers. Low cost nutrient (NaCl, NaNO3) used in
this study reduced the overall production cost.
Coelastrella sp. a pigment producing strain which is photosensitive, was
grown in two stage system with alteration in light intensity and
photoperiod. The biomass was increased when light intensity and
photoperiod increased from low to high, 12:12 to 24:0h in both the
medium. Both the medium promoted lipid production, although the fatty
acid profile varied. B3N medium supplemented with 1.5 % NaCl when
shifted from low to high light intensity did enhance the overall lipid
productivity due to increased biomass. Both the medium have the
potential to produce lipid and carotenoids using varied light intensity and
photoperiod.
Auxenochlorella protothecoides when co cultivated with AMF spore
extract produced high net lipid productivity compared with strain grown
197
Chapter 4
without AMF spore extract. When compared to treatment without AMF
spore extract ,the fatty acid profile obtained from growth in AMF spore
extract varied, producing C18:2, C18:3 and C16:3 as major fatty acids.
AMF spore extract did enhance the carotenoids production when
compared to control.
Chapter 5
198
Chapter 5: Non-destructive extraction of lipids and recovery of active
cell biomass
5.1Introduction Reservoirs of fossil fuels are being depleted providing an opportunity for
biomass- derived fuels including from, given their higher productivity per
unit area (Yen et al., 2013). It is well established that microalgae have
the unique ability to tolerate and consume high levels of carbon dioxide
discharged from various industrial operations in the form of flue gas
(Singh et al., 2011). Microalgal biofuels have already proved successful
in trials, although cost remains the most serious constraint to greater use
of biofuels (Scott et al., 2010). Microalgae are known to produce
pigments such as lutein, zeaxanthin, and astaxanthin, long-chain
polyunsaturated fatty acids (LC- PUFA), and vitamins, which are of
economic importance (Pal et al., 2011) and are used as nutraceuticals
(Priyadarshani and Rath, 2012).
Several extraction methods has been adopted for milking of lipids and
carotenoids from microalgae (Vinayak et al., 2015). Organic solvent
extraction has been widely used for the extraction of lipids and
carotenoids (Cooney et al., 2009). In photosynthetic organisms, the
biocompatibility of some organic solvents has been studied (León et al.,
2001). However, selection of solvent is a critical approach (Leon et al.,
1998). Organic solvent extraction results in decrease of microalgal cell
biomass (Osborne et al., 1990). Two stage extraction using organic -
aqueous phases is a reliable approach based on immiscibility of polar and
non-polar solvents because these systems cut down the costly step of cell
Chapter 5
199
harvesting, whilst the product is continuously extracted. The phase contact
between cells and dodecane is toxic to the cells (Kleinegris et al., 2011).
Thus greater degree of solvent accumulation in the membrane was
equivalent with the higher rate of its toxicity to the cell (Kieboom et al.,
1998), hence solvent biocompatibility and non-toxicity is important
(Cequier-Sanchez et al., 2008). An alternative to lower the high cost of
biofuel production is ―Milking‘‘whereby the cells are subjected to
different conditions that release the metabolites of interest from the
cells without killing the organism. Algal cells can be maintained in a
stationary phase and nitrogen-limiting conditions for longer duration which
favor lipid accumulation in the cells. Furthermore, lipids can be milked in to
the algal suspension without requiring dewatering step (Zhang et al., 2011).
Milking of multiple products from microalgal cells using biocompatible
solvents can overcome some issues faced by organic solvent extraction
of lipids from algae and can make the extraction process highly
economically attractive (Hejazi and Wijffels, 2004). Firstly, the microalgal
cells remain viable and can be recycled in the biphasic sytem by
continuously producing lipids. Secondly, solvent can be recovered to
greater extend which improves the overall economic process (Mojaat et al.,
2008)
Several groups have attempted the biocompatible extraction of lipids
using milking (Santhanam and Shreve, 1994; Hejazi et al., 2002; Le´on
et al., 2003). The use of microalgae for the extraction of high value
products using biocompatible solvents was first studied by Frenz and
co-worker. (1989) and Ramachandra et al. (2009). The milking process
has an advantage that it does not require the use of continuous re-
growing of algal biomass, which take weeks for each cycle (Rickman
Chapter 5
200
et al., 2013). High value products can be milked out using
biocompatible solvents (Frenz et al., 1989), and osmatic shock (Sauer
and Galinski, 1998) in a continuous mode. Milking of lipids using
biocompatible solvents have proven to be effective for continuous
extraction and recovery of lipid and solvent (Hejazi, 2004; Wang et al.,
2009; Kang and Sim, 2007). The use of n-dodecane, which is one of
the best biocompatible solvent for milking of metabolites (da Silva et
al., 2006; Hejazi et al., 2003; Hejazi et al., 2002; Hejazi et al., 2004a;
Hejazi et al., 2004b; Hejazi and Wijffels, 2003). Mojaat et al. (2008), has
been used alone and in combination with other solvents, such as decane
and hexane, to increase the extraction efficiency of the metabolites.
Determining compatibility between the solvent and a specific algal strain
is an important requirement for effective extraction of compounds from
cells. Another consideration is cell wall thickness, which acts as a
barrier to solvent interaction with the internal algal cell components
(Wijffels and Barbosa, 2010), with interactions depending on molecular
structure of solvents (Hejazi at al., 2004).
Algal milking has been primarily applied to two species, Dunaliella
salina and Botryococcus braunii, for the extraction of pigments and
lipids from the aqueous organic phase (Hejazi et al., 2002; Hejazi,
2004). The former produced high amount of β-carotene using dodecane
as a biocompatible solvent. Although diatoms are of interest for milking
of lipids using biocompatible solvent, no experimental data has yet
been published (Vinayak et al., 2015). For successful milking, the
cells must be viable and can be regrown in a two stage system for
continuous milking of lipids from algal cells (Mojaat et al., 2008), and
Chapter 5
201
the mechanism of interaction of solvents with algae cells need to be
well understood (Zhang et al., 2011). Zhang et al. (2011) proposed to
select stress conditions where the species produces high levels of
metabolites. Limitations of lower recovery due to biomass reduction
when these stresses are applied can be compensated for through using
multiple milking cycles. Effective algal milking of microalgae can improve
the current economics of biofuel production making biofuel more
economically vaible (Hejazi et al., 2004).
In the work described we addressed our hypothesis number 3 where in n-
dodecane was used as a biocompatible solvent for milking algal cells to
recover lipids. The effect of solvent on algal growth, biomass, and lipid
was investigated under attenuated growth conditions achieved through
previous studies without negatively impacting cell biomass. We aimed
to milk lipids from the aqueous to the solvent phase. The influence of
solvent on microalgal cell shape was also studied. The isolate of
Scenedesmus bijugus was used in this study because of its potential for
producing lipid in comparatively higher quantities.
5.2 Materials and Methods 5.2.1Organism and culture conditions Scenedesmus bijugus, which accumulated high levels of biomass and
lipid in bioprospecting experiments described in chapter 3, was
selected for milking experiment. The freshwater isolate Scenedesmus
bijugus (from ladakh) belonging to the class Chlorophyceae was
cultivated in BBM medium (Anderson, 2005) in MC 1000-OD under
120 μE/m2/s at 25°C at 16:8 h (L:D) cycle. The pH of the medium was
adjusted to 6.8. Cells of Scenedesmus bijugus were also grown in
Chapter 5
202
medium supplemented with 25mM NaCl, 20μM phosphates and 12 %
cow urine. N-dodecane (Fisher Scientific Pittsburgh, USA) was used in
this experiment, since it is a good lipid extraction biocompatible solvent.
5.2.2 Experimental design Biocompatibility experiments were performed in MC 1000-OD consisted
of 120 mL of 8 panel glass tubes filled with 62 mL of medium and
8 mL of solvent for experimental treatments. The tubes were
maintained at a constant temperature of 25°C with plain air aerated at
the rate of 0.12 mL/min throughout the experiment. For inoculation of
all treatments, algae suspension was allowed to grow in BBM medium
for 10 days until it reaches the exponential phase. After 10 days, a cell
count was performed using Neubauer haemocytometer (Rohem
Instruments, Nashik, Maharashtra, India) and an initial cell count of
3x106 cells/mL was inoculated in all three treatments (T1-T3) as shown
in Table 5.1. The algae - solvent interaction was studied for 12 and 18
days. T1 used only algal suspension without solvent followed by T2
containing algal suspension with n-dodecane. T1 and T2 were
maintained at 120 μE/m2/S light intensity at 25°C under 16:8h (L:D)
photoperiod. On other hand, treatment 3 used the addition of 25mM
NaCl, 20μM phosphates, and 12% cow urine with n-dodecane (Table 5.1)
cultivated under continuous illumination (24:0h) at 250 μE/m2/S at 25°C.
The steps involved throughout the experiment are depicted in Figure
5.1.
5.2.3 Algal growth determination Rapid growth for all the isolates was achieved during their
Chapter 5
203
cultivation. Cells of Scenedesmus bijugus were inoculated with an initial
cell count of 3×106 cells/mL in a MC 1000-OD at 25°C under the
conditions described in section 5.2.1 for each treatment. The OD of
microalgal cells were measured at 680nm with OD viewer software
attached to the cultivator. The growth rate of microalgal isolates was
estimated by fitting the OD for exponential phase in each isolates to an
exponential function (Chapter 3 section 3.2.3). The microalgal cells were
harvested at 12 and 18 day by centrifugation followed by lyophilisation.
5.2.4 Cell count The cell count was performed for Scenedesmus bijugus at 12 and 18
days using a simple dilution plating method for counting the number
of live cells. One mL of algal suspension at 12 and 18 day was diluted
up to 10-6. 100 μL was further plated on agar plate (1.6% w/v) and
incubated for 3 weeks until colonies appear.
5.2.5 Scanning electron microscopy (SEM) SEM was performed on samples of algal cells for T1-T3 treatments at day
12 and 18 to characterize the morphology of Scenedesmus bijugus
before and after solvent contact. Refer for protocol in chapter 2 (section
2.2.4).
5.2.6 Algal lipids analyses
Lipids were analysed in the organic phase and algal cells. Lipids in algal
cells were extracted using the method described in section 2.7 of chapter
3. Method described by Christie and Han. (2010) was employed to
study the FAME profiles in the organic layer. The samples of FAMEs
were analysed by gas chromatography (Agilent 122-2332 column, Santa
Chapter 5
204
Clara, California, USA) with mass spectrometry ( MS) as described in
chapter 3, section 2.8.
Cha
pter
5
Tabl
e 5.
1 Tr
eatm
ents
per
form
ed to
eva
luat
e lo
ng te
rm m
ilkin
g of
alg
ae
Tre
atm
ent c
ode
Tot
al
days
Sp
ecie
s T
reat
men
t
T1
18
Scen
edes
mus
biju
gus
BB
med
ia w
ithou
t rat
e-lim
iting
am
endm
ents
T2
18
Scen
edes
mus
biju
gus
BB
med
ia w
ith n
-dod
ecan
e
T3
18
Scen
edes
mus
biju
gus
BB
med
ia +
25 m
M N
aCl +
20 μM
K2H
PO4
+
cow
urin
e (1
2%) w
ith n
- dod
ecan
e
20
6
207
Chapter 5
Figure 5.1 Flow diagram showing steps involved throughout the experiment
5.3 Results and Discussion
Biocompatibility experiment were performed to milk out lipidsfrom
cells of Scenedesmus bijugus in the solvent phase without killing algal
cells. The biocompatibility of n-dodecane to Scenedesmus bijugus were
investigated by rapid growth determination, biomass production and cell
viability. The effect of n- dodecane on microalgal lipids profile in
organic phase and algal cells were further analysed. SEM studies were
performed to study the effect of solvent on the cell surface. The biomass
at 18 day of cultivation was harvested and re-inoculated in to fresh
medium, grown for another 18 days and was analysed further for
recovery of biomass and fraction of lipids in organic phase.
T1: BBM without n- dodecane
Scenedesmus bijugus cells
Recovery of lipids from cell biomass at
18 day
Inoculated with initial cell count (3X106
cells/mL) in T1-T3
T2: BBM with n- dodecane
Recovery of biomass from fresh medium at
18 day
Growth measurementt @
680nm
T3: BBM amended withstress with n-
Analysis of lipid in organic phase and algal cells at 12 and 18 day
Cell count 12 and 18 day
SEM analysis before and after solvent exposure at 12 and 18 day
208
Chapter 5
5.3.1 Effect of organic solvents on algal growth and cell viability
Dodecane was used as a biocompatible solvent for the culture of
Scenedesmus bijugus in two treatment T2 and T3 (Table 5.1). The
viability of the cells in dodecane was judged by colony count. The color
of the culture medium was green after 24h solvent exposure. The solvent
treated cells were still viable and were similar to control conditions even
after 12 and 18 days exposure (Figure 5.2). The effect of the solvent
treated condition on Scenedesmus bijugus were analysed by
determining the biomass concentration. Figure 5.2 shows the biomass
production in three different treatment at day 12 and 18. As shown in
Figure 5.3 cell growth was not affected by n-dodecane and cells grew
faster than control condition in the presence of medium with solvent at
12 and 18 day. The specific growth rate of Scenedesmus bijugus in
solvent (0.36 μ) and salinity with solvent treated conditions (0.38 μ)
was higher than control conditions (0.28 μ). The presence of organic
solvent is known to increase the oxygen solubility in medium and
thus increase overall photosynthetic rate (León et al., 2001).
Figure 5.2 Culture grown under three different treatments in aqueous phase (T1), aqueous – organic phase (T2) and aqueous (salinity stress)-organic phase at 12 and 18 day.
T1 T1 T2 T2 T3 (12) T3
209
Chapter 5
The net biomass concentration of 3.7 g/L was achieved at 18 day in
medium with solvent treated condition as compared to control without
solvent (2.7 g/L). In salinity with solvent treated conditions the biomass
concentration was less than with medium with solvent at 12 and 18 day
(Figure 5.4). The results of total cell count showed there were
differences in the ability of cells to grow in all the three different
treatments. The cell count as well a growth rate showed that
Scenedesmus bijugus had viable cell growth after contact with solvent
at 12 and 18 day. The cell count increased and reached the highest level
if 63 x 106 cells/mL and 72 X 106 cells/mL in salinity with solvent and
solvent treated algae, respectively, at day 18 compared to control
without solvent (45 x 106 cells/mL).
Figure 5.3 Effect of n-dodecane on Scenedesmus bijugus growth in an aqueous–
organic system and non organic systems and non-organic systems
Abs
orba
nce@
680n
m
3.5 BBM BBM + Solvent BBM +25 mM NaCl + 20 μM phosphates + CU with solvent
3
2.5
2
1.5
1
0.5
0 0 2 4 6 8
Time (days) 10 12 14 16 18 20
210
Chapter 5
Figure 5.4 Biomass productivity of Scenedesmus bijugus at 12 and 18
day with and without solvent treated algae.
5.3.2 Effect of n-dodecane on lipid profiles Lipid milking was monitored by analysing the types and amount of
lipids and fatty acids extracted in the solvent phase and for algal cells.
The milking of Scenedesmus bijugus was able to extract and concentrate
lipid in the solvent phase. Figure 5.6 presents the percentage profile of
fatty acid (% dry wt.) in the solvent and algal cells.
The effect of n-dodecane on fatty acids accumulation in solvent and algal
cells was investigated. Fatty acids were detected in the solvent phase are
listed in Figure 5.6. Chains of fatty acids were extracted to the solvent
phase in T2 at 12 and 18 day, whereas in the T3 treatment few fatty acids
accumulated and only at 18 day. The oil extraction yields were low from
the algal milking experiments and further research is required to enable
this method to produce useful amounts of oil. Palmitic, stearic, oleic and
linoleic acids were the main fatty acids present in solvent phase in both
T2 and T3 treatments.
4.50 4.00 3.50 3.00 2.50 2.00 1.50 1.00 0.50 0.00
BBM BBM + Solvent BBM +25 mM NaCl + 20 μM phosphates + CU with solvent
Treatments 12 day 18 day Recovery after 18 days
Bio
mas
s con
cent
ratio
n(g/
L)
211
Chapter 5
The percentage of fatty acid (dry wt.) increased from day 12 to 18 in
medium with solvent (T2). A high percentage of linoleic acid (16.35 %)
was present in the T2 treatment. This percentage was higher than that
found in algal cells at 18 day in T2 and T1. There was a lag in the milking
of fraction of lipids into the organic phase during the first 12 day of
cultivation in the T3 treatment. In the T3 treatment only a small amount
of lipid extracted into the organic phase after 18 day. The percentage of
longer fatty acids in algal cells did increase (Figure 5.6) and this may be
due to changes in cultivation conditions with high light under continuous
illumination. The large percentage of fatty acid, palmitic and oleic acid in
the algal cells indicates that the use of salinity in the medium was effective
in stimulating lipid accumulation in the cells. The above result indicate
that preliminary tests on the milking of lipids from Scenedesmus bijugus
were successful because of the presence of viable cells even after solvent
exposure.
SEM imaging was performed to observe changes in the S.bijugus cell
morphology after solvent exposure. Figure 5.5 shows SEM images before
and after solvent exposure.
Chap
ter 5
Fi
gure
5.5
sho
ws
SEM
imag
es b
efor
e so
lven
t exp
osur
e (A
), af
ter
12 d
ay o
f so
lven
t exp
osur
e (B
) af
ter
12 d
ays
of s
alin
ity
with
sol
vent
exp
osur
e, (C
) bef
ore
solv
ent e
xpos
ure
at 1
8 da
y, (D
) bef
ore
solv
ent e
xpos
ure
at 1
8 da
y, (E
) afte
r 18
day
solv
ent
expo
sure
,(F) a
fter 1
8 da
y of
salin
ity w
ith so
lven
t exp
osur
e.
21
2
213
Chapter 5
The results in Figure 5.5 show that solvent did impact the shape of the
algal cells at 18 day in T2 and T3 compared to control conditions.
Scenedesmus bijugus was spherical with a rough surface before solvent
exposure. At 12 and 18 day, cells were indended. The indended
morphology was observed in T2 and T3. Despite the indended shape, the
cell membrane was intact, which is consistent with growth rates observed
in all the three treatments. The results of this study indicate that milking
of lipids the S.bijugus strain is possible and provide a proof of concept
toward using this method for the production of lipids from live cells.
Further work is needed to determine if yields can be improved to useful
levels and scale up issues will also need to be determined. S.bijugus
maintained viable cell count throughout 18 days of dodecane exposure
during which the lipids were extracted into the solvent phase.
S.bijugus was exposed to growth media for recovery. Biomass was
recovered at 18 day, however the total net biomass productivity was
less after recovery at 18 day (Figure 5.4) as compared to the previous
cycle. Cultures were able to grow in medium and contained a relatively
simple fatty acid profile. Palmitic, oleic and linoleic acid were extracted
during the recovery at 18 day. The percentage of these fatty acid
extracted are shown in Figure 5.6.
214
Chapter 5
Figure 5.6 Summary of fatty acids identified in solvent phase and algal
cells in long term milking experiments.
0
3
6
9
12
15
18
21
24
27
Fatt
y ac
id c
ompo
sitio
n (%
dry
wt.)
Variables
C14:0 C16:0 C16:1 C16:2 C16:3 C17:0
215
Chapter 5
Figure 5.7 Microalgae the bio-refinery concept. 5.4 Conclusion The results of this study demonstrate the feasibility of milking the
Scenedesmus bijugus strain for the production of lipids. S. bijugus
retained viable cells throughout 18 days of solvent (n-dodecane)
exposure and lipids were extracted into the solvent phase. Palmitic, oleic
and linoleic acid were the major fatty acids extracted into the solvent
phase. The medium with solvent produced high amount of linoleic acid in
the organic phase at 18 days, compared to control conditions without
solvent. The salinity stress conditions of the media resulted in an
increased percentage of oleic acid in the algal cells. Milking with n-
dodecane preferentially extracted few fatty acids in the organic phase
lipid compounds. Milking of Scenedesmus bijugus for biodiesel
production was demonstrated to be feasible. Further investigation need to
216
Chapter 5
be performed in detail.
The results obtained provide advancement in the milking of lipids
from algal cells. Higher cell biomass and higher net extraction efficiency
are major achievements in this study. Cell recovery confirmation and re-
synthesis of lipids and metabolites in recovered biomass indicates the
possibility of developing a continuous system for oil recovery, such
as the concept model shown in Figure 5.7.
217
Chapter 6
Chapter 6: Summary and future aspects
Biodiversity of microalgal isolates is vast and represents an untapped
resource with an array of biochemicals with potential for use in various
industries. The potential for coproduction of lipid and carotenoids, that
may be benefical to human health have gained interest in recent decades.
Diverse habitats provide stringent conditions which help shape the
organism with unique biochemical compositions. Bioprospecting of
microalgal resources from diverse ecologically distinctive locations and
better understanding of the physiological conditions of diverse habitats will
enable us to better exploit these organisms for the production of lipid and
carotenoids. Methods for co-production and separating higher value
compounds such as carotenoids and lipids can offset the cost of algal
biofuel production, making this source more commercially viable.
The main objective of this study was to examine the simultaneous
production of carotenoids and lipids from a collection of microalgae
from diverse habitats. For nutraceutical application isolates producing
lutein and astaxanthin are of interest. An aim of this research was to
isolate organisms with potential of producing lipids and carotenoids and
explore their ability to grow under different low cost stress conditions
or under a two stage cultivation process. Thus several changes in the
medium composition and their influence on biomass, lipids and
carotenoids production were studied. In addition, isolates in the
optimised growth conditions were further selected for harvesting
metabolites such as lipids and bioactives using algal milking. The
multiple milking cycles of lipids enables high recovery of lipids while
218
Chapter 6
maintaining microalgal cell viability. The ultimate aim was not only
improve the cost effectiveness of biodiesel production, but recovery of
multiple products as nutraceuticals, pharmaceutical and cosmetics
significance.
Chapter 2 describes the isolation, identification and preservation of new
strains of microalgae from different geographical region of India and
the World. Freshwater, backwater and terrestrial samples were collected
and screened. Twenty two strains out of two hundred were isolated from
diverse habitats across India and the World. The isolation of microalgae
isolates was carried out using dilution methdod. Each single cell colony
was designated by specific codes. All isolates were further purified on
nutrient agar plates to obtain axenic cultures. Scanning electron
microscopy indicated that the isolates belonged to the Chlorophyceae
and Scenedesmaceae families. The two hundred isolates were further
analysed using 18S rDNA analysis to confirm their identity using
universal primers. Twenty two isolates out of two hundred were found
to be unique, based on 18S rDNA sequencing using designed primers.
The purification, identification and preservation methods described in this
chapter were successful for obtaining and storing a range of diverse green
microalgae.
Chapter 3 describes the evaluation of the twenty two new isolates for
their ability to produce lipids with biofuel potential and the carotenoids
lutein and astaxanthin. Scenedesmus bijugus (from ladakh) showed the
highest biomass productivity, and potential to produce multiple products,
of the twenty two isolates studied. ATR-FTIR microscopy was
219
Chapter 6
performed with twenty isolates to determine the biochemical
compositions of algal cells. Unsaturated fatty acid peaks were present in
spectrum of all isolates, however peak intensity varied among the
isolates. Lipid localisation studies were performed using confocal
microscopy and showed the presence of lipids within cells and cell
membrane of all twenty two isolates. Due to thick cell walls in some of
the isolates, preventing cell penetration, the dye was not effective in
detecting the presence of lipid. GC analysis showed the presence of
comparatively high lipid levels in Scenedesmus sp. (P58), S. bijugus
(from Ladakh), Coelastrella sp. (P63), and Chlorella sorokiniana
(USA). Microalgal isolates from highly chemically polluted as well as
backwater habitats demonstrated the potential to produce large quantities
of lipids and biomass. The major fatty acids present in each of the
twenty two isolates were palmitic (C16:0), stearic (C18:0), oleic
(C18:1), linoleic (C18:2), and linolenic (C18:3), composing up to 90%
of total fatty acid content. Four isolates showed good biofuel
characteristics, based on HPLC and GC analysis. These were S. bijugus,
Chlorella sp., Auxenochlorella protothecoides and Coelastrella sp.,
which were determined to be the best producer for simultaneous
production of lipids and the carotenoids lutein and astaxanthin. On
the basis of desired biofuel properties, high lipid and carotenoids
productivity, the 4 best isolates were selected for further studies with the
aim of to further enhancing their lipid and carotenoid productivity. The
data obtained from from this study provide important inputs for further
optimization in culture medium to employ suitable stress in all four isolates
in order to obtain higher carotenoids and lipid productivity.
220
Chapter 6
In chapter 4, the four isolates selected from the bioprospecting
experiments were grown under a range of conditions to enhance lipid
and carotenoid production .We investigated the ability of S.bijugus
(from ladakh) to grow on medium containing different combination of
nitrate, salinity stress and cow urine and determined the impact of
these conditions on the production of lipids and carotenoids in a single
growth cycle. Salinity stress improved the fatty acid profile and
carotenoids production in these species. Nitrogen deficient medium
with salinity stress and the addition of cow urine as a low cost
nitrogen source increased lipids and biomass productivity. Higher
biomass was achieved in the medium containing cow urine as the nitrogen
source. Palmitic, stearic, oleic, linoleic and linolenic acid were the
most abundant fatty acids found for all treatments. Oleic acid levels
varied the most with changing media. Chlorella sp. was grown in a two
stage system. The strain showed improved biomass growth, lipid and
carotenoid production in medium rich in nitrates amended with salinity
stress, compared to that of control condition.
This treatment increased the linoleic (C18:2) and alpha-linolenic acid
(C18:3) levels. This strain grew well on low cost nutrient sources and
indicated potential for cost effective large scale production of
metabolites. The Coelastrella sp. was grown in two different medium in a
two stage system by altering light intensity and photoperiods. Both
medium (12:12h) with low light to (24:0h) or high light resulted in
increased biomass growth, lipids and carotenoids production, and a fatty
acid profile with biofuel characteristics. Cells grown from high (24:0h)
to low light (12:12h) in a single run produced lower levels of biomass,
221
Chapter 6
lipids and carotenoids. Auxenochlorella protothecoides when co
cultivated with AMF spore extract produced high net lipid and biomass
productivity with desired biofuel properties. Moreover, cow urine and
sodium nitrate could be used as potential nitrogen source for enhanced lipid
yield. On other hand, salinity, nitrogen defeceint conditions can promote
the enhanced carotenoids yield. Hence, cow urine and salinity together
demonstrated itself as a low cost culture medium for cultivation of four
isolates.
In this studyinvestigation was done on the abilities of a diverse collection
of isolates from different habitats to produce lipids and carotenoids
simultaneously in the same biomass. This strategy addressed a cost barrier
for biofuel production by investigating the production of valuable co-
products from these strains. The present study also showed that the use
of stress conditions and low cost media can enhance the production of
metabolites, while also improving cost economics.
Simultaneous production of lipids and carotenoids was carried out with
four novel isolates selected in this work. Previous studies where two
stage fermentation or stress factors were employed did not yield
biomass enhancement or measure metabolites simultaneously. Our study
indicates that a biorefinery approach from naturally occurring microalgal
isolates could be potentially feasible. The market for metabolites such as
lutein, astaxanthin, omega 3 fatty acids is relatively large compared with
other bioactive ingredients and therefore these are good target
compounds to offset the cost of biofuel production in microalgae.
222
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