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Downstream processing of lipids and
lipases from thraustochytrids
by
Avinesh Reddy Byreddy (M.Sc., M. Phil)
Submitted in fulfilment of the requirements for the degree of
Doctor of Philosophy
Deakin University
September, 2015
Acknowledgements
I would like to express my immense gratitude to Principal supervisor Professor Colin
James Barrow for providing me with this exciting opportunity to undertake the most
rewarding and challenging research under your supervision. Thank you very much for
the belief you had in me and the encouragement you have given me over the last few
years will stay with me.
I am grateful to my Principal co-supervisor, Associate Professor Munish Puri for
providing direction, designing experiments, valuable discussions surrounding
achieved results, and getting publications from the carried research work. I also thank
him for his scientific advices, his constructive feedback on my writing and
encouragement continually given over the last few years.
I would also like to thank Dr Madhusudan Rao Nalam (Senior Scientist, Centre for
Cellular and Molecular Biology, Hyderabad, India) for his insightful advices in
phospholipase research.
I would like to acknowledge an award of Deakin university postgraduate scholarship
(DUPRS), Deakin University, Australia for supporting my PhD research.
The journey made in the last 4 years would not have been possible without the support
of many kind hearted and good witted people around me. To my dear, “Super chooks”,
for your care and emotional support in all the good and bad times. I wish to express
my thanks to my fellow students and other postdocs from Barrow group. I am thankful
to the Technical staff at LES and Centre for Chemistry and Biotechnology for their
support and help.
Last but not least, I am extremely grateful to my dear wife Suchitra and my parents
Rama Mohan Reddy and Jayalakshmi and lovable sister Sowjanya for continues
support and encouragement throughout this study.
i
List of publications
1. Avinesh R. Byreddy, Colin Barrow, Munish Puri (2016), Bead milling for
lipid recovery from thraustochytrid cells and selective hydrolysis of
Schizochytrium DT3 oil using lipase. Bioresource Technology. 200, 464-469.
2. Avinesh R. Byreddy, Adarsha Gupta, Colin Barrow, Munish Puri (2015),
Comparison of cell disruption methods for improving lipid extraction from
thraustochytrid strains. Marine Drugs, 13, 5111-5127
3. Adarsha Gupta, Dilip Singh, Avinesh R. Byreddy, Tamilsevi Thyagrajan,
Shailedra Sonkar, Anshu S Mathur, Deepak K Tuli, Colin J Barrow, Munish
Puri (2015), Comparison of Australian and Indian thraustochytrids for omega-
3 fatty acids, enzymes and biodiesel production, Biotechnology Journal.
Accepted, 14th Oct.
Manuscripts Communicated/prepared
1. Avinesh R. Byreddy, Adarsha Gupta, Colin Barrow, Munish Puri (2015), A
quick colorimetric method for total lipid quantification in thraustochytrids.
New Biotechnology Journal.
2. Tushar R M, Avinesh R. Byreddy, Colin Barrow, NM Rao (2015),
Application of phospholipase A1 for concentration of polyunsaturated fatty
acids from fish oil.
Conference Proceedings
1. Avinesh Byreddy, Colin J Barrow and Munish Puri, Evaluating suitability of
bead milling on lipid recovery from marine microalgae for omega- 3 fatty acid
concentration. AAOCS meeting: 9-11 September 2015.
2. Avinesh Byreddy, Munish Puri, NM Rao, Colin Barrow, Application of
phospholipases for selective concentration of omega-3 fatty acids from fish oil.
Presented in 4th international conference on Novel Enzymes Organized by
Ghent University, Belgium 2014.
3. Munish Puri, Adarsha Gupta, Tamilselvi Thyagrajan, Avinesh Byreddy, Colin
J Barrow, Optimising lipid yields in microalgae screened from Australian
ii
marine environment for producing next generation biofuels. SMBB
conference, Mexico 2014
4. Avinesh Byreddy, Munish Puri, Colin Barrow, Production and extraction of
lipases from the Schizochytrium sp. S31. AISRF conference, March 12-14,
2014
5. Avinesh Byreddy, Munish Puri, Colin Barrow, Screening and comparison of
cell disruption methods for the extraction of lipases from marine microbes.
Organised by IFM, Deakin University, Nov 2012.
iii
Sequences deposited in GenBank database
1. DT1 Ulkenia sp. KF682136
2. DT2 Ulkenia sp. KF682137
iv
Table of contents
Acknowledgements ......................................................................................................i
List of publications .....................................................................................................ii
Sequences deposited in GenBank database.............................................................iv
Table of contents.........................................................................................................v
List of figures .............................................................................................................ix
List of tables ..............................................................................................................xii
List of abbreviations................................................................................................xiv
Abstract ....................................................................................................................xviChapter 1: Introduction and Literature review ......................................................1
1.1. Introduction....................................................................................................2
1.2. Omega-3 FAs.....................................................................................................3
1.2.1. Health benefits of omega-3 FAs .................................................................4
1.2.2. Dietary sources of omega-3 FAs.................................................................5
1.2.3. Techniques used for the concentration of omega-3 FAs.............................7
1.2.4. Microalgae as an alternative source for omega-3 fatty acids......................9
1.3. Thraustochytrids ..............................................................................................10
1.3.1. Potential of Thraustochytrids ....................................................................12
1.3.2. Strain selection and sustainable lipid production......................................12
1.3.3. Estimation of lipid content and fatty acid profile .....................................13
1.4. Extraction of lipids from dry cell biomass ......................................................15
1.4.1. Selection of suitable extraction solvents ...................................................15
1.4.2. Lipid extraction from wet biomass ...........................................................19
1.4.3. Microalgae cell disruption for lipid extraction .........................................20
1.4.3. Effect of cell disruption methods on lipid extraction................................21
1.5. Lipases .............................................................................................................25
1.5.1. Marine lipases ...........................................................................................25
1.5.2. Isolation of lipase producing microbes and screening methods ...............26
1.5.3. Extraction of intracellular lipases .............................................................28
1.5.2. Lipase based methods for concentration of omega-3 fatty acids ..............28
1.6. Research aim and objectives............................................................................33
v
Chapter 2: A quick colorimetric method for total lipid quantification in thraustochytrids........................................................................................................34
2.1. Introduction..................................................................................................35
2.2. Materials and Methods ................................................................................36
2.2.1. Chemicals used .........................................................................................36
2.2.2. Cultivation of thraustochytrids..................................................................37
2.2.3. Standard curve preparation .......................................................................37
2.2.4. Quantification of thraustochytrid lipids by SPV.......................................37
2.2.5. Lipid extraction .........................................................................................38
2.3. Results and discussion .....................................................................................39
2.3.1. Biomass and lipid production ...................................................................39
2.3.2 Microalgae oil quantification by SPV method...........................................40
2.3.3. Lipid quantification in thraustochytrid whole cells via SPV method .......42
2.3.4. Validating the accuracy of the SPV method .............................................44
2.4. Conclusion .......................................................................................................46
Chapter 3: Comparison of cell disruption methods for improving lipid extraction from thraustochytrid strains.................................................................47
3.1. Introduction..................................................................................................48
3.2. Experimental Section.......................................................................................49
3.2.1. Chemicals..................................................................................................49
3.2.2. Strain selection and biomass production...................................................50
3.2.3. Lipid extraction from thraustochytrids by organic solvents .....................50
3.2.4. Cell disruption for lipid extraction............................................................51
3.5. Fatty acids methyl esters (FAMEs) analysis ...................................................53
3.3. Results and Discussion ....................................................................................53
3.3.1. Biomass and lipid production ...................................................................53
3.3.2. Lipid extraction from thraustochytrid by organic solvents .......................54
3.3.3. Comparison of lipid extraction methods...................................................56
3.3.4. Fatty acid composition of extracted lipid..................................................60
3.3.5. Prediction of biodiesel properties .............................................................63
3.3.6. Energy analysis .........................................................................................65
3.4. Conclusion .......................................................................................................66
Chapter 4: Optimised cell disruption methods for lipase extraction from Australian thraustochytrids ....................................................................................67
4.1. Introduction......................................................................................................68
4.2. Materials and methods.....................................................................................69
4.2.1. Chemicals..................................................................................................69vi
4.2.2. Strain selection and cultivation .................................................................69
4.2.3. Enzymatic profiling of isolates .................................................................70
4.2.4. Lipase production......................................................................................70
4.2.5. Cell disruption methods for lipase release ................................................70
4.2.6. Microscopic observation of the native and disrupted thraustochytrid cells.............................................................................................................................71
4.2.7. Lipase and protein assay ...........................................................................71
4.3. Results and discussion .....................................................................................72
4.3.1. Cell disruption for lipase extraction..........................................................76
4.3.2. Comparison of different cell disruption methods for lipase extraction ....82
4.4. Conclusion .......................................................................................................83
Chapter 5: Tween 80 influences the production of intracellular lipase by Schizochytrium S31 in a stirred tank reactor .........................................................84
5.1. Introduction......................................................................................................85
5.2. Materials and methods.....................................................................................86
5.2.1. Microorganism and maintenance of culture..............................................86
5.2.2. Schizochytrium S31 growth and lipase production ...................................86
5.2.3. Estimation of Schizochytrium S31 growth................................................87
5.2.4. Optimisation of cell disruption through sonication...................................87
5.2.5. Lipase assay and protein estimation..........................................................88
5.2.6. Lipid extraction .........................................................................................88
5.3. Results and discussion .....................................................................................88
5.3.1. Optimisation of culture conditions for lipase production .........................88
5.3.2. Production of lipase in stirred tank reactor ...............................................90
5.3.3. Disruption of Schizochytrium S31 using sonication .................................92
5.3.4. Hydrolysis of p-Nitrophenyl esters ...........................................................95
5.4. Conclusion .......................................................................................................96
Chapter 6: Bead milling for lipid recovery from thraustochytrids cells.............97
6.1. Introduction......................................................................................................98
6.2. Materials and Methods ....................................................................................99
6.2.1. Chemicals..................................................................................................99
6.2.2. Thraustochytrids biomass cultivation .....................................................100
6.2.3. Cell disruption by bead milling...............................................................100
6.2.4. Microscopic observation .........................................................................101
6.2.5. Lipid extraction .......................................................................................101
6.2.6. Lipase catalysed hydrolysis of thraustochytrid oil..................................101
vii
6.2.7. Thin Layer Chromatography for lipid class analysis ..............................102
6.2.8. Gas Chromatography analysis for fatty acid composition ......................102
6.3. Results and discussion ...................................................................................102
6.3.1. Thraustochytrid cultivation .....................................................................102
6.3.2. Cell disruption by bead mill....................................................................103
6.3.3. Lipid profile analysis ..............................................................................107
6.3.4. Selective hydrolysis of Schizochytrium DT3 oil using Candida rugosalipase .................................................................................................................108
6.4. Conclusion .....................................................................................................111
Chapter 7: Phospholipase A1 as a catalyst to enrich omega-3 fatty acids in anchovy and algal oil ..............................................................................................113
7.1. Introduction....................................................................................................114
7.2. Material and Methods ....................................................................................116
7.2.1. Chemicals................................................................................................116
7.2.2. Hydrolysis of anchovy oil .......................................................................116
7.2.3. Thraustochytrid oil hydrolysis by phospholipase A1 .............................117
7.2.4. IATROSCAN analysis ............................................................................117
7.2.5. Methylation and GC analysis ..................................................................117
7.2.6. Analysis of positional distribution of fatty acid by NMR.......................118
7.3. Results and discussion ...................................................................................118
7.3.1. Lipase activity of Phospholipase A1.......................................................118
7.3.2. Hydrolysis of Anchovy oil by PLA1 ......................................................119
7.3.3. 13C NMR studies of the anchovy oil hydrolysis by PLA1 and BSL.......123
7.3.4. Thraustochytrid oil hydrolysis by phospholipase A1 .............................128
7.4. Conclusion .....................................................................................................130
Chapter 8. Summary and future aspects..............................................................131
viii
List of figures
Chapter 1:
Figure. 1.1 Chemical structures of omega-3 FAs, ALA; C18:3n-3 (cis-9,cis-12-cis-
15-octadecatrienoic acid, EPA; C20:5n- -eicosapentaenoic acid)and
DHA; C22:6n- -docosahexaenoic acid).........................................3
Figure. 1.2. Microscopic images of thraustochytrid cells; (A) Schizochytrium S31,
(B) Schizochytrium DT3 and (C) Thraustochytrium AMCQS5-5. ............................11
Figure. 1.3. Classification of cell disruption methods................................................21
Chapter 2:
Figure. 2.1. Schematic diagram of SPV method. .......................................................38
Figure. 2.2. Standard curve representing relationship between Schizochytrium oil
concentration and increase in the absorbance. ...........................................................40
Figure. 2.3. Comparison of various lipids and other compounds with sulfo-phospho-
vanillin reaction. .........................................................................................................41
Figure. 2.4. Profile of colorimetric detection of thraustochytrid whole cell lipids. ...43
Figure. 2.5. Change of lipid colour after reaction with sulfo-phospho-vanillin
reagent. .......................................................................................................................44
Chapter 3:
Figure. 3.1. Lipid extraction percentage from dry biomass using various solvents
from Schizochytrium sp. S31......................................................................................56
Figure. 3.2. Effect of different cell disruption methods for lipid extraction from
thraustochytrids. .........................................................................................................57
Figure. 3.3. Effect of cell disruption on thraustochytrids cells. Thraustochytrid cells
before (A) and after (B) cell disruption (scale bar 20 μm).........................................58
Figure. 3.4a. Effect of different cell disruption methods on fatty acid profiles of
Schizochytrium sp. S31...............................................................................................61
Figure. 3.4b. Effect different cell disruption methods on SFA, MUFAs and PUFAs of
Schizochytrium sp. S31...............................................................................................62
Figure. 3.5a. Effect of different cell disruption methods on fatty acid profiles of
Thraustochytrium sp. AMCQS5-5. ............................................................................62
Figure. 3.5b. Effect of different cell disruption methods on SFA, MUFAs and PUFAs
of Thraustochytrium sp. AMCQS5-5.........................................................................63 ix
Chapter 4:
Figure. 4.1. Cluster analysis of thraustochytrids based on various enzyme activities.
....................................................................................................................................75
Figure. 4.2. Microscopic observation of thraustochytrid after cell disruption. ..........77
Figure. 4.3. Effect of bead vortexing on thraustochytrid cell disruption and lipase
activity. .......................................................................................................................78
Figure. 4.4. Effect of grinding on thraustochytrid cell disruption and lipase activity.
....................................................................................................................................79
Figure. 4.5. Effect of sonication on thraustochytrid cell disruption and lipase activity.
....................................................................................................................................81
Figure. 4.6. Effect of homogenisation on thraustochytrid cell disruption and lipase
activity. .......................................................................................................................82
Chapter 5:
Figure. 5.1. Effect of Tween 80 concentration on lipase production by
Schizochytrium S31 at shake flask, pH 6.0 and incubation temperature 25 oC..........89
Figure. 5.2. Fermentation profile for the production of lipase by Schizochytrium S31
in a stirred tank reactor……………………………………………………………...92
Figure. 5.3. Biomass, lipid and DHA production reached a maximum after five days
of incubation...............................................................................................................91
Figure. 5.4. Fatty acid composition of Schizochytrium S31.......................................92
Figure. 5.5. Cell disruption optimisation of Schizochytrium S31 a) as a function of
sonication power, b) time duration.............................................................................95
Figure. 5.6. Substrate specificity of lipase from Schizochytrium S31........................96
Chapter 6:
Figure. 6.1. Schematic diagram of experimental process followed from
thraustochytrid cell disruption to enzymatic omega-3 fatty acids concentration. ....104
Figure. 6.2. Thraustochytrid cell disruption (40 X magnification) employing bead
mill............................................................................................................................104
Figure. 6.3. Optimisation of lipid extraction by bead mill as function of biomass
concentration.. ..........................................................................................................105
x
Figure. 6.4. Effect of bead mill agitator speed (rpm) on lipid yield from
thraustochytrid..........................................................................................................106
Figure. 6.5. Fatty acid composition of lipid extracted from thraustochytrids S31 and
DT3...........................................................................................................................108
Figure. 6.6. Capillary chromatography (iatrascan) profile of thraustochytrid oil ....109
Figure. 6.7. Optimisation of enzyme concetrtation. .................................................110
Figure. 6.8. Lipid class analysis of thraustochytrid oil after hydrolysis with Candida
rugosa lipase at different time intervals. ..................................................................110
Figure. 6.9. AG and FFA are separated hydrolysed oil and analysed by gas
chromatography........................................................................................................111
Chapter 7:
Figure.7.1 Phospholipase A1 activity towards p-NP esters with different acyl chains.
..................................................................................................................................119
Figure. 7.2. Time course of hydrolysis of anchovy fish oil by PLA1 and BSL A ...120
Figure. 7.3. Time course of hydrolysis of anchovy oil and fatty acid distribution in
the hydrolysate. ........................................................................................................122
Figure. 7.4. Percent hydrolysis of various fatty acids (from Fig.7.3) at 32%
hydrolysis of anchovy oil by PLA1 (light) and BSL (dark).....................................124
Figure. 7.5. 13C NMR spectra of the unhydrolysed oil fraction before (blue) and
after (red) 30% hydrolysis of anchovy oil by PLA1 (A) and BSL (B). ...................125
Figure. 7.6. Percent accumulation of different fatty acid classes in TG fraction after
30% hydrolysis of anchovy oil by PLA1 (dark) and BSL (light). ...........................126
Figure. 7.7. Lipase hydrolysis of structured triglyceride1, 3-dioeoyl-2-myristic
triglyceride................................................................................................................127
Figure. 7.8. Lipid class analysis of thraustochytrid oil after hydrolysis with
phospholipase at different time intervals..................................................................129
Figure. 7.9. Fatty acid profiles of hydrolysed and unhydrolysed DT3 oil. ..............130
xi
List of tables
Chapter 1:
Table 1.1. Contents of EPA and DHA in some fish species. .......................................6
Table 1.2. Dietary recommendation of omega-3 FAs by various scientific bodies. ....7
Table 1.3. Recent studies investigating the use of organic solvent in the extraction of
microalgal lipids. ........................................................................................................16
Table 1.4. Extraction of lipids improved by cell disruption methods. .......................24
Table 1.5. Main patents relating production of omega-3 fatty acids processing using
lipases from the last 10 years. ....................................................................................29
Chapter 2:
Table 2.1. Biomass and lipid production in glycerol medium by thraustochytrids....39
Table 2.2. Comparison of thraustochytrid lipid content by SPV, gravimetric and GC
methods. .....................................................................................................................45
Chapter 3:
Table 3.1. Biomass, lipid contents and productivity of thraustochytrids. ..................54
Table 3.2. Comparison of cell disruption methods employed to extract total lipids
from different microalgae...........................................................................................58
Table 3.3. Advantages and disadvantages of the investigated cell disruption methods.
....................................................................................................................................60
Table 3.4. Biodiesel properties of the given thraustochytrids strains. .......................64
Table 3.5. Energy consumption comparison. .............................................................66
Chapter 4:
Table 4.1. Lipase activity of thraustochytrids new isolates and ATCC cultures.
Lipase activity was determined by p-NPP assay using whole cell as catalyst. ..........76
Table 4.2. Disruption of thraustochytrid cells for lipase extraction. ..........................83
Chapter 5:
Table 5.1. Effect of sonication on protein release and lipase activity from
Schizochytrium S31. ...................................................................................................93
xii
Chapter 6:
Table 6.1. Energy consumption analysis for different mass concentrations and lipid
yields. .......................................................................................................................106
Chapter 7:
Table 7.1. Positional distribution of various fatty acids in anchovy oil. ..................123
xiii
List of abbreviations
°C - Degree Celsius
% - Percentage
- Micrometre
nm - nanometre
mg - Milligram
- Microgram
ml - Millilitre
- Micro litre
M - Micro mole
nM - Nano mole
L - Litre
g L-1 - Grams per litre
mg L-1 - Milligrams per litreL-1 - Micrograms per litre
Kg - Kilo gram
h - Hours
min - Minutes
sec - Seconds
- Alpha
- Gamma
- Omega
DHA - Docosahexaenoic acid
EPA - Eicosapentaenoic acid
AA - Arachidonic acid
OA - Oleic acid
GLA - Gamma-linolenic acid
DGLA - Di-homo gamma-linolenic acid
ALA - Alpha-linolenic acid
LA - Linoleic acid
SFAs - Saturated fatty acids
MUFAs - Monounsaturated fatty acids
PUFAs - Polyunsaturated fatty acids
FAs - Fatty acidsxiv
TFA - Total fatty acids
TAGs - Triacylglycerols
DAGs - Diacylglycerol
MAGs - Monoacylglycerol
PLA1 - Phospholipase A1
BSL - Bacillus subtilis lipase
DU - Degree of unsaturation
SV - Saponification value
IV - Iodine value
CN - Cetane number
LCDF - Long chain saturated factor
CFPP - Cold filter plugging point
CP - Cloud point
APE - Allylic position equivalents
BAPE - Bis-Allylic position equivalents
OS - Oxidation stability
HHV - Higher heating value
KV - Kinematic Viscosity
MJ - Mega joules
BHT - Butylated hydroxytoluene
GYP - Glucose, Yeast extract, Peptone
sp. - Species
GC - Gas Chromatography
FID - Flame Ionisation Detector
DCW - Dry cell weight
v/v - Volume by volume
w/v - Weight by volume
w/w - weight by weight
SD - Standard deviation
OD - Optical density
FAMEs - Fatty acid methyl esters
ATCC - American Type Culture Collection
xv
Abstract
Marine microalgae are a renewable alternative source to fish oil for the
sustainable production of omega-3 fatty acids. The potential use of marine microalgae
for nutraceutical and biofuel production is of global interest owing to increasing
concerns with sustainable production and environment impact. Thraustochytrid are
common marine heterotrophs, capable of producing large amounts of lipid,
particularly omega-3 fatty acids. These organisms are also potential sources of
bioactive proteins, enzymes, pigments and exopolysaccharides. The development
of rapid lipid quantification method for the isolation of new strains from diverse
marine habitats is important for future cost-effective production of these bioactive
metabolites. Cell disruption methods for improving lipid and lipase extraction has the
potential use in economising large scale algal processing for biofuel production.
Further, development of enzymatic methods for concentrating omega-3 fatty acids
from fish and algal oils is desirable because of their beneficial health effects.
Isolation and screening of suitable microalgae with high lipid productivity are
important for achieving sustainable commercial production of omega-3 oils and
biofuel. Current methods of lipid quantification are time consuming and costly.
Quantification of microalgal lipids using the sulpho-phospho-vanillin (SPV) reaction
is a rapid colorimetric method. This method was successfully tested on in-house
thraustochytrid strains. Conventional gravimetric analysis confirmed the accuracy of
the colorimetric method with R2 = 0.99. The total lipid contents of thraustochytrids
were determined gravimetrically was DT3 33.3%, S31 25.8%, AMCQS5-5 22.8% and
PRA 296 18.0%, respectively. Carbohydrates, proteins and glycerol did not interfere
with the SPV reaction.
Lipid extraction is an integral part of biodiesel production, as it facilitates the
release of fatty acids from algal cells. We evaluated the extraction efficiency of various
solvents and solvent combinations for lipid extraction from Schizochytrium sp. S31
and Thraustochytrium sp. AMCQS5-5. The maximum lipid extraction yield was 22%
using a chloroform:methanol ratio of 2:1. The highest lipid extraction yields were
obtained using osmotic shock method with 48.7% lipid yield from Schizochytrium sp.
S31 and 29.1% from Thraustochytrium sp. AMCQS5-5. Saturated and
monounsaturated fatty acid contents were more than 60% in Schizochytrium sp. S31
which suggests their suitability for biodiesel production.xvi
Marine microorganisms are a potential source of enzymes with structural
stability, high activity at low temperature and unique substrate selectivity. We
screened for a range of enzymatic activities from the seven thraustochytrid strains. The
relationship between isolate(s) and enzyme activity was investigated using cluster
analysis. The two strains AMCQ-4b27 (34 IU/g) and AMCQS1-9 (36 IU/g) exhibited
the highest lipase activity of the strains investigated. Four different cell disruption
methods, bead vortexing, grinding, sonication and homogenization, were evaluated for
their impact on lipase extraction yields. Sonication was found to be the best method
for enhancing lipase extraction yields from Thraustochytrium sp AMCQ-4b27 (0.903
IU/μg) and AMCQS1-9 (1.44 IU/μg).
The effect of Tween 80 as a carbon source was investigated with regard to
biomass, lipase and lipid productivity in Schizochytrium sp. S31. Tween 80 (1%) and
120 h of incubation was the optimum period for biomass, lipid and lipase productivity
in a stirred tank reactor. The yields obtained were 0.9 g L-1 of biomass, 300 mg g-1 of
lipid and 39 U g-1 of lipase activity. Sonication was optimised in terms of time and
acoustic power to maximise the yield of extracted lipase. The extracted lipase from
Schizochytrium S31 had unusual fatty acid selectivity in that it preferentially
hydrolysed DHA over EPA.
A method for lipid extraction from marine thraustochytrids using a bead mill
and enzymatic concentration of omega-3 fatty acids from the thraustochytrid oil was
investigated. The optimised lipid extraction conditions were, bead size 0.4-0.6 μm,
4500 rpm, 4 min of processing time at 5% biomass concentration. The maximum lipid
yield achieved at optimum conditions were 40.5% for Schizochytrium sp. S31 (ATCC)
and 49.4% for Schizochytrium sp. DT3 (in-house isolate). DT3 oil contained 39.8%
DHA as a percentage of lipid, a higher DHA percentage than S31. Partial hydrolysis
of DT3 oil using Candida rugosa lipase was performed to enrich omega-3
polyunsaturated fatty acids (PUFAs) in the glyceride portion. Total omega-3 fatty acid
content was increased to 88.7%.
For enrichment of omega-3 fatty acids in fish and algal oils, we have explored
the possibility of using phospholipase A1 (PLA1), a sn-1 position specific
phospholipid hydrolysing enzyme that also demonstrates lipase activity. PLA1 from
Thermomyces species was initially tested for its substrate specificity with various p-xvii
nitrophenyl esters. In this study, lipase from Bacillus susbtilis (BSL) was taken as a
control. The pH stat based hydrolysis followed by gas chromatographic
characterisation of the hydrolysate of Anchovy fish oil indicated a selective retention
of omega-3 fatty acids in the triglyceride fraction by PLA1, whereas such
discrimination was not observed with BSL. Selective accumulation of omega-3 fatty
acids was also confirmed by 13C NMR spectra. Position-wise analysis of fatty acids in
the unhydrolysed fractions suggests that PLA1 preferably retains docosahexaenoic
acid. Phospholipase A1 was used to catalyse the hydrolysis of thraustochytrid DT3 oil
to enrich DHA in glyceride portion, with short chain fatty acids being preferentially
cleaved from the triacylglyceride. This results show that phospholipase A1 can be used
to concentrate DHA from triacylglycerides.
xviii
Chapter 1
Chapter 1: Introduction and Literature review
1
Chapter 1 1.1. Introduction
Omega-3 fatty acids (FAs) are essential FAs, meaning that the human body
cannot synthesise them, so they must be obtained from the diet. Omega-3 FAs are well
known for their health benefits (Swanson et al., 2012). For example, high consumption
of omega-3 fatty acids in their diet decrease the risks of sudden death particularly in
those who already have had a heart attack (Blondeau, 2015; Hoen et al., 2013). DHA
is added to the infant formulas, as it is necessary for eye and brain development. These
fatty acids may also decrease the risks of as depression, schizophrenia, Alzheimer’s
and Parkinson’s diseases (Nichols et al., 2014; Tai et al., 2013). Multiple health
benefits associated with omega-3 FAs have increased the demand for these ingredients
in nutraceuticals and functional food industries. It has been forecast that the
consumption of omega-3 FAs in 2014 was 134.7 thousand metric tons valued at US$
2.5 billion. By 2020, it is been projected that the demand further will increase
significantly to 241 thousand metric tons with a value of US$ 4.96 billion (Market
watch, 2014). Most of the health benefits associated with omega-3 fatty acids are
attributed to the long-chain FAs eicosapentaenoic acid (EPA, C20:5-3) and
docosahexaenoic acid (DHA, C22:6-3). The most widely and naturally available
sources of EPA and DHA are fish oils. Concentration of EPA and DHA from fish oils
is normally achieved via chemical methods, which are not environmentally friendly,
also result in partial degradation of long chain FAs and production of toxic waste. This
has led researchers to seek an alternative, environmentally friendly method (Namal
Senanayake, 2013; Shahidi & Wanasundara, 1998). Enzymatic concentration of EPA
and DHA from fish oil using selective lipases is ecofriendly and causes less damage
to chemically sensitive long chain FAs (Shahidi & Wanasundara, 1998).
Production of EPA and DHA from fish oils has resulted in increased pressure
on fish species and number. Moreover, it has been recognised that mass fishing is not
a sustainable approach for EPA and DHA production (Lenihan-Geels et al., 2013). In
addition, these fish oils can be contaminated with heavy metal (methyl mercury) that
are hazardous to human health, particular to pregnant and lactating women’s (Lin et
al., 2014; Mahaffey et al., 2011). A viable alternative source of omega-3 FAs are
heterotrophic microalgae (Adarme-Vega et al., 2012). Commercial scale production
has been performed by using Crypthecodinium cohnii and Schizochytrium sp. as a
source for DHA (Infant formulas) and EPA (Food and beverages) (Kuratko et al.,
2
Chapter 1 2013; Kuratko & Salem, 2013). These algae can be grown in suitably sterile conditions
and oil extracted in a manner that is suitable for use in infant formula and dietary
supplements. Biomass harvesting and lipid extraction are expensive steps in the
production of oil from algal biomass (Kim et al., 2013). Effective cell disruption
methods and efficient solvent extraction methods are critical for high lipid yield and
cost effective production (Ryckebosch et al., 2014).
1.2. Omega-3 FAs
The term ‘Omega-3’ refers that the first double bond between the third and
fourth carbon atoms from methyl end, typically in cis configuration, hence called
omega-3 FAs. In cis configuration the hydrogen atoms are attached to double bonds at
the same side (Figure 1.1). Natural PUFAs consist of an even number of carbon atoms
with various unsaturation points. The most common omega-3 FAs are DHA C22:6,
EPA C20:5 and -linolenic acid (ALA) C18:3 (Poudyal et al., 2011; Ratnayake &
Galli, 2009). ALA is a precursor molecule in the biosynthesis of omega-3 FAs.
Figure. 1.1 Chemical structures of omega-3 FAs, ALA; C18:3n-3 (cis-9,cis-12-cis-15-
octadecatrienoic acid, EPA; C20:5n- -eicosapentaenoic acid)and
DHA; C22:6n-3 -docosahexaenoic acid). Modified from (Akanbi et
al., 2014b).
Generally omega-3 FAs are stored in adipose tissue in triacylglyceride (TAG)
form and present in all cellular membrane in phospholipid and related forms. Cell
membrane phospholipid composition determines the physiological characteristics of
the cell membrane, including changes in response to the external environment and
Alpha-linolenic acidALA (C18:3n-3)
Eicosapentaenoic acidEPA (C20:5n-3)
Docosahexaenoic acidDHA (C22:6n-3)
3
Chapter 1 functional activities of membrane-bound proteins (Murphy, 1990). The proportion of
EPA and DHA to all other FAs present in cell membranes differs and impacts the
function of cells, particularly storage and transport (Browning et al., 2012). DHA is
predominantly present in specific regions such as brain and eye. So for example, DHA
constitutes 18% of FA in adult human brain grey matter (Innis, 2007; Skinner et al.,
1993). With demonstrated physiological roles and established clinical benefits, the link
between human health and EPA and DHA is relatively well established.
1.2.1. Health benefits of omega-3 FAs
In the beginning of the 20th century dietary fats were recognised as a good
source of energy and fat soluble vitamins. This view later changed following various
studies, including a study conducted on rats reported that ‘dietary fatty acids were
required to prevent deficiency disease’ (Burr & Burr, 1929). The health benefits of
omega-3 FAs have been evaluated by a number of clinical studies. There is evidence
for the efficacy of omega-3 FAs in the prevention of sudden death from cardiovascular
disease and in ameliorating rheumatologic conditions (Barrow, 2010; Bhangle &
Kolasinski, 2011; Cicero et al., 2015). An early study on fat consumption and health
with Greenland Inuit indicated the cardiovascular benefits of omega-3 FAs. Further
studies produced more evidence for the cardiovascular benefits associated with
omega-3 FAs, including the reduction of the risk of arrhythmias (Asif, 2014),
decreasing platelet aggregation (Gao et al., 2013), lowering plasma triglycerides (Qi
et al., 2008), decreasing blood pressure (Cabo et al., 2012).
The consumption of dietary omega-3 FAs by ulcerative colitis and Crohn’s
disease patients has been shown to produce beneficial weight gain and significant
improvement in disease activity (Papadia et al., 2010). Omega-3 FAs have been
recently shown to have anti-cancer activity, particularly against colorectal cancer
(Cockbain et al., 2012). Many research investigations have been conducted to
understand the positive effects of omega-3 FAs on growth, learning and behavioural
problems of young children. Supplementation of omega-3 FAs during pregnancy
resulted in an increase in birth size (Makrides et al., 2011). Brain and eye development
in infants is particularly influenced by DHA, which has led to the addition of DHA to
infant formulae. Learning capacity of school going children is increased by taking
DHA enriched food supplements (Birch et al., 2007; Nunes et al., 2014; Richardson &
Montgomery, 2005). A number of studies have provided evidence for the anti-
4
Chapter 1 inflammatory effects of omega-3 FAs. Omega-3 FAs reduce the production of pro-
inflammatory molecules (eicosanoids) and increase the production of anti-
inflammatory molecules such as resolvins and protectins (Calder, 2012). The
combined effect of these multiple health benefits has been a dramatic increase in the
market demand for omega-3 FAs.
1.2.2. Dietary sources of omega-3 FAs
The main traditional sources of omega-3 FAs are marine fish. The marine
ecosystem is a rich source of omega-3 FAs, which are naturally produced by
microalgae and accumulated in fish and higher animal via the food chain (Lenihan-
Geels et al., 2013). Some other common sources include crustaceans, such as lobsters,
krill and molluscs, such as oysters and squid. Some land animals and plant oils also
contain omega-3 FAs in small quantities. Plant oils, such as canola, walnut oil, olive
oil and flaxseed oils are rich source of the omega-3 fatty acid ALA, a precursor for the
biosynthesis of EPA and DHA. To meet current requirements for men and women, the
recommended consumption of olive oil and flaxseed oil varies from 158–229 ml/d
(olive oil) to 2.2–3.2 ml/d (flaxseed oil) (USDA, 2006). The recommended intake of
omega-3 FAs from seeds and nuts to meet ALA recommended consumption for
women and men varies, from 12.2–17.6 g/d of English walnuts, 4.8–7.0 g/d of
flaxseeds, 612–890 g/d of pumpkin seeds (USDA, 2006). Many studies indicate that
the conversion of ALA to long chain FAs EPA and DHA is very low (Burdge &
Calder, 2005), because of this, diet rich in preformed EPA and DHA are required to
achieve the associated health benefits.
Marine derived fish oils are an excellent source for EPA and DHA. The
percentages of EPA and DHA in total fish oils vary depending on the fish species.
Farmed fish have more fat than wild fish, but the level of omega-3 in that fat varies
depending on fish feed use. Farm fish often has lower omega-3 levels than wild fish
even, although in one study farm-raised Atlantic salmon has 2.128 g SFA/85 g and
1.825 g EPA+DHA/85 g; whereas wild Atlantic salmon has 1.068 g SFA/85 g and
1.564 g EPA+DHA/85 g, respectively (Gebauer et al., 2006). Some of popular wild
caught fish and their EPA and DHA contents listed in table 1.1.
5
Chapter 1 Table 1.1. Contents of EPA and DHA in some fish species.
Common name Scientific name g/100g of food stuff portion
EPA DHA
Mackerel Scomber scombrus 1.10 2.56
Red mullet Mullus surmuletus 0.91 1.66
Sardine Sardina pilchardus 0.62 1.12
Salmon Salmo salar 0.50 1.00
Ton Thunnus thinnus 0.24 0.98
Fresh anchovy Engraulis
encrasicolus
0.14 0.80
Sea bream Pagellus bogaraveo 0.12 0.61
Cod Gadus morrhua 0.23 0.47
Hake Merluccius
merluccius
0.10 0.54
Conger eel Conger conger 0.15 0.43
Swordfish Luvarus imperialis 0.15 0.30
Dogfish Galeorhinus geleus 0.04 0
This table has been reproduced and modified from Rubio et al., (2010)
Dietary recommendation for omega-3 FAs advised by various scientific bodies
are given in table 1.1. The Dietary Guidelines for Americans 2005 report states,
‘‘Evidence suggests consuming approximately two servings of fish per week
(approximately 227 g; 8 ounces total) may reduce the risk of mortality from CHD
(Coronary heart disease) and that consuming EPA and DHA may reduce the risk of
mortality from cardiovascular diseases in people who have already experienced a
cardiac event’’. Another important challenge is meeting the recommended
consumption of omega-3 FAs in vegetarians and people who do not enjoy the taste of
fish. Alternative ways of getting omega-3 FAs for vegetarian is consuming fish oil
supplements and plant oils with significant omega-3 fatty acid content (Gebauer et al.,
2006; Maurer et al., 2012).
6
Chapter 1 Table 1.2. Dietary recommendation of omega-3 FAs by various scientific bodies.
S.
No
Professional body (year) Recommendation
1 UK Scientific Advisory
Committee on Nutrition (2004)
Minimum of 2 portions of fish/week that
is equivalent to 450 mg EPA + DHA
2 World Health Organization
(2003)
Up to two fish meals/week (400–1000 mg
EPA+DHA)
3 US National Academies of
Science, Institute of Medicine
(2002)
-linolenic acid/day (140 mg
EPA+DHA)
4 American Heart Association
(2002)
1g/day omega-3 FA for secondary
prevention of
CHD
This table has been reproduced from Yashodra et al., (2009)
1.2.3. Techniques used for the concentration of omega-3 FAs
Fish is a proven natural source of omega-3 FAs, but its consumption may not
meet recommended levels. A prior concentration and purification step is valuable,
therefore, to convert fish oils into a suitable chemical form which can be easily
metabolised in the human body, with the added benefit of improved oxidative stability.
Some studies suggest that the acylglyceride form of omega-3 FAs are better absorbed
than as ethyl esters and free fatty acid forms. In addition to that, some studies reported
that their oxidative stability has been increased in acylglyceride form (Dyerberg et al.,
2010). However, eating large amount of fish is also associated with the risk of heavy
metal – including mercury absorption, which may discourage people from eating fish.
(Lin et al., 2014). To achieve highly purified omega-3 fatty acids concentrate, physical,
chemical and enzymatic methods may be employed. The common methods employed
for the concentration and purification of omega-3 fatty acids from marine oils include
urea adduction, chromatographic methods, low-temperature fractional crystallization,
7
Chapter 1 supercritical fluid extraction, distillation, and enzymatic and integrated methods
(Namal Senanayake, 2013).
Urea adduction is simple and efficient technique for the concentration of
omega-3 fatty acids from marine fish oils. The principle behind this method is to
hydrolyse fish oil, using ethanolic KOH or NaOH, then mixing free FAs with urea
solution to form a complex. Saturated and monounsaturated fatty acids form complex
structures with urea and PUFAs include omega-3 FAs remain in the liquid portion,
which is highly enriched with omega-3 fatty acids. These two portions can be separated
by filtration and concentrated omega-3 FAs portion can be collected (Shahidi and
Senanayake, 2006). The major disadvantage of this method is that it produces large
amounts of urea waste, the disposal of which is expensive (Zuta et al., 2003).
Another important method for the concentration of omega-3 fatty acids from
fish oils is chromatographic techniques. This method involves saponification, followed
by treatment with urea solution. The non-complexed free fatty acids are converted into
fatty acid methyl esters and separated by chromatography (Namal Senanayake, 2013).
Guil-Guerrero and Belarbi used silver nitrate-impregnated silica gel column to purify
EPA and DHA from cod liver oil. Multiple washings with mobile phases led to the
recovery of DHA and EPA, 64% and 29.6%, with resulting purities of 100% and
90.6%, respectively (Guil-Guerrero & Belarbi, 2001).
Low-temperature fractional crystallisation is a simple method that depends on
differences in melting points of different fatty acids. The melting point depends on the
degree of unsaturation of each fatty acid. In general, the more saturate the fatty acids
the higher the melting point. The omega-3 FAs EPA and DHA melting points were -
54 and -44.5 oC, compared with 18:1 (Oleic acid) and 18:0 (Steric acid) of 13.4 and
69.6 oC, respectively (Namal Senanayake, 2013). As temperature decreases for a
mixture of saturated and unsaturated fatty acids, saturated fatty acids first crystalise
and the liquid portion enriched with unsaturated fatty acids (Shahidi and Senanayake,
2006).
Fractional distillation is a straight forward and widely accepted method for the
concentration of omega-3 FAs from a mixture under ultralow pressure. Based on their
boiling point and molecular weight, fatty acids are separated at reduced pressure and
elevated temperature of about 250 oC. This method allows highly selective separation
to concentrate selective fatty acids, but its high operating temperatures results in
8
Chapter 1 increased operating costs and damage to the oxidatively sensitive omega-3 FAs
(Breivik, 2007).
Large scale concentration of omega-3 FAs by physical or chemical methods is
associated with some drawbacks in terms of low yield, use of large volumes of
chemical, production of large volumes of waste effluents, high operation costs and
damage to the resulting fatty acids (Senanayake, 2000). Enzymatic methods such as
the use of lipase in concentrating omga-3 FAs is an alternative green method. Lipase
work under mild operating conditions, which make them suitable for the concentration
of omega-3 FAs from fish oils. Lipases have been used for many years in lipid
modification and food processing. Lipase based concentration involves either cleaving
short chain fatty acids from glyceride or selectively hydrolysing EPA and DHA from
glyceride. Thraustochytrid represent a group of marine microalgae and a source from
which to isolate lipases with novel activities. A novel lipase with selectivity towards
EPA and DHA would be useful and no such lipase has been characterised from any
source. Thraustochytrid are a potential source of omega-3 specific lipases. A detailed
review is given in (Section 1.3.6) on lipase mediated concentration of omega-3 fatty
acids.
1.2.4. Microalgae as an alternative source for omega-3 fatty acids
The current commercial source of omega-3 fatty acids for human consumption
is marine fish. Although most fish oil comes from anchovy, which is considered a
sustainable source, there are still concerns regarding overfishing, particularly for larger
fish such as tuna. There are also concerns with the presence of contaminants such as
mercury and polychlorinated biphenyls (PCBs), which are harmful to consumers
(Seabert et al., 2014). Furthermore, the fish oils are not suitable for vegetarians due to
their fish origins. New alternative sources must be found to supply the growing global
omega-3 market (Ratledge, 2013). Various alternative sources such as bacteria, fungi,
plants and microalgae have been explored for commercial production of single cell
oils (SCOs) (Adarme-Vega et al., 2012; Delarue & Guriec, 2014).
Marine microalgae are the primary producer of EPA and DHA in the marine
ecosystem. These naturally grow under a variety of culture conditions including
autotrophic, mixotrophic and heterotrophic (Buono et al., 2014). Use of microalgae as
a source of omega-3 FAs production has several advantages over traditional sources.
They can be grown in a controlled environment with readily available substrates in the 9
Chapter 1 fermentation process. They do not compete for land and fresh water and also eliminate
the risk of chemical pollutant contamination (Ryckebosch et al., 2012a). There are a
number of algal species with higher EPA and DHA contents. Thraustochytrids species
produce elevated quantities of omega-3 FAs with potential commercial application in
infant formulas, food, cosmetic and pharmaceutical products (Sijtsma & de Swaaf,
2004).
1.3. Thraustochytrids
Thraustochytrids are marine heterotrophic fungi like microorganisms, widely
distributed in a variety of habitats such as mangrove detritus, rocky shores, coral reefs,
salt marshes, sandy sediment, coastal waters, and deep sea (Liu et al., 2014a;
Raghukumar & Damare, 2011). The lipid content of thraustochytrids ranges from 50–
77% on a dry weight basis with more than 90% as triacylglycerides. Thraustochytrids
have recently emerged as a potential alternative source for the production of omega-3
fatty acids, particularly for DHA (Lee Chang et al., 2014). For example, Lee chang et
al, 2013, reported that the DHA percentage of total fatty acid content was more than
40% (w/w) in Aurantiochytrium sp. TC20 (Lee Chang et al., 2013). Another study
conducted by Ryu et al, (2013), observed that the DHA content of Aurantiochytrium
sp. KRS101 was 38 % (w/w) of total fatty acids (Ryu et al., 2013). These high DHA
levels indicate that thraustochytrids are potential sources for commercial DHA
production. Some of the important properties of thraustochytrids which could
influence successfully commercial scale production of omega-3 fatty acids include
high growth rates, a high proportion of DHA/EPA as percentage of total lipids and
unaffected growth at low salinity conditions (William et al., 2005).
Thraustochytrids are fungi like microorganisms, but they are classified under
the Labyrinthulomycetes, because ultrastructurally, they are closely related to
heterokont algae (Honda et al., 1999). This phylum contains 8 genera:
Thraustochytrium, Schizochytrium, Ulkenia, Labyrinthuloides, Labyrinthula,
Japonochytrium, Aplanochytriun and Althornia. Thraustochytrium and
Schizochytrium have a rapid growth rate and capable of producing large amount of
DHA (Azevedo & Corral, 1997; Wong et al., 2008). Microscopic image of
thraustochytrids used in this study are presented in Fig. 1.2.
10
Chapter 1
Figure. 1.2. Microscopic images of thraustochytrid cells; (A) Schizochytrium S31, (B)
Schizochytrium DT3 and (C) Thraustochytrium AMCQS5-5.
Many strains available today for commercial production of EPA and DHA are
the result of intensive procedures such as collection, isolation and screening. Extensive
screening and characterisation of thraustochytrid from a wide variety of samples led
to identification of thraustochytrids that can accumulate about 50% of total lipid, with
more than 25% of DHA (Raghukumar, 2008). Thraustochytrids are typically abundant
in marine sediments where large amounts of decaying organic matter accumulate,
resulting in enzymatic degradation (Lucia & Fernando, 2002). Many studies have
focused on the isolation and characterisation of thraustochytrids from various marine
habitats such as oceanic and coastal marine environments (Burja et al., 2006; Yang et
11
Chapter 1 al., 2010) as well as low temperature environments (Zhou et al., 2010) and the colonial
tunicate Botryllus schlosseri (Rabinowitz et al., 2006). There are two important
techniques used for the isolation of thraustochytrids from marine samples, direct
plating and pollen baiting. Direct plating techniques involve the spreading of samples
on agar plates under sterilised conditions (Bremer, 2000). Pollen baiting is the most
common method for isolation of thraustochytrids by which, sterilised pollen grains are
spread in marine samples with anti-bacterial and fungal agents and incubated for 1-2
weeks. Thraustochytrids cell growth can be observed on the surface of the pollen grain,
after which follows isolation of pure cultures on selective agar medium (Wilkens &
Maas, 2012).
1.3.1. Potential of Thraustochytrids
Currently the most common microalgae used for commercial production of
omega-3 fatty acids are marine derived, particularly from family members of
Thraustochytriaceae and Crypthecodiniaceae. Thraustochytrids are known to produce
several commercially interesting biotechnological compounds such as omega-3 fatty
acids, carotenoids, sterols, steroids and surfactants and also enzymes and
exopolysaccharides (Fan & Chen, 2007; Gupta et al., 2012a). The company DSM
commercially produces DHA rich oil Life’s DHATM using a Schizochytrium sp that
accumulate about 40-45% of DHA and 2% of EPA, for the food, beverage and
supplement industries. DSM uses a different strain of Schizochytrium, which produce
DHA and EPA rich oils in 2:1 ratio, to compete with anchovy fish oil that generally
has a higher EPA than DHA level. The trade name of this oil is Life’s OmegaTM, and
it contains 24% DHA and 12% EPA (Winwood, 2013).
Carotenoids are commonly used as a food colouring agent, particularly
astaxanthin that is used in aquaculture for salmon coloration. Carotenoids produced
either synthetically or derived from microbial production. Production of carotenoids
from microbial fermentation is an attractive area of research interest, as carotenoids
have been found to have protective activity against cancer, aging, ulcers, heart attack,
and coronary artery disease (Rao & Rao, 2007).
1.3.2. Strain selection and sustainable lipid production
Screening of high lipid producing microalgae is a key step for toward the
success of large scale cultivation and biodiesel production from microalgae. These 12
Chapter 1 microalgae are specific to particular environments and certain climate conditions
(Brennan & Owende, 2010; Williams & Laurens, 2010). These organisms occur in
various environments such as freshwater, lacustrine, brackish, marine, maturation
ponds and hyper-saline conditions (Mutanda et al., 2011). During the selection of
suitable microalgae for lipid production, organisms which have high biomass
productivity and lipid accumulation are preferred for mass cultivation than high lipid
storage with low productivity (Amaro et al., 2011; Mata et al., 2010). In addition, there
are other factors which have to be considered such as lipid profile and the capacity to
grow under specific conditions. Some microalgae produce fatty acids, making them
suitable for biodiesel production, whereas others, such as polyunsaturated fatty acids,
are not suitable because their unsaturation bonds are more prone to oxidative
degradation (Chisti, 2007). Biodiesel production depends on lipid profiles, which can
be regulated by the nutritional supply, processing and growth conditions (Amaro et al.,
2011). In some microalgal species, lipid accumulation is maximum during the
stationary phase. This can be induced by limiting one of the growth controlling
nutrients such as nitrogen, phosphorous and silicon (Guschina & Harwood, 2006).
Microalgae accumulate more lipid when subjected to stress conditions and by limiting
one of the growth nutrients, which enhances the production of lipid in microalgae
(Brennan & Owende, 2010; Pruvost et al., 2011).
In addition to bioprospecting for high lipid producing microalgae, finding a
cheaper raw materials for lipid production can reduce production cost. Researchers
have used molasses, crude glycerol and agricultural waste as a carbon source for lipid
production from microorganisms (Karatay & Dönmez, 2010).
1.3.3. Estimation of lipid content and fatty acid profile
Conventional lipid quantification methods involve microalgal biomass being
exposed to extraction solvents that extract lipids from the microalgal cellular matrix.
The whole mixture is centrifuged to separate the lipid layer from water and cell debris,
and the solvent evaporated. The microalgal crude oil thereby obtained is measured
gravimetrically and calculated as a percentage of dry biomass. This method only gives
the amount of lipid present in microalgae cells and not the lipid profile. To analyse the
lipid profile, obtained crude oil is subjected to transesterification process to convert
microalgal lipids to fatty acid methyl esters (FAMEs). The lipid composition and
quantities can be determined using gas chromatography (Bendikiene et al., 2011; Vyas
13
Chapter 1 et al., 2010). This method provides accurate quantity of lipid weight, but requires many
chemicals and multi-step process. In particular, this method is not favourable in
screening microalgae strains for high lipid production.
1.3.3.1. Rapid quantification of lipids
Microalgae strain selection depends on biomass and lipid productivity since
these are the key factors involved in economical biodiesel production. These factors
are critical to achieve, because the conditions favour high biomass may not favour
lipid accumulation, and vice versa (Huo et al., 2011). The biomass and lipid
production, particularly triacylglycerols, can be modulated by varying the culture
conditions such as nutrient limitation. In addition metabolic engineering can be applied
as a relatively new strategy for enhancing lipid accumulation in different microbial
species (Liang & Jiang, 2013). Therefore, a rapid and reliable lipid quantification
technique is required in identifying potential lipid producing strain from
environmental samples and screening large numbers of mutants. Traditional methods
for lipid quantification from microalgae are labour intensive, require solvents and
treatment steps (Wawrik & Harriman, 2010).
Staining of intracellular lipid using fluorescent lipophilic dyes, such as Nile red
for lipid quantification, popularly used for the dyes affinity towards neutral lipids (Han
et al., 2011; Rumin et al., 2015). Lipid content has been successfully quantified in
certain microalgae using Nile red dye, but unsuccessful with other microalgae because
of their thick cell wall that prevent the penetration of the dye. At elevated temperature,
Chen et al (2009) assessed the effectiveness of different solvents as a stain carrier and
found DMSO as good stain carrier for the species Chlorella vulgaris. Using this
optimised method, they successfully quantified lipids in microalgae using 96 well
plates (Chen et al., 2009). In a different study by Chen et al (2011), both conventional
and improved Nile red methods were ineffective in quantifying the lipid content in
green algae such as Pseudochlorococcum sp. and Scenedesmus dimorphus. This
problem was solved by treating the samples with microwaves for 50-60 seconds prior
to lipid quantification (Chen et al., 2011). Several studies have successfully quantified
the lipid content in various microalgae using Nile red methods (Bertozzini et al., 2011;
Gerde et al., 2012; Kou et al., 2013; Satpati & Pal, 2014).
Colorimetric quantification of lipids is achieved by sulfo-phospho-vanillin
methods (SPV). The SPV method was initially introduced by Chabrol and Charonnat14
Chapter 1 (1973) to quantify the total serum lipids in humans (B. Chabrol, 1937; Vatassery et al.,
1981). This method was further improved by Frings and Dunn (1970) and is still
widely used for serum lipid quantification.
1.4. Extraction of lipids from dry cell biomass
Marine microalgae are single-celled microorganisms with potential industrial
applications: as a source of aquaculture feedstock and production of valuable bioactive
compounds such as lipids, carotenoids and enzymes (Amin, 2009; Cao et al., 2013;
Mercer & Armenta, 2011). Lipid extraction is the most costly and critical step in
biodiesel production. Generally, organic solvent extraction method is most commonly
used as it is well established (Bligh & Dyer, 1959; Folch et al., 1957). However,
effective extraction of total lipids from microalgae is not possible due to the rigid cell
wall. In case of slow extraction due to the presence of strong cell wall, it is advised to
perform an extraction dynamic study in order to improve extraction efficiency (Cho et
al., 2012a). The most commonly used organic solvents are benzene, hexane, acetone
and chloroform and these have shown to be effective for lipid extraction from
microalgae by disrupting their cell walls (Ramluckan et al., 2014). Selection of the
solvent for lipid extraction should take into account its ability to enter into the algal
cells efficiently and match the polarity of the desired compounds (for instance, hexane
for non-polar lipids). Physical contact with solvent and lipid material can be achieved
by mechanically disrupting the microalgal cells and adding the solvent later (Li et al.,
2014b). The effect of cell disruption in lipid extraction from microalgae has been
reported and shown to significant improve lipid extraction yields (dos Santos et al.,
2015; Grimi et al., 2014; Halim et al., 2013; Lee et al., 2010b; Yap et al., 2014).
1.4.1. Selection of suitable extraction solvents
In order to optimise lipid extraction yields solvent selection is important.
Chloroform / methanol (1: 2 v/v) is the most commonly used and reliable organic
solvent mixture for total lipid extraction from microalgae (Bligh & Dyer, 1959). Using
this organic solvent system, residual endogenous water in the microalgal cells acts as
a ternary component that enables the complete extraction of both neutral and polar
lipids. This method does not require the complete drying of microalgal biomass. Once
the cell debris is removed, more chloroform and water are added to induce biphasic
partitioning. The lower organic phase (chloroform) contains most of the lipids (both
15
Chapter 1 neutral and polar) while the upper aqueous phase (water with methanol) constitutes
most of the non-lipids (proteins and carbohydrates) (Medina et al., 1998). The
chloroform and methanol lipid extraction process is fast and quantitative. However,
the use of chloroform associated with health, security, and regulatory problems, and
as such, its usage is undesirable. Folch et al, originally developed chloroform and
methanol lipid extraction procedure to extract lipids from brain tissue (Folch et al.,
1951). The lipid extraction yield depends on the solvent mixture used in the extraction
process. As a result, its efficiency in lipid extraction from microalgal biomass need to
be further investigated.
Jeon et al, examined 15 different solvents and analysed their efficiency in lipid
extraction from Chlorella vulgaris biomass. Lipid extraction yields increased to 25%
with methanol:dichloromethane as a solvent mixture compared with other solvents
used (Jeon et al., 2013). Mandal et al, also studied the effect of different solvents and
their combinations in lipid recovery from Scenedesmus obliquus. Around 13% of lipid
extracted using chloroform and methanol (2:1) solvent (Mandal et al., 2013).
Ramluckan et al, investigated the efficiency of 13 solvents and their combinations with
a range of polarities and solubilities. The greatest lipid extraction efficiency (11.76)
was obtained upon using 1:1 mixture of chloroform:ethanol (Ramluckan et al., 2014).
Table 1.3 summarises the recent studies investigating the effect of various solvents on
lipid extraction.
Table 1.3. Recent studies investigating the use of organic solvent in the extraction of
microalgal lipids.
16
Chapter 1
Marine
Microorganis
ms
Dried
biomas
s (g)
Organic solvents v/v Reaction conditions
Duration
(min)/agitation
(rpm)/Temp (oC)
Wt% of
total
extracted
lipids
Reference
Schizochytriu
m S31
0.05 Chloroform/ethanol (2:1) NA 22 (Byreddy et
al., 2015)
Chlorella sp. 1 Chloroform/ethanol (1:1) 3 h/NA/boiling point 11.76 (Ramlucka
n et al.,
2014)
Scenedesmus
obliquus
1 Chloroform/ethanol (2:1) NA/NA/room
temperature
12.9 (Mandal et
al., 2013)
Nannochloro
psis gaditana
Tetraselmis
suecica
Desmodesmu
s communis
0.05 N,N-
dimethylcyclohexylamine
50 mg/ml
(Biomass/solvent)
24 h/stirring/room
temperature
29.2
57.9
31.9
(Samori et
al., 2013)
Chlorella
vulgaris
50 dichloromethane:methano
l (1:1)
1 h/stirring/37 oC 25 (Jeon et al.,
2013)
Arthrospira
platensis
1 chloroform/methanol
(2:1)
NA 20 (Baunillo et
al., 2012)
Pavlova sp 10 Water/methanol/ethyl
acetate (10:24:48)
NS 44.7; (Cheng et
al., 2011a)
Chlorococcu
m sp
4 Hexane,
hexane/isopropanol (3:2)
450/800/25 6.8;
hexane/isop
ropanol
(Halim et
al., 2011)
Synechocystis
PCC 6803
15 Chloroform/methanol/
water ( 1:2:0.8)
24 h/stirring/room
temperature
4.2 (Sheng et
al., 2011b)
Schizochytriu
m limacinum
1 Water/chloroform/methan
ol (4:6:12)
NA 57 (Johnson &
Wen, 2009)
17
Chapter 1
Aurantiochyt
rium sp.
strain T66
28-32 Methanol/chloroform/wat
er (2:2:1)
NS/NS/40 55;
Methanol/c
hloroform/
water
(Jakobsen
et al., 2008)
Phaeodactylu
m
tricornutum
10 Ethanol 1440/500/25 6.3 (Fajardo et
al., 2007)
S. mangrovei
IAo-1
100-
200
Chloroform/methanol
(2:1)
NS/Ns/38 33.2 (Leano et
al., 2003)
Chlorella sp 0.1 Soxhelt: methylene
chloride:methanol (2:1)
Batch 1:
chloroform/methanol/50
mM phosphate buffer
(35:70:28)
Batch 2: n-hexane/IPA/
distilled water (70:47.7:3)
Soxhlet: 180/NS/NS
batch 1: 2160/NS/NS
batch
2: overnight/NS/NS
11.9;
Soxhlet
with
methylene
chloride/me
thanol (2:1)
(Guckert et
al., 1988)
Botryococcus
braunii
0.12 Chloroform/methanol
(2:1), hexane/isopropanol
(3:2),
dichloroethane/methanol
(1:1),
dichloroethane/ethanol
(1:1),
acetone/dichloromethane
(1:1)
50/high/NS 28.6;
chloroform/
methanol
(2:1)
(Lee et al.,
1998)
18
Chapter 1
1.4.2. Lipid extraction from wet biomass
Drying microalgae biomass prior to lipid extraction accounts for 90% of the
total production energy cost in dry algal lipid extraction for biodiesel production.
Microalgae biomass drying is an energy intensive process, more than 25% of energy
required for this process can be saved by using wet algal biomass (Du et al., 2015).
There are few studies on microalgae lipid extraction using wet biomass. Yao et al,
investigated the use isopropanol as a potential solvent in extracting lipids from
Nannochloropsis sp. wet biomass and achieved 70% of lipid extraction (Yao et al.,
2012). Dejoye, et al, developed a new environmentally safer method, called
simultaneous distillation and extraction process for lipid extraction from wet
microalgae Nannochloropsis oculata and Dunaliella salina. This method used
alternative solvents such as d- -pinene and p-cymene. The lipid recovery
yields were compared with standard soxhlet extraction and Bligh & Dyer method and
observed similar lipid yields (Dejoye Tanzi et al., 2013). Yang, et al, proposed the use
of ethanol as an organic solvent for lipid extraction from wet microalgae Picochlorum
sp. This study investigated the effect of extraction temperature and time and the
Isochrysis
galbana
5 Chloroform/methanol/wat
er (1:2:0.8)
hexane/ethanol (1:2.5),
hexane/ethanol (1:0.9),
butanol, ethanol,
EtOH/water (1:1)
60/constant/25 8.9;
chloroform/
methanol/w
ater
(1:2:0.8),
(Grima et
al., 1994)
Chaetoceros
muelleri
and
Monoraphidi
um
minutum
60 1-butanol, ethanol,
hexane/2-propanol
(2/3), water/
methanol/chloroform
as a control system
90/high (NS)/close to
boiling point of each
chemical
Control
100%
extraction.
2nd
Highest
efficiency
with 1-
butanol
at 94%
(Nagle &
Lemke,
1990)
19
Chapter 1 influence of solvent biomass ratios on extraction efficiency and lipid class, with results
indicating that the extracted lipid was comparable with Bligh & Dyer method (Yang
et al., 2014). Another recent study confirms the suitability and efficiency of ethanol as
an extraction solvent for lipid extraction from some microalgae (Yang et al., 2015).
Sathish, et al, developed and modified a lipid extraction procedure from wet
microalgae biomass using acid and base hydrolysis. This method was capable of
extracting 79% of total transesterified lipids from Chlorella and Scenedesmus sp.
(Sathish et al., 2015; Sathish & Sims, 2012).
Recently, oil extraction from wet biomass and direct biodiesel conversion
called direct (in-situ) transesterification, has been investigated (Park et al., 2015). Chen
et al, developed a novel method for microalgal biodiesel production using
Chlamydomonas sp. JSC4 with 68.7 wt% water content as the feedstock. This method
involves microwave disruption, partial dewatering (via combination of methanol
treatment and low-speed centrifugation), oil extraction, and transesterification without
removal of solvent and they achieved 97% biodiesel conversion (Chen et al., 2015a).
A recent study by Kim et al, investigated the effect of factors influencing the direct
biodiesel conversion from wet Nannochloropsis salina biomass. The reaction
conditions such as temperature, reaction time, and solvent and acid quantity were
optimised and 90% of total conversion was obtained (Kim et al., 2015).
1.4.3. Microalgae cell disruption for lipid extraction
A large number of lipid extraction procedures are used for microalgal biomass
and their extraction efficiencies have been documented. All microalgae are single cell
organisms having an individual cell wall. Furthermore, some species of microalgae
have high lipid levels but they are located inside a tough cell wall. The presence of
tough cell walls of microalgae increase the lipid extraction cost, since a cell disruption
step is required for effective lipid extraction. Cell disruption is an energy consuming
process but an important step in liberating intracellular lipid molecules from
microalgae (Lee et al., 2012). A number of cell disruption methods are available for
lipid extraction from microalgae. Different microalgae have different disruption
propensities and as a result, no single method of disruption can be universally applied
to the various microalgae species (Halim et al., 2012b).
Cell disruption methods (Figure. 1.3.) generally used at the laboratory scale are
classified according to the working procedure by which microalgal cells are disrupted, 20
Chapter 1 for example mechanical and non-mechanical (Chisti & Moo-Young, 1986; Günerken
et al., 2015; Middelberg, 1995). Mechanical methods include bead mill, press, high-
pressure homogenisation, ultrasonication, autoclave, lyophilisation, and microwave,
while non-mechanical methods often involve lysing the microalgal cells with acids,
alkalis, enzymes, or osmotic shocks (Chisti & Moo-Young, 1986; Günerken et al.,
2015).
Figure. 1.3. Classification of cell disruption methods. Modified from (Chisti & Moo-
Young, 1986).
1.4.3. Effect of cell disruption methods on lipid extraction
Microalgal cell wall disruption in a non-specific manner is generally achieved
by mechanical methods using solid-shear forces. The basic principle of bead milling
involves physically grinding the microalgal cells against the solid surfaces of glass
beads in a violent agitation (Chisti & Moo-Young, 1986). Some key factors include
high throughput process, high biomass loading, good temperature control, easy scale
up procedures, low labour intensity and high disruption efficiency, which in
combination make bead milling a suitable method of implementing large scale
applications (Günerken et al., 2015; Jahanshahi et al., 2002). The method has been
used for years to disrupt microorganisms. The size of beads used in the disruption
21
Chapter 1 process is important. The optimal diameter of the beads for bacterial cell disruption is
0.1 mm and 0.5 mm for yeast and other unicellular organisms. The cell disruption
efficiency can be further improved by using zirconia-silica, zirconium oxide or
titanium carbide made beads. This is because of their hardness and density. In addition,
the separation of beads from the agitated solution is easily achieved due to the high
density of beads (Lee et al., 2012c).
High pressure homogenisation is an effective and simple cell disruption
method. In this process microalgal suspension pumped through narrow orifice of a
valve under high pressure. Cell disruption is achieved by high-pressure impact (shear
forces) of the accelerated fluid jet on the stationary valve surface as well as
hydrodynamic cavitation from the pressure drop induced shear stress (Chisti & Moo-
Young, 1986). Some recent studies observed that high working pressure and cycle
number have a positive effects on cell disruption efficiency (Halim et al., 2012b). In
addition, microalgae biomass concentration and species have major effect on specific
energy consumption (Halim et al., 2013). High speed homogenization is a simple and
effective method by which a stirring device which can rotate at high rpm with in a
stator-rotor assembly. The stator-rotor are usually made of stainless steel with a variety
of designs. High speed homogenization achieves cell disruption via hydrodynamic
cavitation, generated by stirring at high rpm, and shear forces at the solid-liquid
interphase. Hydrodynamic cavitation occurs when the impeller tip reaches
approximately 8500 rpm due to the local pressure decreases nearly down to the vapour
pressure of the liquid (Gogate & Pandit, 2008; Kumar & Pandit, 1999).
Microalgal cells can be disrupted by an ultrasonication process that is
transmission of sonic waves. An ultrasonicator converts the electrical energy into
mechanical vibrations. The probe intensifies the mechanical vibrations, resulting in the
formation of pressure waves in the liquids. This mechanism forms significant amounts
of cavities (bubbles) in the liquid, which expand during the low pressure phase and
collapse violently during the high pressure phase. This phenomenon is referred as
“cavitation”, which releases pressure and heat. Although the cavitational collapse lasts
only for a few microseconds, the cumulative effect of bubbles is to release high levels
of energy into the liquid (Chisti & Moo-Young, 1986).
Generally the non-mechanical cell disruption methods involve use of
chemicals, enzymes and osmotic shock (Monks et al., 2013). These methods primarily
22
Chapter 1 depend on the interaction between chemicals or enzymes and cell walls of the
microalgae, which modify the outer membrane layer and release the intracellular
components. Enzymatic cell disruption is a desirable process due to biological
specificity, mild operating conditions, low energy requirements, low capital
investment. The use of enzymes normally circumvents the need to use aggressive
physical conditions such as high shear stress and elevated temperatures (Choi et al.,
2010; Harun & Danquah, 2011). The discovery of the enzyme lysozyme led interest
in the application of lytic enzymes in cell lysis. During enzymatic cell lysis, enzymes
bind to cell membranes at specific sites and hydrolyse specific bonds leading to
degradation of cell membrane (Choonia & Lele, 2013; Fu et al., 2010; McKenzie &
White Jr, 1991). A variety of chemical agents such as such as antibiotics, chelating
agents, chaotropes, detergents, solvents, hypochlorites, acids and alkali can cause cell
disruption. The efficiency of the cell disruption depends on the selectivity of the
chemical agent and composition of the microorganisms (Jalalirad, 2013; Middelberg,
1995).
Lee at al., (2010) assessed the effect of prior mechanical cell disruption on lipid
extraction for Botryococcus sp., Chlorella vulgaris, and Scenedesmus sp. using
chloroform and methanol (2:1 v/v) as an extraction solvent. Among the methods used
(autoclave, bead beating, microwave, sonication, and osmotic shocks), the microwave
method was resulted in the higher lipid extraction yields in all three microalgal species.
Another study by Zheng et al., (2011) investigated the effect of different mechanical
and non-mechanical cell disruption methods for release of lipids from a marine species
of Chlorella vulgaris. Overall, grinding in liquid nitrogen was identified as the most
effective method in terms of disruption efficiency.
A similar study was conducted by Halim et al, (2012), in which four cell
disruption methods were compared to determine their lipid extraction efficiency.
Among the four methods (Bead beating, ultrasonication, acid treatment and high-
pressure homogenisation), high-pressure homogenisation was the most effective in
disrupting Chlorococcum cells. Summary of recent studies on microalgal cell
disruption and their lipid extraction efficiencies are given in table. 1.4.
23
Chapter 1 Table 1.4. Extraction of lipids improved by cell disruption methods.
Microalgae Cell disruption
method
Maximum
lipid yield (%)
Reference
Schizochytrium
S31
Thraustochytrium
AMCQS5-5
Osmotic shock 48.7
29.1
(Byreddy et al.,
2015)
Chlorella vulgaris Ultrasonication 55 (dos Santos et al.,
2015)
Scenedesmus sp Freeze drying +
Microwave
29.6 (Guldhe et al.,
2014)
Chlorella vulgaris pressure-assisted
ozonation
27 (Huang et al.,
2014)
Mixed culture Microwaves 33.7 (Florentino de
Souza Silva et
al., 2014)
Chlorella vulgaris H2O2 + FeSO4 17.34 (Steriti et al.,
2014)
Nannochloropsis
sp
Microwave 38.3 (Wahidin et al.,
2014)
C. vulgaris Ultrasonication 52.5 (Araujo et al.,
2013)
Chlamydomonas
reinhardtii
Osmotic shock NA (Yoo et al.,
2012b)
Microcystis sp Sonication 53.89 (Supriya &
Ramachandra,
2012)
C. vulgaris (SAG
211-12)
Grinding+microwaves+
sonication
9.82 (Sostaric et al.,
2012)
Synechocystis
PCC 6803
Microwave+pulsed
electric fields
9-13 (Sheng et al.,
2012)
Chlorella vulgaris Microwaves NA (Prommuak et
al., 2012)
24
Chapter 1 Scenedesmus sp high-pressure
homogenization
24.9 (Cho et al.,
2012b)
Synechocystis
PCC 6803
pulsed electric field 25-75 (Sheng et al.,
2011a)
Chlorella sp
Nostoc sp
Tolypothrix sp
Sonication
Sonication
Microwave
21
18
16
(Prabakaran &
Ravindran,
2011)
Botryococcus sp
Chlorella vulgaris
Scenedesmus sp
Microwave
28.6
10
11
(Lee et al.,
2010b)
Scenedesmus
dimorphus
Chlorella
protothecoides
Wet milling
Bead-beater
25.3
18.8
(Shen et al.,
2009)
1.5. Lipases
Lipase have emerged as an important biocatalyst with potential applications
in many industries including organic synthesis, paper manufacturing, oleochemistry,
dairy, cosmetics, perfume, biosensors, and detergents (Hasan et al., 2006). Lipases
catalyse the breakdown of fats into free fatty acids and glycerol at the water oil
interface. Lipase also synthesise ester from free fatty acids and glycerol at the
hydrophobic interface. Lipases often exhibit high chemo-, region- and enantio-
selectivity at different conditions contributing to their utility as industrial biocatalysts
(Gotor-Fernández et al., 2006).
1.5.1. Marine lipases
The marine ecosystem and marine microorganisms in particular represents an
enormous pool of potential enzyme candidates for industrial and biotechnological
applications. Marine microbes are an important source of bioactive compounds (Soliev
et al., 2011). The demand for the novel enzymes is also increasing, due to their utility
in the production of chemically and structurally important molecules. Marine
microorganisms are a potential source for the isolation of enzymes with novel
properties, because of their stability and their potential for wider applications than
25
Chapter 1 those of enzymes isolated from plant or animal sources (Bull et al., 2000; Lam, 2006).
The potential demand for enzymes in industrial applications is increasing every year.
Marine environments provides unique genetic structures and life habitats such as
nutrient rich regions, nutrient limited regions, high salinity, high pressure, low
temperature and special light conditions. Microorganisms must adapt to survive in the
marine environment and so new biodiversity is created (Kennedy et al., 2008; Zhang
& Kim, 2010).
Lipases are found in all forms of life, includes bacteria, fungi, plants and
animals (Wang et al., 2008). Microorganisms are potent sources of enzymes by
comparison with plants and animals, because of the availability of their wide variety
of catalytic activities, the high yields possible, relative ease of genetic manipulation,
regular supply due to the absence of seasonal fluctuations and rapid growth of
microorganisms (Iftikhar et al., 2010). Microbial lipases are normally more stable than
the other lipase sources which makes them more suitable for industrial applications
(Peng et al., 2014). Fungi are the most studied source for the production of commercial
lipases. Some of the common lipase producing fungi belong to the following genus,
Rhizopus sp., Aspergillus sp., Penicillium sp., Geotrichum sp., Mucor sp., and
Rhizomucor sp. Candida antarctica and candida rugosa lipases are most extensively
studied fungal species (Gopinath et al., 2013; Singh & Mukhopadhyay, 2012). Lipase
production depends on the carbon source, nitrogen source, pH and temperature of the
growth medium (Cihangir & Sarikaya, 2004). Unusual lipase producing
microorganisms can be isolated from the extreme environments, such as sites
contaminated with oil, dairy waste, vegetable oil waste and industrial waste (Sztajer et
al., 1988).
1.5.2. Isolation of lipase producing microbes and screening methods
The increasing demand for lipases in industrial applications has led the
identification of novel lipases with unique properties. Screening for specific lipase
activity and new lipases is facilitated by the development of rapid, reliable, specific,
selective, and sensitive analytical methods for evaluating their catalytic activity
(Stoytcheva et al., 2012). Quantifying lipase activity can be challenging since lipases
are water soluble enzymes but act on water insoluble substrates (Kanchana et al.,
2011). Methods based on using chromogenic and fluorogenic substrate are amongst
the simplest and most widely used methods for screening for lipase activity (Hasan et 26
Chapter 1 al., 2009). A preliminary plate method is normally used in the initial screening for
lipase producing strains. The medium in the agar plates are supplemented with lipid
substrate (olive oil or tributyrin) and microorganisms are streaked on agar plate and
periodically observed (Singh et al., 2006). Microbes that are producing lipase can
make a halo in the plate and then these strains are selected for lipase production in the
liquid medium (Cardenas et al., 2001); Hasan et al., 2009).
Lipases hydrolyse ester bonds and liberate free fatty acids that can be
titrimetrically measured by neutralizing the released free fatty acids by manually
adding NaOH or by a pH stat method (Borkar et al., 2009). In spectrophotometric
methods, substrate are designed to give a colour compound upon enzyme hydrolysis.
Lipase activity can be measured in microbial culture supernatant by using p-
nitrophenyl esters of various chain lengths. The lipolytic activity on carboxylic ester
leads to the release of yellow coloured p-nitrophenol that can be monitored
quantitatively by spectrophotometry (Winkler & Stuckmann, 1979).
Spectrofluorometric methods are rapid, simple and sensitive. Substrates used in this
method are acylglycerol analogs coated with fluorophores such as dansyl, resorufin,
or 4-methylumbelliferone (MUF). These analogues are released by lipolytic activity,
which can be measured fluorometrically (Li et al., 2014a).
Chromatographic techniques can be used to measure lipase activity by
quantifying the released free fatty acids from triacylglycerols. Most general techniques
use thin-layer chromatography, gas-liquid chromatography, HPLC or gas
chromatography- mass spectrometry (Hao et al., 2007; Stoytcheva et al., 2012). Lipase
activity estimation by HPLC was developed by Maurich et al 1999, using beta-
naphthyl or para nitrophenol esters. Substrate were incubated with the enzymes and
released naphthol was quantified by reverse phase HPLC. Gas chromatography is used
to determine the products released by the enzymatic reaction (Stoytcheva et al., 2012).
Since lipases are inducible enzymes, the medium composition, particularly the carbon
source, influences lipase production. Most microbial lipases are extracellular lipases.
Lipase production can be induced by adding lipase substrates into the media, such as
triacylglycerols, fatty acids, hydrolysable esters, tweens, bile salts, and glycerol (Gupta
et al., 2004). In addition, the nitrogen source and some important micronutrients also
play a crucial role in the production of lipases. Lipase production levels can be
increased by the optimisation of culture condition, generally optimised by changing
27
Chapter 1 one variable at a time while keeping other variables constant. However, this systematic
approach is time consuming and doesn’t effectively explore the interaction between
variables. A more versatile approach is the Plackett-Burman (PB) method, which is an
efficient and widely accepted approach for the optimisation of multiple culture
variables (Mazzucotelli et al., 2015).
1.5.3. Extraction of intracellular lipases
Enzymes that are synthesized and remained inside the cells are called
intracellular enzymes. To extract intracellular enzymes, the cells have to be broken by
mechanical or non-mechanical methods to release the products. Some cell disruption
methods are sonication, homogenisation, bead beating, French press, or chemical and
enzymatic methods (Safi et al., 2014b). Jermsuntiea et al, extracted the intracellular
lipase from Mortierella alliacea by homogenising with glass beads and investigated
its substrate selectivity. The novel lipase preferentially hydrolysing monoacylglycerols
containing long-chain unsaturated fatty acids and phosphatidylcholine (Jermsuntiea et
al., 2011). Perez et al, isolated and purified a novel halophilic lipase from
Marinobacter lipolyticus SM19 by ultrasonic treatment. This lipase was employed for
hydrolysing fish oil and preferentially hydrolysed EPA from triglycerides over DHA
(Pérez et al., 2011).
1.5.2. Lipase based methods for concentration of omega-3 fatty acids
Lipases are able to catalyse unique reactions such as hydrolysis, ethanolysis or
transesterification of triglycerides. Fatty acids are naturally distributed in the glyceride
‘backbone’ of several marine oils, and these unique properties of lipases are very
useful both in omega-3 concentration from fish oil and for the production of structured
lipids (Ando et al., 1992; Tengku-Rozaina & Birch, 2014). Lipase based methods can
improve the quality of fish oil concentrates by causing less damage to EPA and DHA
during processing, which impacts the sensory quality and stability of the final products
(Venkateshwarlu et al., 2004). EPA and DHA have multiple double bonds in the cis-
configuration and these are very prone to oxidation, polymerization or isomerisation.
Concentrated omega-3 oils can contain high levels of products such as polymers and
trans fatty acids due to oxidative degradation (Kralovec et al., 2010). An effective way
of concentrating omega-3 fatty acids (EPA and DHA) is selective hydrolysis or
esterification of shorter chain fatty acids and their separation from the omea-3 fatty
28
Chapter 1 acid enriched glycerides (Carvalho et al., 2009; Kralovec et al., 2010). Many
commercial lipases available from fungi and bacteria discriminate against omega-3
fatty acids in that they preferentially hydrolyse shorter chain fats (Haraldsson, 1999).
Saturated and monounsaturated fatty acids are partially selectively cleaved from
triglycerides and long-chain omega-3 fatty acids primarily remain in the glyceride
portion as residual mono-, di- and triacylglycerols. The resistance of omega-3 fatty
acids to lipase hydrolysis by lipases is related to the presence of cis double bonds that
causes steric hindrance and results in bending of the fatty acid chains, which brings
the terminal methyl groups close to the ester bond (Okada & Morrissey, 2007). A more
direct approach is selective hydrolysis of omega-3 fatty acids. However, no omega-3
selective lipases have been characterised, although fish digestive juice may contain
such a lipase (Halldorsson et al., 2004). A more recent study also observed the
presence of a lipase with preferential EPA and DHA selectivity from fish waste
(Kurtovic & Marshall, 2013). A novel lipase isolated from the halophilic bacterium
Marinobacter lipolyticus SM19 showed high efficiency in the enrichment of EPA from
fish oil. These results indicate that selective concentration of omega-3 fatty acids from
marine oils using lipases is possible through selective EPA and/or DHA hydrolysis.
Table 1.5. Main patents relating production of omega-3 fatty acids processing using
lipases from the last 10 years.
S.
NO
Year Title Patent Number Inventors
1 2015 Separation of omega-3 fatty
acids
WO2015024055
A1
Colin James
Barrow, Taiwo
Olusesan Akanbi
2 2014 Method of producing oil/fat
comprising highly-
unsaturated fatty acids by
means of lipase
EP 2682453 A1 Nobushige Doisaki,
Hideo Ikemoto,
Kazuhiko Hata,
Shinji Tokiwa,
Kazunori
Matsushima
3 2013 Process for production of
oil or fat containing highly
EP 2578691 A1 Hideo Ikemoto,
Nobushige Doisaki,
Yasuo Umehara
29
Chapter 1
unsaturated fatty acid using
lipase
4 2013 Process for separating
polyunsaturated fatty acids
from long chain unsaturated
or less saturated fatty acids
EP 2585570 A1 Gudmundur G.
Haraldsson, Bjorn
Kristinsson
5 2013 Method for enrichment of
eicosapentaenoic acid and
docosahexaenoic acid in
source oils
WO
2013043641 A1
Subramaniam
Sathivel, Huaixia
Yin
6 2012 Enrichment of marine oils
with omega-3
polyunsaturated fatty acids
by lipase-catalysed
hydrolysis
WO2012087153
A1
Kahveci Derya, Xu
Xuebing
7 2011 High-Purity Purification
Method for Omega-3
Highly Unsaturated Fatty
Acids
US
20110091947
A1
Gap Jin Kim, Hong
Joo Son, Wu Song
Whang, Yoon Mo
Koo, Jin Il Kim, Jin
Hyo Yang,
8 2011 Processes to generate
compositions of enriched
fatty acids
WO
2011067666 A1
Inge Bruheim,
Svein Rye
9 2011 Method for producing oil
containing polyunsaturated
fatty acid using lipase
CA 2800674 A1 Hideo Ikemoto,
Nobushige Doisaki,
Yasuo Umehara
10 2010 Triglycerides with high
content of unsaturated fatty
acids
WO
2010115764 A2
Per Munk Nielsen,
Jesper Brask,
Steffen Ernst
30
Chapter 1
11 2010 A polyunsaturated fatty
acid (PUFA) enriched
marine oil comprising
eicosapentaenoic acid
(EPA) and docosahexaenoic
acid (DHA), and a process
of production thereof
EP 2147088 A1 Patrick
Adlercreutz, Ann-
Marie Lyberg
12 2010 Production method of oil or
fat containing
polyunsaturated fatty acid-
containing triglyceride
US 7767427 B2 Kengo Akimoto,
Motoo Sumida,
Kenichi
Higashiyama,
Shigeaki Fujikawa
13 2010 A polyunsaturated fatty
acid (PUFA) enriched
marine oil comprising
eicosapentaenoic acid
(EPA) and docosahexaenoic
acid (DHA), and a process
of production thereof
EP 2147088 A1 Patrick
Adlercreutz, Ann-
Marie Lyberg
14 2010 Enzymatic modification of
oil
EP 2183376 A2 Colin James
Barrow, Jaroslav
A. Kralovec,
Weijie Wang
15 2010 Method for production of
epa-enriched oil and dha-
enriched oil
EP 2172558 A1 Kiyomi Furihata,
Hiroyuki
Kawahara, Hideaki
Yamaguchi, Hideo
Ikemoto,
Nobushige Doisaki
16 2009 Method for production of
condensed polyunsaturated
fatty acid oil
EP 2006389 A9 Nobushige Doisaki,
Kiyomi Furihata,
Hiroyuki Kawahara
31
Chapter 1
17 2008 Process of selective
enzymatic enrichment of a
mixture containing omega-3
WO
2008093378 A1
Giovanni Cotticelli,
Raul Salvetti
There are numerous research articles and patents published in relation to the
production of omega-3 fatty acids from marine oils using lipases. Table. 1.6
summarises the patents filed in the last ten years on the production of omega-3 fatty
acids using lipase catalysed reaction. Linder et al, hydrolysed salmon oil using a sn-1,
3 specific lipase to concentrate PUFA 39.20 mol% to 43.05 mol% (Linder et al., 2002).
Another study by Gámez-Meza et al. 2003, investigated the enzymatic hydrolysis of
sardine oil and concentration using urea complexation. They also compared the effect
Pseudomonas lipase hydrolysis with chemical hydrolysis using KOH, ethanol and
hexane. This study concluded that the immobilized lipase with 10% protein was the
most effective system by which to achieve the best purity of omega-3 fatty acids
(86.58%) with an acceptable yield (78.0%). Although chemical hydrolysis yielded
higher omega-3 recovery (90.5%), the purity was lower (83.13%). Moreover, the
lipase based method resulted in easier separation of the products using a packed-bed
reactor (Gámez-Meza et al., 2003). A recent study optimised the hydrolysis of anchovy
fish oil and the concentration of omega-3 fatty acids using TL 100 lipase, from which
24% DHA was concentrated in the glycerol portion under optimised conditions
(Akanbi et al., 2013). Kahveci and Xu, reported the repeated hydrolysis of salmon fish
oil after distillation using Candida rugosa lipases in a 1 L stirred tank reactor resulted
in PUFA content of 38.7mol%, double of the initial level (Kahveci & Xu, 2011). These
approaches show the potential of lipase based methods to improve the nutritional value
of marine oils. Compared to existing chemical processes, lipase based approaches are
more environmentally friendly. Further research is required to obtain more selective
lipases and to optimise suitable production and immobilisation methods.
32
Chapter 1 1.6. Research aim and objectives
This research was focused on downstream processing of lipid and lipase from
thraustochytrids for the concentration of omega-3 fatty acids. The main objectives
were:
1. To establish a rapid lipid quantification method for the screening and
identification of thraustochytrid strains from marine samples collected from
Queenscliff, Victoria.
2. To determine which enzymatic activities are present in these thraustochytrid in
house isolates.
3. To develop methods for the extraction of total lipids and lipases from selected
thraustochytrids.
4. To investigate lipase production from thraustochytrid in a stirred tank reactor.
5. To develop a novel enzymatic method for concentrating omega-3 fatty acids
from marine oils, including thraustochytrid oils.
33
Chapter 2
Chapter 2: A quick colorimetric method for total lipid
quantification in thraustochytrids
34
Chapter 2 3.1. Introduction
Biodiesel is a renewable fuel of commercial interest, driven in part by the rising
cost of fossils and negative environmental impacts associated with conventional fuel
sources (Sadeghinezhad et al., 2014). First generation biodiesel was produced from
biomass, vegetable oils, animal fats and waste oils by transesterification (van Eijck et
al., 2014). The use of edible oils for first-generation biodiesel has resulted in the
undesired effect of increasing global competition for food crops (Kumar & Sharma,
2015; Pinzi et al., 2014). Second-generation biofuels utilise non-competing crops such
as jatropha, karanja, jojoba, mahua as well as the use of waste cooking oil, grease, and
animal fats. While the use of “inedible” crops reduces competition with food crops,
second-generation biofuels still compete for arable land, which ultimately negatively
impacts the world food supply and can contribute to the destruction of soil resources.
In addition, these feedstock cannot meet current energy demands (Nautiyal et al., 2014;
Zhu et al., 2014). Sustainable alternatives must be found that can provide the large fuel
volumes required, while not compromising the use of agricultural land for food
production. Microalgae have attracted attention as a means of addressing rapidly
growing demand for feedstock in biodiesel production (Baganz et al., 2014; Bellou et
al., 2014; Chen et al., 2015b). Microalgae are easy to cultivate and can be grown on
non-arable land, as well as accumulate more lipids as a percentage of their biomass
than plants (Challagulla et al., 2015).
Production of biodiesel from heterotrophic microalgae requires expensive
nutrients, particularly glucose as a carbon source (Slade & Bauen, 2013). Glucose can
constitutes 80 % of the total medium cost and up to a third of total fermentation
production costs (Li et al., 2007). To reduce process costs and assist in the
commercialisation of biodiesel production from heterotrophic microalgae, sustainable
low-cost carbon sources are required. One approach is to couple microalgae cultivation
with the use of inexpensive carbon sources such as carbon rich waste water, or use
glycerol derived from the biodiesel industry (Di Caprio et al., 2015; Xin et al., 2010).
Production costs can also be decreased by discovering new microalgae with improved
lipid productivity (Ahmad et al., 2015). An important reason for the slow development
in potential microalgae strain isolation and identification is due to lack of quick and
high-throughput assays (Cheng et al., 2011b). Existing techniques used for the
screening and quantification of microbial lipids include gravimetric, chromatographic
35
Chapter 2 and fluorescence dye methods (Bligh & Dyer, 1959; Han et al., 2011; Ren et al., 2015).
Analysis of microbial lipids by gravimetric method is usually effective, but has
limitations. The gravimetric method is time-intensive, requires the use of toxic
chemicals and utilises relatively high levels of biomass. Spectrophotometric
approaches can use a fluorescent dye, Nile Red, for lipid quantification. However, the
sensitivity of this method is poor and microbial cell components such as proteins,
pigments and some other environmental factors may interfere with lipid quantification
using these methods (Cheng et al., 2011b). A disadvantage of using fluorescent dye is
it can photo-bleach during exposure to light (Govender et al., 2012). Microbial lipid
quantification using triethanol-amine copper salts is an alternative method, however,
analyses of fatty acids in the ultraviolet region results without any colour development
(Chen & Vaidyanathan, 2012).
Colorimetric sulfo-phospho-vanillin (SPV) method for lipid quantification
firstly introduced by Charbo and Charonnat, 1937 for lipid quatification in human
cerebrospinal fluid samples (Vatassery et al., 1981). Later, this method was modified
for quantification of lipids in various samples include serum, foods and ecological
samples . Van handel modified SPV methods
into micro scale for total lipids quantification in single mosquitos (Van Handel, 1985).
Thraustochytrids, marine heterokonts and classified as oleaginous
microorganism, have been identified as a potential candidate for feedstock in the
heterotrophic production of lipids (Chen et al., 2015d). Extensive screening from
marine habitats has led to the discovery of strains which produce more than 50% of
their biomass as lipids (Raghukumar, 2008). In this study, we developed a colorimetric
sulfo-phospho-vanillin (SPV) method (initially developed for the serum lipids
quantification (Charbol & Charonnat, 1937)) for quantifying of lipids in
thraustochytrids and other oils containing materials. The accuracy of the SPV method
was compared with gravimetric and gas chromatography analysis. This method is
useful for rapid screening of microalgal strains for biodiesel production.
3.2. Materials and Methods
2.2.1. Chemicals used
All chemicals and medium components used in this study were procured from
Sigma-Aldrich, Australia. Instant ocean sea salts were procured from Aquarium 36
Chapter 2 Systems Inc., USA. Schizochytrium oil (batch number 11071602J) was obtained from
Ocean Nutrition Canada, Canada.
2.2.2. Cultivation of thraustochytrids
A total of four thraustochytrids, Schizochytrium sp. S31 (ATCC 20888),
Thraustochytrium sp. PRA 296, Thraustochytrium sp. AMCQS5-5 and
Schizochytrium sp. DT3 (in-house isolates) were selected for the current study.
Thraustochytrium S31 and PRA 296 were procured from the ATCC and used as
standard cultures, whereas AMCQS5-5 (JX993841) and DT3 (KF682125) were
isolated from Australian marine samples (Gupta et al., 2013). Seed culture was
prepared in GYP medium containing: Glucose 0.5 %, yeast extract 0.2 %, peptone 0.2
% in 50 % artificial seawater (Instant Ocean sea salts) and incubated at 150 rpm, 25
°C for 48 h. The inoculum (5 %) was later transferred into production media containing
glycerol (1 %), yeast extract (0.2 %) and peptone (0.2 %) and incubated at 150 rpm,
25 °C for 120 h. The cultures were harvested by centrifugation at 8000 rpm for 10 min,
freeze dried and stored at -20 °C for further use.
2.2.3. Standard curve preparation
For preparing a standard curve, commercial Schizochytrium oil was dissolved
in chloroform (10 mg in 10 ml) to a final concentration 1 mg mL-1. For the preparation
of the standard curve, different concentrations (15-90 μg) of lipid samples were taken
in clean glass tubes. The tubes were incubated at 60 °C for 10 min to evaporate the
chloroform. 100 μl of distilled water was added to each tube. Lipid quantification was
done by SPV reaction.
2.2.4. Quantification of thraustochytrid lipids by SPV
Vanillin phosphoric acid reagent was prepared by dissolving 0.120 g of vanillin
in 20 ml of distilled water and the final volume was adjusted to 100 ml with 85 %
phosphoric acid. This reagent was stored in the dark until further use. SPV reagent was
prepared fresh, which resulted in high activity with lipid samples. A known amount of
thraustochytrid dried biomass was suspended in distilled water and added to each test
tube (0.2-1 mg). Tubes were incubated at 100 °C for 10 min and cooled for 5 min in
an ice bath. Vanillin-phosphoric acid reagent (5 ml) was added to each tube and
37
Chapter 2 incubated in a shaker incubator at 200 rpm at 37 °C for 15 min. Absorbance was
measured at 530 nm.
The major steps involved in this method are: (a) phospho-vanillin reagent
preparation using phosphoric acid and vanillin, (b) addition of concentrated sulfuric
acid to a sample that contains lipids of interest, and heating of the mixture, (c) addition
of phospho-vanillin reagent, and (d) measurement of absorbance at 530 nm (Knight et
al., 1972). Fig. 2.1 illustrates in details the steps involved in the SPV method.
Figure. 2.1. Schematic diagram of SPV method. Concentrated sulphuric reacts with
unsaturated lipids microalgae and forms a carbonium ion. Phosphoric acid and vanillin
reacts and form a phosphate ester, which increases the reactivity of carbonyl group.
Carbonium ion and carbonyl group (from phospho vanillin reaction) reaction results
in formation of a stable colour compound.
2.2.5. Lipid extraction
Lipid was extracted and quantified from freeze dried biomass every 48 h
according to a previously published protocol (Lewis et al., 2000) with some 38
Chapter 2 modifications. Freeze-dried bio solvent (chloroform
and methanol in 2:1 ratio) and the sample vortexed for 2 min, followed by
centrifugation at 13,000 rpm for 15 min. This extraction process was repeated three
times. The three supernatants were collected and dried at 50 °C. The lipid percentage
was measured gravimetrically.
All the experiments were performed in triplicates. Results are represented as
mean values and standard deviation is < 10%.
2.3. Results and discussion
2.3.1. Biomass and lipid production
Production of biodiesel from inexpensive carbon sources will reduce oil
production costs. Glycerol is an economical carbon sources for microbial fermentation
(Leite et al., 2015) and has been tested as a carbon source for growing thraustochytrids
(Gupta et al., 2013; Scott et al., 2011). After 5 days, the biomass obtained were 938
mg L-1 d -1 in DT3, 950 mg L-1 d -1 in S31, 740 mg L-1 d -1 in AMCQS5-5 and 256 mg L-
1 d -1 in PRA-296 (Table 1). Schizochytrium sp. S31 and DT3 produced similar biomass
yields, whereas Thraustochytrium sp. AMCQS5-5 and PRA-296 exhibited lower
biomass productivity, depending on how efficiently the strain utilised the carbon
source to produce biomass. However, lipid productivity was documented to be higher
in DT3 at 287.96 mg L-1 d -1 when compared to other strains. The next highest biomass
was produced by S31 (227.05 mg L-1 d -1) followed by AMCQS5-5 (158.36 mg L-1 d -
1) and PRA-296 (51.71 mg L-1 d -1) (Table 1).
Table 2.1. Biomass and lipid production in glycerol medium by thraustochytrids.
Item DT3 S31 AMCQS5-5 PRA-296
Biomass (g L-1) 4.69 ± 0.09 4.75 ± 0.1 3.7 ± 0.01 1.28 ± 0.005
Biomass productivity (mg L-1 d -1) 938 ± 0.19 950 ± 0.02 740 ± 0.002 256 ± 0.001
Average lipid content (mg L-1) 1439 ± 92 1135 ± 68 791 ± 49 258 ± 15
Lipid productivity (mg L-1 d -1) 287 ± 28 227 ±17 158 ± 11 51 ± 4
39
Chapter 2 2.3.2 Microalgae oil quantification by SPV method
The colorimetric method was first validated to generate the calibration curve
with known quantities of Schizochytrium oil. A known quantity of lipid standard in the
range of 15-90 μg was taken in clean test tubes and placed in a hot air oven at 60 °C
to evaporate chloroform, followed by the SPV reaction protocol described in the
experimental section. Lipid content was measured by plotting absorbance at 530 nm
and lipid concentration. A high correlation coefficient was obtained with R2 value
0.9939 (Fig. 2.2). This demonstrates the high accuracy of the SPV method in
quantifying lipids in aqueous samples.
Figure. 2.2. Standard curve representing relationship between Schizochytrium oil
concentration and increase in the absorbance. The lipid concentrations were from 15
to 90 μg. Standard curve representing relationship between Schizochytrium oil
concentration and increase in the absorbance.
Interference by other cellular and medium components such as protein (BSA),
carbohydrate (Glucose) and glycerol did not occur with the SPV reagent (Fig. 2.3),
which is consistent with a previous study (Izard & Limberger, 2003). These medium
compounds were used as false positives. Thus, other compounds in the medium
showed no effect on the absorbance value.
y = 0.0033x - 0.0258R² = 0.9939
0
0.05
0.1
0.15
0.2
0.25
0.3
0 20 40 60 80 100
Abs
orba
nce
at 5
30 n
m
Schizochytrium oil (μg)
40
Chapter 2
Figure. 2.3. Comparison of various lipids and other compounds with sulfo-phospho-
vanillin reaction. Triolein yielded intense colour compared with other lipid standards.
Compounds with no bonds in their structure do not react with the SPV reagent.
The basic principle underlying the SPV method is a three step reaction. When
the lipids are heated in the presence of sulphuric acid at boiling temperature (low
temperature leads to lower reaction rates), a carbonium ion is formed. An aromatic
phosphate ester is formed from the reaction of vanillin and phosphoric acid. The
activated carbonyl group of phospho-vanillin reagent reacts with the carbonium ions
to form a stable pink colour, which has a maximum absorbance at 530 nm (Knight et
al., 1972). Colour formation depends mainly on the molecular structure of fatty acids
present in the lipids. The molecules which do not possess double bonds do not
detectably react with sulfo-phospho-vanillin reagent. Colour intensity varies from one
lipid to another due to differences in fatty acid structure. It was observed that the colour
intensity of the triolein was greater than that of the polyunsaturated fatty acids EPA
and DHA (Fig. 2). The presence of multiple double bonds in EPA and DHA did not
yield more colour due to steric hindrance near these double bonds. Saturated fatty acids
such as palmitate and stearate did not react with sulfo-phospho-vanillin reagent (Frings
& Dunn, 1970; Knight et al., 1972). Among the plant oils tested in this study, olive oil
0
0.05
0.1
0.15
0.2
0.25
0.3
0.35
0.4
0.45
0.5
0 20 40 60 80
Abs
orba
nce
(530
nm
)
Different oil samples (μg)
SunflowerOliveCanolaTrioleinEPADHAPalmitateSterateGlycerolGlucoseBSA
False positives
Lipid samples
41
Chapter 2 resulted in higher levels of colour formation than did canola and sunflower oil,
probably due to the presence of more than 70% oleic acid in olive oil. In general, higher
levels of unsaturated fatty acids lead to higher absorbance, although the presence of
multiple double bonds in a single fatty acid (polyunsaturated) may lead to the
formation of intense colour (Knight et al., 1972). The presence of various fatty acids
in microbial lipids limits the accuracy of the method. This can be rectified by selecting
a suitable oil with the same fatty acid content as a reference. Previous studies have
reported the quantification of bacterial and microalgal lipids with reference to triolein
and canola oil, respectively (Izard & Limberger, 2003; Mishra et al., 2014). We used
commercial Schizochytrium oil as reference for quantification of lipids in
thraustochytrids.
2.3.3. Lipid quantification in thraustochytrid whole cells via SPV method
The established colorimetric method was applied to the quantification of lipids
in whole thraustochytrid cells. A known quantity of dried biomass of all four
thraustochytrid cultures was suspended in distilled water at a concentration of 20 mg
ml-1. Aliquots of 10-50 μl were taken in clean test tubes and the final volume was made
up to 100 μl with distilled water. Fig. 3 shows the correlation coefficient between
biomass concentration and absorbance with an R2 > 0.99. All four cultures showed an
increase in the intensity of pink colour formation with respect to the increasing
biomass concentration (Fig. 2.4). Lipid content in all four cultures was quantified by
converting the absorbance into mass using a reference curve with a low standard
deviation. Colour change from low concentration to high concentration of biomass
(lipid) in four different thraustochytrids shown in Fig. 2.5.
42
Chapter 2
Figure. 2.4. Profile of colorimetric detection of thraustochytrid whole cell lipids. The
relationship between the concentration of biomass and absorbance at 530 nm for
DT3 (R2= 0.992), S31 (R2=0.999), AMCQS5-5 (R2=0.993) and PRA-296 (R2 =
0.998)
y = 0.993x + 0.0039R² = 0.9926
y = 0.888x - 0.0154R² = 0.999
y = 0.7308x + 0.0138R² = 0.9932
y = 0.516x + 0.0121R² = 0.9985
0
0.2
0.4
0.6
0.8
1
1.2
0 0.2 0.4 0.6 0.8 1 1.2
Abs
orba
nce
at 5
30
Biomass (mg)
DT3 S31AMCQS5-5 PRA 296
43
Chapter 2
Figure. 2.5. Change of lipid colour after reaction with sulfo-phospho-vanillin reagent.
Increase of colour represents the correlation between the concentrations of lipid in
thraustochytrids. (a) DT3, (b) S31, (c) AMCQS5-5 and (d) PRA-296.
Strain development and lipid quantification are major milestones on the path
to sustainable production of biodiesel from the microbial sources. A rapid lipid
quantification technique is required to screen potential lipid producing strains from
environmental samples. In this study, we used a colorimetric method (SPV) to quantify
the total lipids in an aqueous medium containing thraustochytrids, a potential lipid
producer. The lowest amount of biomass used for quantifying the lipid content was 50
μg. This enables us to quantify the lipids from the thraustochytrids cells without an
extraction step. Moreover, the colorimetric method not only quantified lipid content
accurately, but also required less than one hour to complete. In contrast, gravimetric
quantification of lipids, extraction and GC analysis are more time and labour intensive.
2.3.4. Validating the accuracy of the SPV method
Gravimetric quantification of lipids from the four selected thraustochytrids was
used in order to validate the accuracy of the SPV method. Thraustochytrid lipids were
44
Chapter 2 extracted using a conventional solvent extraction method with modifications (Lewis
et al., 2000). The total lipid percentages determined gravimetrically from the four
thraustochytrids DT3, S31, AMCQS5-5 and PRA-296, were 33.3, 25.8, 22.8 and 18.0,
respectively. Lipid percentages determined also by the SPV method showed similar
quantities in all thraustochytrids. Lipid contents determined by SPV, gravimetric
methods and gas chromatography were compared (Table 2.2). The lipid profiles
determined by gas chromatography were 27.3, 21.8, 20.3 and 18.4, respectively. In
two thraustochytrid strains, AMCQS5-5 and PRA 296, the lipid percentages were the
same as those obtained using the SPV method. However, in DT3 and S31 there was
small (1.9-2.6 %) difference in the lipid percentages. These results obtained from
standard gravimetric method for lipid quantification compared with the SPV method
in quantifying the total lipids from thraustochytrid whole cells, and the variation was
less than 3%, which confirms the suitability of SPV method in screening microbial
samples.
Table 2.2. Comparison of thraustochytrid lipid content by SPV, gravimetric and GC
methods.
Thraustochytrid
strains
Lipid
percentage
by SPV
method
Lipid
percentage by
gravimetric
method
Variation Lipid
percentage by
gas
chromatography
DT3 33.3 ± 1.2 30.7 ± 0.59 2.6 27.3 ± 1.3
S31 25.8 ± 0.9 23.9 ± 0.86 1.9 21.8 ± 0.84
AMCQS5-5 22.8 ± 1.1 21.4 ± 0.78 1.4 20.36 ± 0.9
PRA-296 18.0 ± 1.4 20.2 ± 0.62 2.2 18.47 ± 0.81
Microalgal lipids are useful for biodiesel production. Isolation and
identification of the high lipid-producing microalgae is a key step in achieving
economically sustainable biodiesel production. Quantification of lipids from
microalgae must be performed frequently in microbial screening and other research
areas, such as in the screening of genetically engineered microbes for high lipid
production and so higher throughput accurate methods are important.
45
Chapter 2 2.4. Conclusion
In conclusion, this study demonstrated the use of the sulfo-phsopho-vanillin
reaction to quantify lipid accumulation in thraustochytrids. The SPV method resulted
in similar lipid content determination in thraustochytrid whole cells as did gravimetric/
gas chromatography analysis. Triolein contains only mono-unsaturated fatty acids and
the colour intensity was greater than for polyunsaturated fatty acids with multiple
double bonds. Carbohydrates, proteins and glycerol did not interfere with the SPV
reaction. SPV is a rapid method that requires low levels of chemicals and so is useful
for the screening of microalgal cultures for biodiesel production.
46
Chapter 3
Chapter 3: Comparison of cell disruption methods for
improving lipid extraction from thraustochytrid strains
47
Chapter 3 2.1. Introduction
The limited availability of fossil fuels and increasing greenhouse gases
emission has led to interest in alternative biomass derived energy. As an alternative
fuel, biodiesel is promising (Azeem et al., 2016). Biomass derived biofuels are
classified as solid (bio-char), liquid (ethanol, vegetable oil, and biodiesel), and gaseous
(biogas, biosyngas and biohydrogen) fuels (Mubarak et al., 2015). Biodiesel is
renewable and a relatively clean energy in terms of carbon dioxide generation and
greenhouse gas emissions. Biodiesel accounts for 10% of total biofuel production
globally and its estimated production is about 6 billion litres/year (Nogueira, 2011).
The present approach to biodiesel (defined as the monoalkyl esters of long-chain fatty
acids) production involves transesterification of plant oils such as soybean oil,
sunflower oil and rapeseed oil with methanol using alkali catalysts (Daroch et al.,
2013; Meher et al., 2006). The use of plant oils for biodiesel production has been
associated with some drawbacks such as high viscosity, low volatility and deposition
in combustion chambers (Lin et al., 2011). Also, the use of edible oils for biodiesel
production has led to an increase in the price of oils, initiating a food versus fuel debate
regarding biofuel sustainability. Feedstock useful for biofuel include soapstocks, acid
oils, tallow oils, used cooking oils, various animal fats, non-edible plant oils and
microbes including algae (Chisti, 2007; Silitonga et al., 2013).
Microalgae are promising vehicles for the production of biodiesel and possess
advantages such as higher growth rate and productivity, grow in various environments
(fresh, brackish or salt water), do not compete for land, and have high oil productivity
(20-50% by dry weight basis) compared to conventional crops (Singh et al., 2011).
Microalgae lipid content differs between microalgae species. Microalgae lipid
composition and fatty acids profiles depends on culture media composition (carbon
source and nitrogen source), temperature, pH and aeration (Halim et al., 2012a). The
process of lipid extraction from microalgae is an energy intensive and expensive
because the use of solvents and recovery methods in downstream processing.
Selection of efficient microalgae species and suitable lipid extraction methods are
important for commercial biodiesel production (Griffiths & Harrison, 2009; McMillan
et al., 2013). Biodiesel production from microalgae involves four major stages that are
cultivation, cell harvest, lipid extraction and finally conversion of lipids into biodiesel
(Lee et al., 2010b; Schnurr & Allen, 2015). Therefore, a suitable lipid extraction
48
Chapter 3 technique is a prerequisite for microalgal lipid extraction. Lipid extraction efficiency
is dependent on the polarity of the solvent and combination of solvent mixture (Lee et
al., 2010b; Lee et al., 1998; Lewis et al., 2000). The combination of a polar and non-
polar solvent mixture can in some cases extract more lipids from microalgae
(Ramluckan et al., 2014; Ryckebosch et al., 2012b). For example, the Bligh and Dyer
method uses chloroform and methanol for lipid extraction from a range of biological
samples (Ramluckan et al., 2014). The use of chloroform and ethanol in a 1:1 ratio
provided maximum lipid extraction from Chlorella sp. (Ramluckan et al., 2014),
whereas, a combination of dichloromethane and ethanol increased lipid extraction
efficiency by 25% in the same organism (R et al., 2015).
Microalgae cell walls block the release of lipid present inside the cells. To get
higher product recovery and quality lipids with lower operating costs from microbial
cells, a suitable cell disruption method is required. Cell disruption enhances the release
of intracellular lipids from microalgae by improving the access of the extracting
solvent to fatty acids (Halim et al., 2012b). Cell disruption methods used in microalgae
lipid extraction are classified as chemical and mechanical. Chemical cell disruption
methods involve use of acids, alkalis, enzymes or osmotic shocks and mechanical
methods use microwave, ultrasonication, bead mill, drying, or supercritical fluid
extraction. The choice of method influences lipid extraction yields from a range of
microalgae (Lee & Han, 2015; Lee et al., 2010b; McMillan et al., 2013; Zheng et al.,
2011).
The aim of this study was to compare various organic solvents and cell
disruption methods for effective lipid extraction from a new strain of thraustochytrids
(a newly isolated strain from the Queenscliff region, Victoria, Australia). This strain
was used as a representative microalgae in this work due to its ability to accumulate
high levels of lipids (Gupta et al., 2013). In addition, microalgae biomass was used for
different extraction methods that were successfully used for efficient algal lipid
extraction in previous studies. Finally, the conversion of lipids to fatty acid methyl
ester (FAME) was performed to determine yield and fatty acid distribution after
extraction.
3.2. Experimental Section
3.2.1. Chemicals
49
Chapter 3
All the chemicals used in this study were of analytical grade. Medium
components such as glucose, yeast extract and mycological peptone (Sigma-Aldrich,
USA) and sea salt (Instant Ocean, USA) were used for biomass production while
solvents such as acetone, ether, hexane (Merck, Australia), methanol and ethyl acetate
from Fischer and Honeywell, Australia.
3.2.2. Strain selection and biomass production
Schizochytrium sp. S31 (ATCC 20888) was procured from American Type
Culture Collection (ATCC) and used as standard culture. Thraustochytrids used in this
study were maintained on GYP medium consisting (g L-1): glucose 5, yeast extract 2,
mycological peptone 2, agar 10 and artificial seawater 50% at 25 °C and sub-cultured
after 15 days.
Thraustochytrium sp. AMCQS5-5 (an in house isolate; Genbank accession
number JX993841), was grown in a medium containing (g L-1): glucose 5, peptone 2,
yeast extract 2 and artificial sea water 50% for inoculum preparation with shaking at
150 rpm for 2 days at 25 oC. The medium was autoclaved at 121 oC for 20 min.
Inoculum (5% v/v) was used to inoculate production medium (100 ml contained in 500
ml flask) and incubated for 5 days in a shake flask at 25 oC and 150 rpm. The resultant
biomass was harvested by centrifugation (4000 rpm for 15 min) and was freeze-dried
and kept at -20 oC until further use.
The thraustochytrids grown in culture medium were harvested at the interval
of 24 h up to 120 h. Optical density at 600 nm and dry cell weight (DCW) was
measured at 24 h intervals. A calibration curve was plotted between OD and dry cell
weight. Results are presented as mean ± SD of duplicates repeated twice.
The biomass and lipid productivity was calculated from the formula mentioned below:
Productivity = Biomass or Lipid content/T1-T0
Where, T1 = Final day of biomass harvesting and T0 = Initial day of incubation
3.2.3. Lipid extraction from thraustochytrids by organic solvents
The nine solvents, chloroform, dicholoromethane, diethylether, ethanol,
heptane, hexane, isopropanol, methanol and toluene at ratios of 1:1, 1:2 and 2:1 were
tested for maximising extraction of total lipids. For solvent extraction, 50 mg of freeze
50
Chapter 3 dried biomass of thraustochytrid was blended with 3 ml of various solvents. The
mixture was vortexed for 2 min and the sample was then centrifuged at 4000 rpm for
15 min. Supernatant (organic phase) was carefully collected in the pre-weighed glass
vials and the solvent was evaporated under nitrogen gas at room temperature. Lipid
content (% dry weight basis) was determined gravimetrically. To determine the
optimal organic solvent mixture for lipid extraction from thraustochytrids, different
ratios of the best three single solvents were investigated. For chloroform-methanol, the
Bligh and Dyer method was followed (Bligh & Dyer, 1959).
3.2.4. Cell disruption for lipid extraction
Freeze dried biomass was blended with 3 ml of chloroform and methanol (2:1)
and disrupted by means of different cell disruption methods as detailed below. After
each treatment, lipid extraction from thraustochytrids was done according to Gupta
and co-workers (Gupta et al., 2012c). After centrifugation (4000 rpm for 15 min), the
upper layer was collected and dried under nitrogen gas. Lipid content (% dry weight
basis) was determined gravimetrically.
3.2.4.1. Grinding with liquid nitrogen
A sample of freeze dried thraustochytrid biomass (50 mg) was taken in the
ceramic mortar. About 10-15 ml liquid nitrogen was added and the sample was allowed
to thaw and grinded with pestle for 2 min. After grinding the lipid was extracted using
organic solvents.
33.2.4.2. Bead vortexing
Thraustochytrid cell suspension (50 mg) was taken in glass tube (35 ml) and 3
ml of solvent and 1 ml of beads (zirconia beads, size 0.4-0.6 mm, Klausen Pty Ltd,
Australia) were added and contents were vortexed for 20 min, using a vortex in 30
second bursts. Samples were kept on ice in between the bursts and lipid was extracted
using organic solvents.
3.2.4.3. Sonication
Thraustochytrid cell biomass (50 mg) was suspended in 3 ml of solvent in a 15
ml centrifuge tube. Sample was sonicated at 20 kHz, 40% amplitude and the pulse was
51
Chapter 3 40 seconds on and 20 seconds off with total working time of 20 min (Sonics, USA).
Sample tubes were kept in ice during the sonication process to prevent overheating.
3.2.4.4. Osmotic shock
Thraustochytrid suspension was disrupted using an osmotic shock method. 50
mg of thraustochytrid biomass was suspended in 3 ml of 10% NaCl solution (mass
concentration 16.6 Kg/m-3, 0.6 Kg of NaCl) and vortexed for 2 min and incubated for
48 h at room temperature, followed by solvent extraction. Cell disrupting pressure was
calculated by the Morse equation:
kPa), i is van 't Hoff factor, M is molar concentration of NaCl, R is 0.0821 L atm/K
mol, and T is absolute temperature in °K at which the osmotic shock was performed.
Osmotic shock was calculated using an online osmotic pressure calculation hosted by
Georgia State University.
3.2.4.5. Water bath
3 ml samples containing 50 mg of biomass taken in 15 ml centrifuge tube was
placed in preheated water bath (Ratek Instruments Pty Ltd, Australia) to induce
thermolysis. Samples were kept in a water bath for 20 min at 90 °C. Three tubes were
treated simultaneously as replicates without shaking.
3.2.4.6. Shake mill
Thraustochytrid cell suspension was disrupted in plastic sample bottle using
shake mill (SPEX Mill 8000M, USA). The biomass and bead ratio was 3:1 ratio
(Zirconia beads 0.4-0.6 mm) at maximum speed (1060 cycle/min) and exposed for 5
min.
All the cell slurries were observed under the microscope using differential
interference contrast (Axio-imager, Zeiss, Germany) to check the disruption of
thraustochytrid cells. Microbial smear was prepared on the glass slide, air dried and
observed under a microscope. All experiments were performed three times.
52
Chapter 3
3.5. Fatty acids methyl esters (FAMEs) analysis
Fatty acids were converted to methyl esters by acid-catalysed trans-
esterification according to a published method (Han, 2010). 1 ml toluene was added to
methanol (2 ml) was also added to the tube and kept for overnight incubation at 50 °C.
Fatty acid methyl esters (FAMEs) were extracted into hexane. The hexane layer was
removed and dried over sodium sulphate. FAMEs were concentrated using nitrogen
gas. The samples were analysed by a GC-FID system (Agilent Technologies, 6890N,
Santa Clara, CA, USA). The GC was equipped with a capillary column (SGE, BPX70,
ier gas at a flow rate
of 1.5 ml min 1. The injector was maintained at 250 °C and a sample volume
was injected. Fatty acid peaks were identified on comparison of retention time data
with external standards (Sigma-Aldrich, St. Louis, MO, USA) and corrected using
theoretical relative FID response factors (Ackman, 2002). Peaks were quantified with
Chemstation chromatography software (Agilent Technologies, Santa Clara, CA,
USA). Results are presented as mean ± SD of triplicates.
3.3. Results and Discussion
3.3.1. Biomass and lipid production
Schizochytrium sp. S31 showed highest biomass productivity at 0.81 g L-1 d-1
and Thraustochytrium sp. AMCQS5-5 at 0.64 g L-1 d-1 at the end of 5 days (Table 1).
The lipid productivity of Schizochytrium sp. S31 and Thraustochytrium sp. AMCQS5-
5 were 100.74 mg L-1 d-1 and 64.2 mg L-1 d-1, respectively. Selection of suitable
microalgae strain with adequate biomass and oil productivity is important for cost
effective biodiesel production. Our results on biomass and lipid productivity are in
agreement with previous findings by Vello et al. 2014 (0.30 g L-1 day-1 and 34.53 to
230.38 mg L-1 day-1) that demonstrated the suitability of Chlorella strain as a
promising candidate for biodiesel production (Vello et al., 2014).
53
Chapter 3 Table 3.1. Biomass, lipid contents and productivity of thraustochytrids.
Properties Thraustochytrid strains
Schizochytrium sp. S31 Thraustochytrium sp. AMCQS5-5
Dry weight (g L-1) 4.06 3.23
Biomass productivity (g L-1 d-1) 0.88 0.64
Average lipid content (mg L-1) 503.7 321.3
Lipid productivity (mg L-1 d-1) 100.74 64.2
Lipid content and fatty acid profile of a microorganism is dependent on the
growth conditions (Rodolfi et al., 2009; Salama et al., 2013). Medium composition
influences the percentage of total lipid and type of fatty acids in the microbe. For
example, the addition of tween 80 in the production medium led to the accumulation
of oleic acid in the thraustochytrids (Taoka et al., 2011). A recent study reported the
effect of seasonal variation and nitrogen limitation in the total lipid production and
fatty acid composition of Nannochloropsis oculata. They observed an increased
accumulation (up to 90%) of saturated and mono-unsaturated fatty acids (Olofsson et
al., 2014). Since the aim of this work was to find a suitable disruption method for lipid
extraction, higher biomass productivity was not pursued further.
3.3.2. Lipid extraction from thraustochytrid by organic solvents
The lipid extraction by organic solvents was examined to confirm the lipid-
extraction characteristics of thraustochytrids. Fatty acids present in the lipid govern
the polarity, based on the principle “like dissolves like”, thus a suitable solvent should
be identified for total lipid extraction, however a universal solvent cannot be applied
to all microbes with different fatty acid composition. Total lipid extraction yields vary
primarily with solvent polarity (Phukan et al., 2011).
To understand the efficacy of organic solvents in lipid extraction from
Schizochytrium sp. S31, nine solvents and their combinations (based on high lipid
yield) were selected, since the lipid extraction is highly dependent on the polarity of
the solvents and their ratios (Ramluckan et al., 2014). The effect of single solvent as
54
Chapter 3 well as the combinations of the best 3 solvents with respect to the lipid extraction from
Schizochytrium sp. S31 is shown in Figure 1. Among the single organic solvents,
maximum lipid (12.5%) was extracted with hexane, followed by 11 % in heptane and
9.7% in chloroform. However, among the combination of solvents, the mixture of
chloroform: methanol (2:1) showed maximum lipid extraction (22%), followed by
chloroform and hexane (2:1) with 13.4% (Fig. 3.1). Chloroform and hexane showed
comparatively low efficacy in lipid extraction indicating that a range of polar to non-
polar lipids were present in Schizochytrium sp. S31. A similar trend was observed
during lipid extraction from Chlorella sp. (Ramluckan et al., 2014). A mixture of
hexane:heptane (1:1) extracted the least amount of lipid (2.2%). Higher lipid yields
with chloroform: methanol indicates the presence of more polar and neutral lipids in
the algae. It was observed that the percentage of polar and neutral lipids were
approximately 78% and non-polar lipids were 22% in some algae (Kale, 2013).
Combination of polar and non-polar solvents could extract more lipids than individual
solvents (Lewis et al., 2000; Ryckebosch et al., 2012b). However, a study (Shen et al.,
2009) contradicts this conclusion, indicating that the combination of hexane and
ethanol could not extract more lipid than hexane alone from Scenedesmus dimorphus
and Chlorella protothecoides. This suggests the efficiency of lipid extraction depends
on the algal species and their lipid compositions. Also, results obtained from
chloroform and methanol (2:1) indicate the presence of more polar and neutral lipids
in algae (Ramluckan et al., 2014). Mixtures of chloroform:methanol extract
hydrocarbons, carotenoids, chlorophyll, sterols, triglycerides, fatty acids,
phospholipids and glycolipids (dos Santos et al., 2015; Singh et al., 2013).
55
Chapter 3
Figure. 3.1. Lipid extraction percentage from dry biomass using various solvents from
Schizochytrium sp. S31. The symbols represent Chl: Choloroform, Hex: Hexane, Hep:
Heptane, Met: Methanol.
3.3.3. Comparison of lipid extraction methods
Six methods were evaluated in order to determine the efficiency of cell
disruption methods for total lipid extraction from thraustochytrids. The effectiveness
of the cell disruption methods were quantified using lipid yield percentages. Cell
disruption breaks the cells and improves the accessibility to the intracellular
components for extraction (Wang et al., 2015). Figure 2 shows the percentage of total
lipids extracted as a function of cell disruption methods. All the cell disruption
methods used in this study were able to disrupt thraustochytrid cells, although lipid
yield varied.
0
5
10
15
20
25
30
Tota
l lip
id c
onte
nt (d
cw, %
)
Solvents and their combination used
56
Chapter 3
Figure. 3.2. Effect of different cell disruption methods for lipid extraction from
thraustochytrids. Average lipid extraction of each method was reported as % of total
lipids extracted.
Thraustochytrid cells subjected to disruption resulted in rupture of the cell
walls and release of intracellular components (Figure 3.3a and b). It was observed that
lipid content of Schizochytrium sp. S31 was higher than that of Thraustochytrium sp.
AMCQS5-5. Maximum lipid was extracted from both Schizochytrium sp. S31 (48.7%)
and Thraustochytrium sp. AMCQS5-5 (29.1%) cells using osmotic shock (Figure 2).
Grinding, sonication, shake mill and water bath treatments extracted 44.6%, 31%,
30.5% and 20.8% of lipids, respectively. Bead vortexing resulted in 25% lipid
extraction from Thraustochytrium sp. AMCQS5-5 cells, whereas other methods (water
bath, grinding, shake mill and sonication) resulted in lower yields. Osmotic shock
resulted in a 2.2- folds increment in lipid extraction from Schizochytrium sp. S31 and
a 2.8-folds increase from Thraustochytrium sp. AMCQS5-5, compared to control. A
similar study for a Chlorella sp. indicated that osmotic shock was an effective method
for extracting lipids (Prabakaran & Ravindran, 2011). This method also consumes less
energy than traditional methods. In a recent study, when Chlamydomonas reinhardtii
cells were incubated in a high osmotic environment, it led to a 2-folds improvement in
lipid extraction (Yoo et al., 2012). Table 3.2 summarises some cell disruption methods
reported for lipid extraction from microalgae. Due to differences in cell wall structure,
0
10
20
30
40
50
60
Control Grinding Beadvortexing
Osmoticshock
Water bath Sonication Shake mill
Tota
l lip
ids c
onte
nt (%
)
Cell disruption methods
S31 AMCQS5-5
57
Chapter 3 not all microalgae respond the same to pretreatment. For example, Lee et al. (2010)
observed that microwave treatment was optimum for the disruption of Botryococcus
sp., Chlorella vulgaris and Scenedesmus sp. cells. Another study showed that grinding
with liquid nitrogen facilitated higher levels of lipid extraction from Chlorella vulgaris
(Zheng et al., 2011). Available literature suggests that cell disruption methods improve
lipid extraction from microalgae, and efficiency depends on microalgae species, age
of the culture and composition of cell wall. Therefore, results obtained from one
species cannot be generalised to all other species (Halim et al., 2012b).
Figure. 3.3. Effect of cell disruption on thraustochytrids cells. Thraustochytrid cells
before (A) and after (B) cell disruption (scale bar 20 μm).
Table 3.2. Comparison of cell disruption methods employed to extract total lipids from
different microalgae.
S.
No.
Cell
disruption
methods
used
Efficient
method
Organism used Lipid
content
(%)
Reference
1 Autoclaving
Bead beating
Microwaves
Sonication
Osmotic
shock
Microwaves Botryococcus sp.
Chlorella vulgaris
Scenedesmus sp.
28.6
11
11.5
(Lee et al.,
2010b)
58
Chapter 3 2 Sonication
Osmotic
shock
Microwave
Autoclave
Bead beating
Sonication Chlorella sp.
Nostoc sp.
Tolypothrix sp.
20.1
18.2
14
(Prabakaran
&
Ravindran,
2011)
3 Grinding
Sonication
Bead Milling
Enzymatic
Lysis
Microwaves
Grinding Chlorella vulgaris 29 (Zheng et
al., 2011)
4 Grinding
Bead
vortexing
Osmotic
shock
Water bath
Sonication
Shake mill
Osmotic
shock
Schizochytrium
sp. S31,
Thraustochytrium
sp. AMCQS5-5
48.7
29.1
This study
There are many lab scale cell disruption methods discussed in the literature, however,
only a few mechanical methods either alone or with the intervention of enzymes/
chemicals can be scaled up for industrial applications. For instance, bead mill, high
pressure homogenizer and Hughes press are used extensively at large scale, which
reduce unit operation steps compared to chemical and enzymatic methods (Chisti &
Moo-Young, 1986; Schutte & Kula, 1990). An osmotic shock method was
implemented in thraustochytrid cell disruption and lipid extraction, to reduce the
energy consumption and production cost. During the osmotic shock treatment, the
resulting wastewater can be recycled through reverse osmosis technology (Arnal et al.,
2005). The same method has been applied at pilot-scale for enhancing the release of
ectoine (Sauer & Galinski, 1998). A recent study by Jayaranja and Rekha (2015)
indicated that osmotic shock was a suitable method for maximising the extraction of
intracellular products, and this method can also be industrially scaled up (Kar &
59
Chapter 3 Singhal, 2015). The advantages and disadvantages of selected investigated methods
are summarised in Table 3.3.
Table 3.3. Advantages and disadvantages of the investigated cell disruption methods.
Cell disruption
methods
Advantages Disadvantages
Manual grinding Quickest and efficient
- 2 min process
- Localised heating caused
denaturation of molecules
Bead vortexing - Can be established easily
and relatively effective
- High heat generation,
- Incomplete cell lysis
Osmotic shock - Lower energy
consumption
- Easier scale-up
- Generation of waste salt
water
- Time consuming
Water bath - Maximum disruption
- Easy in handling at lab
scale
- Increases the viscosity
- Energy intensive
Sonication - Faster extraction
- suitable for all cell type
- Damage chemical
structure of molecules
Shake mill - Rapid method - High energy intensive
- High heat generation
3.3.4. Fatty acid composition of extracted lipid
The fatty acid profiles of the lipids extracted following different cell disruption
methods from thraustochytrids are presented in Figure 4a and 5a. Major fatty acids
such as myristic acid (28.1%), palmitic acid (27.3%), palmitoleic acid (20.7%) and
oleic acid (12.8%) were detected in Schizochytrium sp. S31 (Figure 3.4a) after osmotic
shock disruption. Total saturated, monounsaturated and polyunsaturated fatty acid
contents after osmotic shock facilitated extraction were 58.7%, 34.6% and 4.8%,
respectively, making it a potential feedstock for biodiesel production (Figure 3.4b).
The other prominent fatty acids based on grinding, sonication and shake mill identified
in the lipid extracts were saturated (49-57%), mono-unsaturated (31-35%) and
polyunsaturated fatty acids (2-18%). Saturated and monounsaturated fatty acids are
useful major components for microalgal biodiesel production because of their
60
Chapter 3 relatively high oxidative stability (Cao et al., 2014). The general properties of biodiesel
such as viscosity, specific gravity, cetane number, iodine value, and low temperature
performance metrics are determined by the structure (length and unsaturation) of fatty
acid esters (Hoekman et al., 2012).
Figure. 3.4a. Effect of different cell disruption methods on fatty acid profiles of
Schizochytrium sp. S31.
0
5
10
15
20
25
30
C14:0 C15:0 C16:0 C16:1n7 C18:0 C18:1n9C18:1n7C20:5n3C22:5n6C22:6n3
Tota
l lip
id c
onte
nt (%
)
Major fatty acids
Control Grinding with liquid N2Bead vortexing Osmotic shockWaterbath SonicationShake Mill
61
Chapter 3
Figure. 3.4b. Effect different cell disruption methods on SFA, MUFAs and PUFAs of
Schizochytrium sp. S31.
Figure. 3.5a. Effect of different cell disruption methods on fatty acid profiles of
Thraustochytrium sp. AMCQS5-5.
0
10
20
30
40
50
60
70
SFAs MUFAs PUFAs
Tota
l lip
id c
onte
nt (%
)
Fatty acids composition
Control Grinding with liquid N2Bead vortexing Osmotic shockWaterbath SonicationShake Mill
0
10
20
30
40
50
60
C16:0 C18:0 C18:1n9 C18:1n7 C20:4n6 C20:5n3 C22:5n6 C22:5n3 C22:6n3
Tota
l lip
id c
onte
nt (%
)
Major Fatty acids
Control Grinding with liquid N2Bead vortexing Osmotic shockWaterbath SonicationShake Mill
62
Chapter 3
Figure. 3.5b. Effect of different cell disruption methods on SFA, MUFAs and PUFAs
of Thraustochytrium sp. AMCQS5-5.
In Thraustochytrium sp. AMCQS5-5, palmitic acid (31.6%) and docosahexaenoic acid
(31.5%) were the major fatty acids (Figure 3.5a), with other polyunsaturated fatty acids
(C20:4n6, C20:5n3, C22:5n6 and C22:5n3) ranging from 2-9%, when cells were
disrupted using osmotic shock. Saturated, monounsaturated and polyunsaturated fatty
acid contents were 35.5%, 7.4% and 51%, respectively (Figure 3.5b). The highest
polyunsaturated fatty acid percentages were extracted by grinding and sonication
methods, 67% and 57%, respectively. Only shake mill resulted in a significant
percentage of monounsaturated fatty acids extraction (37.5%). Most methods resulted
in different lipid yields, but no difference in the fatty acid profiles for extraction from
Thraustochytrium sp. AMCQS5-5 (Figure 3.5b). If the omega-3 fatty acids could be
separated from other fatty acids in the Thraustochytrium sp. AMCQS5-5 extract, then
these fatty acids, particularly DHA, could be used as nutritional products and offset
the cost of biofuel production in this strain (Ryckebosch et al., 2012a; Singh et al.,
2015b).
3.3.5. Prediction of biodiesel properties
We analysed fatty acid profiles of two different thraustochytrid strains to
determine the suitability of these microalgae for biodiesel production. The fatty acid
0
10
20
30
40
50
60
70
80
SFAs MUFAs PUFAs
Tota
l lip
id c
onte
nt (%
)
Fatty acid composition
Control Grinding with liquid N2Bead vortexing Osmotic shockWaterbath SonicationShake Mill
63
Chapter 3 values were taken as an input in predicting the biodiesel properties by using an open
access software "Biodiesel Analyzer© Ver. 1.1" (Talebi et al., 2013). Some of the
important parameters of biodiesel are cetane number (CN), Iodine value (IV) and
oxidative stability (OS), which determine the combustion behavior, quality of
biodiesel, stability and performance, respectively (Islam et al., 2013). The CN and IV
values in Schizochytrium sp. S31 (47.4 and 98.61) when compared to
Thraustochytrium sp. AMCQS5-5 (44.01 and 157.44) were observed. According to
ASTM D6751-12, the standard values were 47-51 (CN) and 120 g I2/100g maximum,
indicating suitability of selected strain S31 for biodiesel, however, its further
characterization will be a follow up study. The OS value of 6.5 for Schizochytrium sp.
S31 was higher than 1.65 for Thraustochytrium sp. AMCQS5-5 suggesting oxidation
stability decreased with the increase of polyunsaturated fatty acid content. Other
properties such as saturated fatty acids (SFA), monounsaturated fatty acids (MUFA),
polyunsaturated fatty acids (PUFA), degree of unsaturation (DU), saponification value
(SV), long chain saturated fatty (LCDF), cold filter plugging point (CFPP), cloud point
(CP), allylic position equivalent (APE), bis-allylic position equivalent (BAPE),
Table 3.4. Predicted biodiesel properties of given thraustochytrids strains.
Properties Units Strain S31 Strain AMCQS5-
5
Saturated fatty
acid
% (m/m) 59 44
Monounsaturated
fatty acids
% (m/m) 35 4.6
Polyunsaturated
fatty acids
% (m/m) 14 33
Degree of
unsaturation
63 70
Saponification
value
mg KOH/g oil 233.98 164.73
Iodine value g I2/100 g 98.61 157.44
64
Chapter 3 Cetane number min 47.44 44.01
Long chain
saturated factor
% (m/m) 3.8 5.85
Cold filter plugging
point
oC 4.54 1.90
Cloud point oC 9.74 10.79
Allylic position
equivalents
83 167.60
Bis-Allylic position
equivalents
70 165.20
Oxidation stability h 6.5 1.65
Higher heating
value
oC 42.14 32.33
Kinematic
Viscosity
mm2/s 1.28 0.94
Density Kg/m3 0.94 0.73
3.3.6. Energy analysis
Energy consumption of the investigated cell disruption methods was carried
out to establish their potential as large scale processes. The comparative estimated
energy consumptions and processing times of the investigated cell disruption methods
are presented in Table 5. In comparison, water bath and sonication methods resulted
in highest energy consumption, 2400 MJ Kg-1 dry mass and 1200 MJ Kg-1 dry mass,
respectively. Shake mill energy consumption was estimated to be 690 MJ Kg-1 dry
mass. The osmotic shock method consumed modest energy (4.8 MJ Kg-1) and highest
lipid recovery, and so was the preferred method. NaCl present in the lysate solution
exerted osmotic pressure which was estimated to be 4.21 KPa, using an online osmotic
pressure calculator. Another study showed that the osmotic pressure of 1.9 KPa was
enough to break the microbial cells (Kar & Singhal, 2015). It has been demonstrated
that energy consumption for the microwave (28 MJ Kg-1) and ultrasound (44 MJ Kg-
1) methods enhanced lipid extraction from Chlorella sp. (Olofsson et al., 2014), which
shows that osmotic shock consumed less energy than either of these methods.
65
Chapter 3 Table 3.5. Energy consumption comparison.
Cell disruption methods Lipid yield
(%)
Energy consumption
(MJ Kg-1 dry mass)
Processing time
(min)
Control 22.04 Nila 0
Manual grinding 44.6 ND b 2
Bead vortexing 22.8 48 20
Osmotic shock 48.7 4.8 c 2
Water bath 20.8 2400 20
Sonication 31.05 1200 20
Shake mill 30.5 690 5
Thraustochytrids mass concentration of 16.6 kg/ m-3 was used for energy analysis.
Where Nila represents no energy was consumed; NDb represents physical effort cannot
be quantified; cOsmotic pressure by virtue of salt addition
3.4. Conclusion
In this study, we have shown that some cell disruption methods, particularly
osmotic shock, result in both different oil yields and variation in the percentage of
saturated fatty acids, monounsaturated fatty acids and polyunsaturated fatty acids in
the extracted oil from thraustochytrid species.
66
Chapter 4
Chapter 4: Optimised cell disruption methods for lipase
extraction from Australian thraustochytrids
67
Chapter 4 4.1. Introduction
The marine environment has a unique biodiversity and is a good source of
enzymes with unusual properties with potenial utility in biotechnology (Lee et al.,
2010a). Considerable effort has gone into screening marine extracts for novel bioactive
compounds, with unique structural classes being discovered (Kurtovic & Marshall,
2013; Vester et al., 2014). Marine microorganisms are particularly useful for bioactive
compunds discovery since they can be fermented and so offer ease of scaleup versus
other sources (Sana, 2015). Among marine microbes, thraustochytrids are one of the
most scaleable strains since they are commerically useful for the production of omega-
3 oils and carotenoids. These heterotrophic microorganisms also are a potentially
useful source of new enzymes (Gupta et al., 2012a; Gupta et al., 2013).
Thraustochytrids can accumulate approximately 40% docosahexaeonic acid (DHA) as
a percentage of total lipids (Liu et al., 2014b). Thraustochytrids have been used as
animal feed, aquaculture feed and poultry feed in addition to their use in omega-3 and
carotenoid production (Milledge, 2012; Sprague et al., 2015). Thraustochytrids have
been reported to produce degradative enzymes such as lipases, peptidases,
phosphatases and cellulases, although the utility of their enzymes has not been fully
investigated (Bongiorni et al., 2005; Kanchana et al., 2011).
Cell disruption is an important downstream processing step as it impacts
extraction yields and therefore the cost of bioactive compound production (Lee et al.,
2012c). Various methods–physical, chemical and mechanical–are available for
disrupting microbial cells, with mechanical methods generally used for large-scale
processing (Günerken et al., 2015). The selection of a suitable method of cell
disruption for extracting intracellular products depends on cell wall strength,
intracellular location of products, stability and the final use of the recovered products
(Klimek-Ochab et al., 2011). For example, the effect of chemical and mechanical cell
disruption methods was investigated for extraction of intracellular -D-galactosidase
(Puri et al., 2010).
Lipases are hydrolytic enzymes that catalyse the hydrolysis or synthesis of esters
and possess positional as well as fatty acid selectivity (Ferreira-Dias et al., 2013; Singh
& Mukhopadhyay, 2012). Lipases isolated from marine organisms generally showed
maximum activity at low temperatures and have unusual fatty acid specificity (Vester
et al., 2014). Lie and Lambertsen, observed that the crude enzyme mixture collected
68
Chapter 4 from cod intestine showed an unusual fatty acid selectivity, preferentially hydrolysed
omega-3 fatty acids (Lie & Lambertsen, 1985). Similar fatty acid selectivity has also
been observed from the crude digestive juice from salmon and rainbow trout collected
from the pyloric area of the stomach (Halldorsson et al., 2004). However, in each of
these cases no omega-3 lipase has been characterised. A recent study reported that a
novel lipase isolated from the marine microbe Marinobacter lipolyticus SM19 showed
a high preference for hydrolysing eicosapentaenoic acid (EPA) from fish oil (Pérez et
al., 2011).
We report here the development of a cell disruption method for release of
intracellular lipases from thraustochytrids. The efficiency of different cell disruption
methods for the release of intracellular proteins and their effect on lipolytic activity
was investigated. To the best of our knowledge this is the first report on optimisation
of cell disruption methods for lipase extraction from thraustochytrids.
4.2. Materials and methods
4.2.1. Chemicals
All chemicals such as p-nitrophenyl palmitate (p-NPP), Tween 80, and medium
components (yeast extract and peptone) were procured from Sigma-Aldrich, USA.
Acid washed glass beads from Thomas Scientific, Australia and Sea salts from Instant
Ocean, USA. The API ZYM kit was procured from BioMérieux, Australia Pty. Ltd.
4.2.2. Strain selection and cultivation
Recently isolated strains designated as AMCQS5-3 (Genbank accession
numbers JX993839), AMCQS5-4 (accession numbers JX993840), AMCQS5-5
(accession numbers JX993841), AMCQS1-9 (accession numbers JX993843) and
AMCQ-4b27 (accession numbers JX993842) were used for screening lipase activity
(Gupta et al., 2013). Two microbial cultures, Schizochytrium S31 (S31) and PRA 296,
were procured from the American type of culture collection (ATCC).
Thraustochytrids were maintained in a GYP medium containing glucose (5 g L-
1) , peptone (2 g L-1) and yeast extract (2 g L-1) and artificial sea water (50% v/v) at
20 oC. Stock cultures were subcultured onto fresh plates at regular intervals. All
thraustochytrids were grown in modified Vishniac’s Medium (0.1% glucose, 0.01%
yeast extract, 0.01% peptone, 0.1% gelatine hydrolysate, 1.2% agar in seawater, pH
69
Chapter 4 7.0) for enzyme screening (Raghukumar, 2002). The incubation conditions were 20 oC
and 150 rpm agitation until the cells reach the late exponential growth phase (96 h).
Before harvesting, cells were observed under a microscope for growth.
4.2.3. Enzymatic profiling of isolates
The thraustochytrid cells were harvested at 5000 rpm for 15 min and washed
with sterile artificial sea water. Cells were suspended in artificial sea water and 65 μl
of sub aliquots (corresponds to 4.0 x 105 cells) inoculated into (19 microcupules except
in control) API ZYM strips and incubated at 20 oC overnight. After incubation, 30 μl
of API ZYM A and B were added to the microcupules and incubated for 5 min.
Enzyme activities were measured using a colorimetric scale (Tiquia, 2002).
4.2.4. Lipase production
Thraustochytrid were grown in medium containing peptone (500 mg L-1), yeast extract
(500 mg L-1) and Tween 80 (1% v/v) (Arafiles et al., 2011). The production medium
was pre-adjusted to pH 6.0 using 0.1 M HCl. A volume of 50 ml medium contained
in a 250 ml Erlenmeyer flask was inoculated with a loop full of culture from the plate.
The flask was incubated at 20 oC with shaking at 150 rpm for 48 h. Inoculum (5% v/v)
was used for the production of biomass. The production culture was incubated for 120
h under the same conditions. The biomass was harvested by centrifugation (5000 rpm,
10 min). Harvested biomass was resuspended in distilled water to remove the Tween
80, and this process was repeated three times (Li et al., 2001). The biomass was kept
at -80 oC overnight and freeze-dried (Christ, Germany) for 24 h for further use.
4.2.5. Cell disruption methods for lipase release
4.2.5.1. Bead vortexing
A sample of freeze-dried biomass (100 mg) was suspended in 1 ml of
extraction buffer (100 mM Tris pH 7.2) with EDTA (10 mM) and NaCl (100 mM).
One ml of acid washed glass beads (425-600 μm, Klausen Pty Ltd, Australia) were
added and the contents were vortexed for 10 min, in 30 second burts. Ground biomass
was centrifuged at 12,000 rpm for 60 min at 4 oC. The supernatant was analysed for
lipase activity and protein levels as described below.
70
Chapter 4 4.2.5.2. Grinding with liquid nitrogen
A sample of freeze-dried biomass (100 mg) was suspended in 10-15 ml of
liquid nitrogen and ground with a pestle for 2 min. 1 ml of extraction buffer was added
and the resultant biomass was collected and centrifuged at 12,000 rpm for 60 min at
4 oC. The supernatant was analysed for lipase activity.
4.2.5.3. Sonication
A sample of freeze-dried biomass (100 mg) was suspended in 1 ml of
extraction buffer. The suspension was sonicated at 20 kHz, 40% amplitude and the
pulse was 40 seconds on 20 seconds off through a time-range of 5-25 min (Sonics,
USA). After every 5 min, cells were observed under a microscope to check the level
of disruption. During sonication, the sampling tubes were kept in ice-bath to avoid
heat-mediated denaturation of crude enzyme. The suspension was centrifuged at
12,000 rpm for 60 min at 4 oC. The supernatant was analysed for lipase activity.
4.2.5.4. Homogenization
Homogenization was performed using a rotor-stator type homogenizer
(Unidrive 1000, CAT Scientific, USA). A sample of freeze-dried biomass (100 mg)
was suspended in 1 ml of extraction buffer and the suspension was subjected to
homogenization at 8500 rpm/min for 5-25 min at 5 min intervals. After every 5 min,
the suspension was observed under the microscope. The suspension was centrifuged
at 12,000 rpm for 60 min at 4 oC. The supernatant was analysed for protein
concentration and lipase activity.
4.2.6. Microscopic observation of the native and disrupted thraustochytrid cells
The cell slurries were observed under a microscope using differential
interference contrast (Axio-imager, Zeiss, Germany) to check for disruption.
Microbial smears of normal and disrupted cells were prepared on the glass slide and
air-dried
4.2.7. Lipase and protein assay
Lipase activity was measured from the supernatant obtained after the cell
disruption using p-nitrophenol palmitate pNPP as a substrate, following a previously
described method with some modifications (Winkler & Stuckmann, 1979). The lipase
substrate solution was prepared by adding solution A consisting of 30 mg of pNPP in
10 ml of isopropanol and the solution B consisting of 0.4% Triton X-100, 0.1% Gum 71
Chapter 4 Arabic and 50 mM Tris-HCl buffer, pH 7.0. Solution A (10 ml) was added to solution
B (90 ml) drop by drop with stirring. The substrate solution (2.9 ml) and 0.1 ml of
enzyme solution was incubated at 37 oC for 10 min and absorbance was measured at
410 nm. One unit (IU) of lipase activity was defined as one nmol of pNPP released per
minute under the standard assay conditions.
Protein concentration was estimated from the supernatant using the Bradford
method with bovine serum albumin as the standard (Bradford, 1976). All the enzyme
and protein assays were done in four replicates and the standard error (SE) calculated
using Microsoft excel software.
4.3. Results and discussion
In this study, seven thraustochytrids isolates were screened for 19 different
enzyme activities. An API ZYM kit was used that contained enzyme strips that
consisted of 20 microcupules with 19 different chromogenic substrates and one
control. Table 4.1 shows the wide spectrum of enzyme activities that were exhibited
by the thraustochytrids isolated from Austrlian marine water. The new isolates were
isolated form two different sample collection trips. The investigated thraustochytrid
strains exhibited high esterase, lipase, acid and alkaline phosphatase activities. The
peptide degrading enzyme leucine arylamidase was produced by all strains, except
AMCQS5-4. Maximum lipase activity was observed in AMCQS1-9. Other peptidases
such as cysteine arylamidase and trypsin were produced in all isolates, except
AMCQS5-4 and AMCQS1-9. PRA 296 and AMCQS1- -chymotrypsin
-galactosidase -glucosidase activities were observed in the strains
PRA 296 and AMCQS5-5.
72
Chapter 4 *Table 4.1a. Enzymatic activities of thraustochytrids isolated form Australian marine
waters (First collection trip, October 2011).
Enzyme S31 PRA GTC 7 6a13 S53 S54 4b27 S19 S55Alkaline Phosphatase
LE HE
Esterase HE
Esterase + Lipase
Lipase
Leucine arylamidaseValine arylamidase
Cysteine arylamidaseTrypsin
a-chymotrypsin
Acid phosphatase
Napthol-AS-BI-phosphohydrolase
a-galactosidase
b-galactosidase
b-glucuronidase
a-glucosidase
b-glucosidase
N-acetyl-b-glucosaminidase
a-mannosidase
a-fucosidase
(Indicate what is shown in red, green and straw yellow colour)
*Samples were collected by Dr A Gupta and Dr T Thyagrajan from Barwon region,
Victoria, Australia.
73
Chapter 4 **Table 4.1b. Enzymatic activities of thraustochytrids isolated form Australian marine
waters (Second collection trip, September 2013).
Enzymes PP31 AGT-0
AGT-8
TS-1
TS-7
TS-8
TS-4
DSP5-1
DSP 2-4
DSP1-1B
DSP 2-3
AVI-1
AVI-2
Alkaline PhosphataseEsteraseEsterase + LipaseLipaseLeucine arylamidaseValine arylamidaseCysteine arylamidaseTrypsina-chymotrypsinAcid phosphataseNapthol-AS-BI-phosphohydrolase
-galactosidase-galactosidase-glucuronidase-glucosidase-glucosidase
N-acetyl-b-glucosaminidase
-mannosidase-fucosidase
Footnote: Semi-quantitative screening of thraustochytrid for different enzymatic
activities was done using a commercial enzyme kit. Thraustochytrid cells were
inoculated into the strips and observed for colour formation due to the enzymatic
action. (Green) high activity, (yellow) medium activity; (red) low activity; (no colour)
no activity. GenBank accession numbers for isolates given in Appendix.
Cluster analysis revealed that the enzymatic activities of thraustochytrid strains
were homogenous, with 86% similarity (Fig. 4.1). Bongiorni et al. also observed a
range of enzyme activity in some thraustochytrids (Bongiorni et al., 2005).
**Others bioprocessing lab members Dr A Gupta, Dr T Thyagrajan, Dr D Singh and
Dr S Sonkar were involved in the collection of various isolates from the Queenscliff
region, Victoria.
74
Chapter 4
Figure. 4.1. Cluster analysis of thraustochytrids based on various enzyme activities.
Cluster analysis was done using XLSTAT version and reveals that degree of similarity
was 86% based on presence/absence in their enzymatic profiles.
Lipase production was inhibited by high concentrations of glucose in the
medium (Kanchana et al., 2011). To improve lipase production levels, thraustochytrids
were grown in a Tween 80 containing medium. Intracellular lipase activity was
measured using a colorimetric enzyme assay. The lipase activity from S31 (39 IU/g)
was higher than that observed for the other strains used in the current study. The
maximum lipase activities observed for the thraustochytrid isolates were 34 IU/g for
AMCQ-4b27 and 36 IU/g for AMCQS1-9 (Table 4.1). 18r DNA sequence analysis
identified these strains as a Thraustochytrium sp (AMCQ-4b27) and a Schizochytrium
sp (AMCQS1-9) (Gupta et al., 2013).
75
Chapter 4 Table 4.1. Lipase activity of thraustochytrids new isolates and ATCC cultures. Lipase
activity was determined by p-NPP assay using whole cell as catalyst.
Thraustochytrid strains Lipase activity (IU/g)
In house isolates AMCQS5-3 26±2.0
AMCQS5-4 21±1.0
AMCQS5-5 32±2.0
AMCQS1-9 36±1.6
AMCQ-4b27 34±4.2
ATCC S31 39±3.7
PRA 296 28±2.0
4.3.1. Cell disruption for lipase extraction
Cell disruption is a key factor in the extraction of intracellular enzymes from
microbial cells. Different methods are available in the literature to solubilize the
intracellular proteins that depend on the location of enzymes, its application and
stability (Kurtovic & Marshall, 2013). The thraustochytrids strains examined had a
range of enzymes activities, with some lipase activity. We assessed the effect of cell
disruption techniques on lipase extraction from two thraustochytrid strains. The impact
of different cell disruption methods on enzyme yield varies with enzyme stability and
cell type, so that a balance between maximum cell degradation effectiveness and
minimum protein denaturing needs to be achieved.
The effect of cell disruption methods on lipase extraction from thraustochytrid
was studied via quantifying protein content released, lipase extraction and also by
direct microscopic observation (Fig. 4.2 a-d). Undisrupted cells of AMCQS1-9 and
AMCQ-4b27 remained intact (Fig. 4.2a and b), whereas, cells subjected to disruption
ruptured to release intracellular components (Fig. 4.2c and d).
76
Chapter 4
Figure. 4.2. Microscopic observation of thraustochytrid after cell disruption.
Undisrupted/Intact cells of AMCQS1-9 and AMCQ-4b27 (a and b), and disrupted cells
of AMCQS1-9 and AMCQ-4b27 (c and d) by sonication reaction conditions ( 20 KHz,
40% amplitude, 15 min) were observed after the treatment using differential
interference contrast.
4.3.1.1. Bead vortexing
Thraustochytrid cells AMCQ-4b27 and AMCQS1-9 were disrupted by bead
vortexing. The lipase activity and specific activity after cell disruption by bead
vortexing are shown in Fig. 4.3. More protein was obtained from AMCQ-4b27 cells
(130 μg/ml) than from AMCQS1-9 (63 μg/ml), however, lipase activity was greater in
protein extracted from AMCQS1-9 cells (0.045 IU/μg versus 0.125 IU/μg,
respectively). The effectiveness of bead vortexing versus other cell disruption methods
77
Chapter 4 appears to be cell type dependant. -galactosidase
from Kluyveromyces lactis by disruption with glass beads. They also observed the
volumetric and specific activity were minimum when compared with sonication and
other methods (Becerra et al., 2001).
AMCQ-4b27 AMCQS1-9
6.0
6.5
7.0
7.5
8.0
Acti
vity (
IU/m
l)
Thraustochytrids
Activity Protein yield Specific activity
60
70
80
90
100
110
120
130
140
Prot
ein yi
eld (μ
g/ml)
0.04
0.06
0.08
0.10
0.12
0.14
Spe
cific
activ
ity (I
U/μg
)
Figure. 4.3. Effect of bead vortexing on thraustochytrid cell disruption and lipase
activity. Bead vortexing of thraustochytrid cells resulted in more protein release from
AMCQ-4b27 than AMCQS1-9. S1-9 cells were relatively more susceptible for high
enzyme activity than AMCQ-4b27 cells (in all case SE <10%).
4.3.1.2. Grinding with liquid nitrogen
Grinding with liquid nitrogen is a simple and quick method for disrupting
microbial cells (Zheng et al., 2011). Grinding with liquid nitrogen was faster than other
methods for cell disruption. The total protein extracted by this method was higher than
that obtained by other methods, for AMCQ-4b27 (156 μg/ml) and AMCQS1-9 (107
μg/ml). Fig. 4.4 showed that grinding with liquid nitrogen led to a higher degree of
cell disruption, consistent with higher protein yields. Lipase activity of AMCQ-4b27
(25 IU/ml) was higher than that for AMCQS1-9 (18 IU/ml). However, specific activity
from AMCQ-4b27 (0.160 IU/μg) was lower than that for AMCQS1-9 (0.168 IU/μg).
Lipase from AMCQ-4b27 more readily degraded with temperature than lipase from
AMCQS1-9. Our results are consistent with the study investigated the effect of
different chemical and physical methods for extraction of -D-galactosidase, where
78
Chapter 4 grinding with river sand is effective in enzyme extraction compared with other
methods (Puri et al., 2010).
AMCQ-4b27 AMCQS1-917
18
19
20
21
22
23
24
25
26
Acti
vity (
IU/m
l)
Activity Protein yield Specific activity
Thraustochytrids
100
110
120
130
140
150
160
Prote
in yie
ld (μ
g/ml)
0.160
0.162
0.164
0.166
0.168
Spec
ific a
ctivit
y (IU
/μg)
Figure. 4.4. Effect of grinding on thraustochytrid cell disruption and lipase activity.
AMCQS-4b27 cell were more susceptible to cell disruption compared with AMCQS1-
9. Grinding with liquid nitrogen resulted in more protein released than did other
methods.
4.3.1.3. Sonication
Sonication was previously shown to be an effective method for releasing
intracellular bioactive compounds (Liu et al., 2013). In order to understand the optimal
sonication time for maximum protein release and lipase activity, we varied the
sonication treatment time. The impact of sonication exposure time on thraustochytrid
cells was investigated. Enzyme released by sonication can be calculated as:
R = Rm (1- e –kt)
Where R and Rm are the released enzyme activity and maximum enzyme
activity that can be released, respectively (U/mg/min), k is the disruption constant for
sonication and t is the time of sonication in minutes (Doulah, 1977). Lipase release
from thraustochytrid cells was investigated at an acoustic power of 20 kHz and 40 %
amplitude for 5 to 25 mins. Figure 4.5 [(a) and (b)] shows higher levels of protein
extraction as sonication time increased for thraustochytrids AMCQ-4b27 and
AMCQS1-9 cells. However, lipase activity increased to 15 minutes but plateaued after
that, so that the lipase concentration in extracted protein decreased after 15 minutes 79
Chapter 4 extraction (Figure 4.5). The maximum lipase activity obtained from AMCQ-4b27 was
47 U/ml and from AMCQS1-9 was 42 U/ml. This indicates that either heat produced
during sonication may have denatured lipase, or that lipase extracted more readily than
other protein (Farkade et al., 2006). Decreased lipase activity when acoustic power
was increased beyond 40% amplitude is consistent with heat-induced degradation. A
recent study indicated deactivation of Yarrowia lipolytica lipase upon increasing
sonication time (Kapturowska et al., 2012).
Fig. 4.5 (b)
5 10 15 20 2520
25
30
35
40
45
50
Acti
vity (
IU/m
l)
Activity Protein yield Specific activity
Sonication time (min)
48
50
52
54
56
58
60
62
64
66
Pro
tein y
ield (μg
/ml)
0.5
0.6
0.7
0.8
0.9
Spe
cific
activ
ity (I
U/μg
)Fig. 4.5 (b)
5 10 15 20 250
5
10
15
20
25
30
35
40
45
Acti
vity (
IU/m
l)
Activity Protein yield Specific activity
Sonication time (min)
20
30
40
50
60
Prot
ein yi
eld (μ
g/ml)
0.0
0.2
0.4
0.6
0.8
1.0
1.2
1.4
1.6
Spe
cific
activ
ity (I
U/μg
)
80
Chapter 4 Figure. 4.5. Effect of sonication on thraustochytrid cell disruption and lipase activity.
Time course effect of sonication on the protein release and lipase activity from
AMCQ-4b27 and AMCQS1-9.
4.3.1.4. Homogenization
During homogenization, microbial cells enter into a rotor-stator creating shear
force that disrupts the cells (Lee et al., 2012c). After 25 min homogenization 133 μg/ml
protein was extracted from AMCQ-4b27 cells and 75 μg/ml protein from AMCQS1-9
cells. As expected, protein content increased with increasing number of cycles (Halim
et al., 2012b). Lipase activity increased substantially in AMCQ-4b27 (25 IU/ml) and
AMCQS1-9 (37 IU/ml) until 20 min of homogenization and started declining
thereafter [Fig. 4.6 (a) and (b)], likely due to heat deactivation of the lipases. The
decrease in lipase activity after 20 min of homogenization can be attributed to enzyme
inactivation from the heat produced during the process. Overall lipase activity was
relatively low when compared to sonication, indicating that the combination of shear
forces and surface tension during disruption process increases lipase degradation (Lee
Ying Yeng et al., 2013; Taubert et al., 2000).
Fig. 4.6 (a)
5 10 15 20 25
10
12
14
16
18
20
22
24
26
Acti
vity (
IU/m
l)
Activity Protein yield Specific activity
Homogenization time (min)
40
60
80
100
120
140
Prote
in yie
ld (μ
g/ml)
0.16
0.17
0.18
0.19
0.20
0.21
0.22
Spec
ific a
ctivit
y (IU
/μg)
Fig. 4.6 (b)
81
Chapter 4
5 10 15 20 2510
15
20
25
30
35
40
Acti
vity (
IU/m
l)
Activity Protein yield Specific activity
Homogenization time (min)
30
40
50
60
70
80
Prote
in yie
ld (μ
g/ml)
0.46
0.48
0.50
0.52
0.54
0.56
Spec
ific a
ctivit
y (IU
/μg)
Figure. 4.6. Effect of homogenisation on thraustochytrid cell disruption and lipase
activity. Time course of extraction and its effect on lipase activity by homogenization.
4.3.2. Comparison of different cell disruption methods for lipase extraction
To release intracellular microbial products, such as lipases, efficient cell disruption
methods are required. However, these should not denaturation the compounds of
interest. In this study the four different cell disruption methods, bead vortexing,
grinding with liquid nitrogen, sonication and homogenization, were compared for
impact on protein and lipase recovery from thraustochytrid extraction (Table 4.2).
Sonication resulted in maximum lipase yield from AMCQ-4b27 (0.903 IU/μg) and
AMCQS1-9 (1.44 IU/μg). Lower specific activities were obtained using bead
vortexing (0.045 IU/μg and 0.125 IU/μg), grinding with liquid nitrogen (AMCQ-4b27
0.160 IU/μg and AMCQS1-9 0.168 IU/μg) and homogenization (AMCQ-4b27 0.211
IU/μg and AMCQS1-9 0.556 IU/μg). Extracts from strain AMCQS1-9 showed
approximately 1.5 times higher lipase activity than those from AMCQ-4b27. Overall,
sonication resulted in the highest lipase activity, followed by homogenization,
grinding with liquid nitrogen and bead vortexing.
82
Chapter 4 Table 4.2. Disruption of thraustochytrid cells for lipase extraction.
Cell disruption
method
AMCQ-4b27 AMCQS1-9
Lipase
activity
(IU/ml)
Protein
released
(μg/ml)
Lipase
activity
(IU/ml)
Protein
released
(μg/ml)
Bead vortexing 6±0.7 130±2.1 8±1.0 63±2.0
Grinding with liquid
nitrogen
25±3.0 156±2.4 18±0.5 107±1.6
Sonication 47±0.2 52±3.0 42±3.7 29±1.3
Homogenization 25±5.0 126±17 37±4.0 73±9.0
4.4. Conclusion
Thraustochytrids contain a range of potentially useful enzyme activities,
including lipase activity. Cell disruption methods were used to increase the extraction
yield of intracellular lipase from thraustochytrid cells. We conclude that AMCQ-4b27
cell were sensitive to protein release than the AMCQS1-9. Moreover, AMCQS1-9 has
yielded high total specific lipase activity than the AMCQ-4b27. Sonication resulted in
the higher lipase activity yield, with strain AMCQS1-9 yielding more lipase activity
than did strain AMCQ-4b27. Lipase activity yield did not completely correlate with
protein extraction yield, probably due to partial lipase degradation under conditions of
high temperature or sheer.
83
Chapter 5
Chapter 5: Tween 80 influences the production of
intracellular lipase by Schizochytrium S31 in a stirred tank
reactor
84
Chapter 5 5.1. Introduction
Lipases (triacylglycerol lipases, EC 3.1.1.3) belongs to the hydrolase family of
enzymes that catalyse the hydrolysis of triglycerides into free fatty acids and glycerol
(Schreck & Grunden, 2014). However, under certain conditions, they are also able to
catalyse synthetic reactions (Martins et al., 2014). In addition, lipases have some
important functional properties such as substrate selectivity, regio-selectivity and high
enantio-selectivity (Gupta et al., 2007; Yuan et al., 2014). These properties make them
potential catalyst in numerous industrial processes, such as food, detergent, chemical,
and pharmaceutical industries (Sangeetha et al., 2011; Yuan et al., 2014). Lipases are
ubiquitous enzymes widely distributed in plants, animals and microorganisms
(Mazzucotelli et al., 2015). Among them, microbial lipases have displayed a broad
range of substrate specificities. This property might have evolved through the access
of microbial lipases to different carbon sources (Hasan et al., 2006; Salihu & Alam,
2015). There is a growing demand for the novel lipases from a microbial sources,
because of their ease in production and access to genetic manipulations. Also because
they work under mild reaction conditions without requirement of any co-factors (Pérez
et al., 2011).
The health benefits of omega-3 fatty acids have been evaluated by a number of
clinical studies (Barrow, 2010; Nichols et al., 2014). There is strong evidence for the
efficacy of omega-3 fatty acids in the prevention of sudden death from cardiovascular
disease and rheumatologic conditions (Abuajah et al., 2015; Bhangle & Kolasinski,
2011; De Caterina, 2011). The anti-cancer activity of omega-3 fatty acids, particularly
against colorectal cancer, has recently been shown (Cockbain et al., 2012). Infant brain
and eye development is influenced by DHA, which has led to the addition of DHA to
the infant formulae. Learning capacity of school going children has been shown to
increase by taking DHA enriched food supplements (Qawasmi et al., 2013; Wu et al.,
2015). Oily marine fish are the primary source for the industrial production of omega-
3 fatty acids. These fish derived omega-3 fatty acids from their diet, primarily from
phytoplankton. Researchers have implemented different methods for concentrating
omega-3 fatty acids (Rubio-Rodríguez et al., 2010). A promising green approach for
concentration of omega-3 fatty acids is the use of lipases. Generally the enzymatic
methods are faster than other methods and can be carried out under milder conditions
then methods involving chromatographic separation, molecular distillation or urea
85
Chapter 5 complexation (Kahveci et al., 2010; Kralovec et al., 2010). Commercially available
lipases have been exploited for omega-3 fatty acid concentration (Akanbi et al., 2013;
Casas-Godoy et al., 2014; Fernández-Lorente et al., 2011; Sharma et al., 2013;
Valverde et al., 2013). However, selectivity of fatty acid hydrolysis is poor and no
know lipases selectivity hydrolyse DHA and EPA, a property that would be useful in
selectivity concentrating these fatty acids from fish or microbial oils.
Thraustochytrids produce lipases and because of their high productivity of
omega-3 fatty acids, particularly DHA, they are potential sources of omega-3 selective
lipases, although none have yet been characterised (Bongiorni et al., 2005; Kanchana
et al., 2011). We recently reported lipase activity from a strain of Schizochytrium
named S31 (Table 4.2). In this work we optimised culture conditions to enhance lipase
production from Schizochytrium sp. S31 in a bioreactor and optimised the extraction
of this lipase using sonication. Using a recently developed p-nitrophenyl esters (p-NP)
assay (Nalder et al., 2014) we showed that lipase extracted from S31 preferentially
hydrolysed DHA over EPA, an unusual selectivity for lipases.
5.2. Materials and methods
5.2.1. Microorganism and maintenance of culture
Schizochytrium S31 (ATCC 20888) was procured from the American type
culture collection (ATCC). The stock culture was maintained on GYP medium
consisting of glucose 5, yeast extract 2, mycological peptone 2, agar 10 g L-1 and
artificial seawater 50% at 25 °C and sub-cultured after 15 days. All medium
components and chemicals including p-NP were obtained from Sigma, Australia. The
p-NP fatty acid esters of -linolenic acid (C18:3),
eicosapentaenoic acid (C20:5) and docosahexaenoic acid (C22:6) were synthesised
based on a recently published protocol (Nalder et al., 2014).
5.2.2. Schizochytrium S31 growth and lipase production
Schizochytrium S31 was grown in a medium containing peptone 0.5, yeast
extract 0.5 g L-1 and Tween 80 (1% w/v; as sole carbon source). 50 ml of medium in a
250 ml Erlenmeyer flask was inoculated with a loop full of culture from the plates.
This culture flask was incubated for 48 h at 20 oC and shaken at 150 rpm. A 48 h old
5% seed culture (v/v) was used to inoculate sterilised production medium (100 ml in
86
Chapter 5 500 ml Erlenmeyer flask) containing peptone 0.5, yeast extract 0.5 g L-1 and Tween 80
for the production of lipase. The flasks were incubated in a rotary shaker for up to 9
days. Samples were withdrawn aseptically at regular time intervals and then dried and
weighted and lipase activity determined. For dry weight determination, the entire
content of the liquid culture was transferred to a pre-weighed centrifuge tube and
harvested by centrifugation at 3500 rpm for 10 min and the supernatant was discarded.
Harvested cells were washed thoroughly with phosphate buffer (pH 7.0) and then
freeze-dried for 24 h. The biomass weight was determined gravimetrically. Tween 80
concentrations were estimated based on a colorimetric method (Cheng et al., 2011b).
A standard curve was prepared by varying the Tween 80 concentration and the
unknown sample concentration was determined using the standard curve. Temperature
(15 to 30 oC) and initial pH (5-8) of the medium were optimised to maximise lipase
activity. All experiments were carried out in triplicates.
Fermentation of thraustochytrids was carried out in a 2 L stirred tank reactor
(New Brunswick, USA) with a working volume of 1.4 L at 25 oC with aeration of 0.5
vvm and agitation of 150 rpm. Varying concentration of Tween 80 (0.5 to 1.5%, v/v)
was used to optimise biomass, lipase and lipid production in the bioreactor. Samples
were withdrawn at regular time intervals and analysed for residual substrate. Changes
in pH were also monitored during the course of fermentation.
5.2.3. Estimation of Schizochytrium S31 growth
The growth of the organism was estimated by taking 1-ml aliquot periodically
and measuring the optical density (OD) at 600 nm in a UV-visible spectrophotometer
(Shimadzu1601, Japan) with medium as a blank. The OD value was converted into
cell mass concentration (mg ) using a standard curve.
5.2.4. Optimisation of cell disruption through sonication
Cell disruption of Schizochytrium S31 biomass was performed as previously
described (Byreddy et al., 2015). Thraustochytrid cell biomass (50 mg) was suspended
in 3 ml of solvent in a 15 ml centrifuge tube. Sample was sonicated at 20 kHz, 40%
amplitude and the pulse was 40 sec on and 20 sec off with total working time of 20
min (Sonics, Newtown, CT, USA). Sample tubes were kept on ice during the
sonication process to prevent overheating. Sonication conditions such as time and
87
Chapter 5 acoustic power were optimized. The sonicator used in this study was a horn type with
500 W power output and 20 KHz.
5.2.5. Lipase assay and protein estimation
Lipase activity was measured in the supernatant obtained after cell disruption
using p-nitrophenol palmitate (pNPP) as a substrate, following a previously described
method with some modifications (Winkler & Stuckmann, 1979). The lipase substrate
solution was prepared by adding solution A consisting of 30 mg of pNPP in 10 ml of
isopropanol and the solution B consisting of 0.4% Triton X-100, 0.1% Gum Arabic
and 50 mM Tris-HCl buffer, pH 7.0. Solution A (10 ml) was added to solution B (90
ml) drop by drop with stirring. The substrate solution (2.9 ml) and 0.1 ml of enzyme
solution was incubated at 37 oC for 10 min and absorbance was measured at 410 nm.
One unit (IU) of lipase activity was defined as one nmol of p-nitrophenol released per
minute under the standard assay conditions. The relative activity of crude enzyme is
calculated as a percentage activity divided by the most hydrolysed substrate
(designated 100% for each substrate).
Protein concentration was estimated from the supernatant using the Bradford
method with bovine serum albumin as the standard (Bradford, 1976). All enzyme and
protein assays were done in four replicates and the standard error (SE) calculated using
Microsoft excel software.
5.2.6. Lipid extraction
Lipid was extracted and quantified from freeze-dried biomass every 48 h
according to a previously published protocol (Lewis et al., 2000) with some
modifications. Freeze-
and methanol in 2:1 ratio) and the sample vortexed for 2 min, followed by
centrifugation at 13,000 rpm for 15 min. This extraction process was repeated three
times. The supernatants were collected in a pre-weighed vials and dried in an oven at
50 °C. The lipid percentage was measured gravimetrically. All the experiments were
performed in triplicate.
5.3. Results and discussion
5.3.1. Optimisation of culture conditions for lipase production
88
Chapter 5
Lipase production is influenced by the type of carbon source used, temperature
and initial pH of the culture medium (Orlando Beys Silva et al., 2005). Tween 80 has
been successfully used as a carbon source for the production of lipases from various
microbes (BoekeMa et al., 2007; Li et al., 2001; Sharma et al., 2014). In this study,
Tween 80 was used as the sole carbon source and its concentration was optimised to
maximise biomass and lipase productivity (Fig. 5.1). Tween 80 (1%) resulted in the
highest level of biomass (0.9 g L-1) and lipase (39 IU/g) productivity by
Schizochytrium S31. Similar results were found previously for Acinetobacter
radioresistens (Li et al., 2001). Lipase production did not improve when higher tween
80 concentration (1.5%) was used in the medium, although the reason for this is
unclear. Remaining experiments were performed with 1% tween 80.
Figure. 5.1. Effect of Tween 80 concentration on lipase production by Schizochytrium
S31 at shake flask, pH 6.0 and incubation temperature 25 oC.
Initial pH and temperature are known to influence lipase production (Priji et
al., 2015). In Schizochytrium S31 maximal lipase activity was observed at pH 6 (38
U/g) for the pH range of 5 to 8. Similar results were reported for lipase production
from Thraustochytrium sp. AH-2 and TZ when the medium pH was pre-adjusted to 6
(Kanchana et al., 2011). The effect of incubation temperature on lipase production was
investigated for the four temperatures 15 oC, 20 oC, 25 oC and 30 oC at pH 6 and
agitation 150 rpm, with lipase levels being highest at 25 oC (data not shown). Our
00.10.20.30.40.50.60.70.80.91
05
1015202530354045
0 0.5 1 1.5 2
Bio
mas
s (g/
L)
Lip
ase
activ
ity (I
U/g
)
Tween 80 (%)
Biomass (g/L) lipase (IU/g)
89
Chapter 5 results are comparable with lipase production from Colletotrichum gloeosporioides
where maximum lipase activity at 25 oC (Balaji & Ebenezer, 2008).
5.3.2. Production of lipase in stirred tank reactor
Growth kinetics of Schizochytrium S31 was studied in terms of biomass
production, lipid accumulation in cells, and lipase production. After 48 h
acclimatisation in the medium, dry cell weight went from 0.16 g L-1 to 0.2 g L-1, with
the log phase beginning after this time, up to 120 h when growth slows and the
stationary phase begins (Fig. 5.2). Schizochytrium S31 cells grew well in a
fermentation medium containing Tween 80 (1%) as sole carbon source. Fig. 5.2 shows
the time course of the investigated parameters at different growth phases and also
showed that lipase and lipid production were growth associated. Biomass increased
with incubation time and was maximal (0.9 g) after 5 days of incubation. Lipase
production reached a maximum of 39 U/g in the late exponential phase, then the lipase
activity declined (10.7 U/g) up until 9 days of incubation, perhaps due to product
inhibition. Our results are in agreement with other researchers who observed the
maximum lipase activity of Yarrowia lipolytica KKP 379 in late exponential phase
(after 72 h) (Fabiszewska et al., 2014). Another study conducted by Kanchana et al,
(2011), found that the maximum lipase activity was observed in thraustochytrids
between 4-7 days of incubation (Kanchana et al., 2011). The pH and substrate
concentration of the medium decreased as the incubation time increasing, which may
be due to the hydrolytic activity of lipase on Tween 80 releasing oleic acid for
utilisation. In the bioreactor, lipase production increased with increasing biomass
concentration as a result of increasing Tween 80 concentration from 0.5 to 1 %. In
addition, improved control of environmental factors in the vessel resulted in increased
enzyme production, compared with that achieved in flasks.
90
Chapter 5
0 2 4 6 8 100.1
0.2
0.3
0.4
0.5
0.6
0.7
0.8
0.9
1.0
0 2 4 6 8 103.5
4.0
4.5
5.0
5.5
6.0
6.5
7.0
0 2 4 6 8 10
0
50
100
150
200
250
300
350
Biom
ass (
g/L)
Biomass
pH
Incubation time (day)
pH
0
10
20
30
40
50
60
70
80
90
100
Sub
strat
e de
pletio
n (%
)
Substrate depletion
0
10
20
30
40
Lipas
e Ac
tivity
(IU/
g)
Lipase
Lipid
(mg/
g)
Lipid
Figure. 5.2. Fermentation profile for the production of lipase by Schizochytrium S31
in a stirred tank reactor (Tween 80 containing medium was used as sole carbon source,
pH 6.0 and temperature 25 oC).
Figure. 5.3. Biomass, lipid and DHA production reached a maximum after five days
of incubation. After that the DHA percentage decreased more rapidly than total lipids.
To investigate lipid content and fatty acid composition, biomass was collected
and analysed at different time intervals up to 9 days. DHA content increased with total
0
20
40
60
80
100
120
0
5
10
15
20
25
30
35
1 2 3 4 5 6 7 8 9
Bio
mas
s (%
)
Lipi
d/D
HA
(%)
Incubation time(daya)
Lipid (%) DHA (%) Biomass (%)
91
Chapter 5 lipid content and reached a maximum of 5.4% on day 5. Biomass, lipid and DHA all
decrease after day 5, although DHA decreases more rapidly than does total lipid (Fig.
5.3). Lipid is used for energy and membrane synthesis during starvation (Thompson
Jr, 1996) and DHA is associated with many cellular functions. DHA appears to be used
preferentially in the late stages, which also indicates that lipase is acting on DHA
during this stage (Fig. 5.2 and 5.3). The fatty acid composition of oil from
Schizochytrium S31 grown in Tween 80 medium is given in Fig. 4, with growth in
glucose medium as a control. The oleate in Tween 80 was directly accumulated into
cells (Fig. 5.4). A similar pattern was observed in Thraustochytrium aureum ATCC
34304, where Tween 80 in the medium increased the oleic acid content in lipid (Taoka
et al., 2011). A corresponding decrease in the percentages of other fatty acids was
observed in Tween 80 medium (Fig. 5.4). There was also changes in the ratio of fatty
acids when Tween 80 medium was used, with relative decreases in C16 (26%), C18
(70%) relative to DHA.
Figure. 5.4. Fatty acid composition of Schizochytrium S31 was compared between
lipid extracted from biomass grown GYP medium and that from grown in Tween 80
medium.
5.3.3. Disruption of Schizochytrium S31 using sonication
0
10
20
30
40
50
60
70
80
90
C14:0Myristic
acid
C16:0Palmitic
acid
C16:1n7Palmitoleic
acid
C18:0stearic acid
C18:1n9cOleic acid
C20:5n3EPA
C22:5n6DPA
C22:6n3DHA
Tot
al fa
tty
acid
s (%
)
Fatty acids
Control GYP TW 80
92
Chapter 5
In order to investigate the fatty acid selectivity of lipase from Schizochytrium
S31, sonication was used to release the intracellular lipase. The effect of sonication on
Schizochytrium S31 was evaluated by microscopic observation and cells were shown
to be fully broken at high acoustic power. Table 5.1 shows lipase activity extracted
from thraustochytrid cells at an acoustic power of 20 kHz and 40 % amplitude for
intervals of 5 to 25 mins. Lipase activity increased to 15 minutes but plateaued after
that, indicating partial lipase inactivation with increased time over 15 minutes. Since
sonication releases intracellular enzymes but also degrades, optimising time and power
are important for maximising lipase extraction (Kadkhodaee & Povey, 2008; Mawson
et al., 2011; Nguyen & Le, 2013).
Table 5.1. Effect of sonication on protein release and lipase activity from
Schizochytrium S31.
Cell disruption
methods
Cell disruption
parameters
Lipase
activity
(IU/ml)
Protein
concentratio
n (μg/ml)
Specific
activity
(IU/μg)
Sonication 5 min
10 min
15 min
20 min
22.8±1.4
32.5±1.6
37.8±1.5
26.3±1.3
43±1.5
60±1.8
82±2.1
105±9
0.512
0.533
0.451
0.247
Fig. 5.5 (a) and (b) show the effect of sonication on lipase activity from
Schizochytrium S31 at different levels of sonication time and power. As the sonicator
power increased from 30% to 70%, the total proteins released increased from 53 to
122 μg/ml and the most lipase activity was observed at 50% power (64 IU/ml). Total
proteins released after sonication treatment was 3 folds higher at 50 % (0.75 IU/μg)
sonication power than 70% (0.25 IU/μg) (Fig. 5a). Our results are in agreement with
previous studies showing that the amount of protein release from microbial cells
increased linearly with increasing sonication time and power (Apar & Özbek, 2008;
Iida et al., 2008). The highest protein extraction levels from Schizochytrium was
observed after 15 min of sonication time at 70% of sonication power. However, both
total lipase activity and specific lipase activity decreased at 70% power, with extended
time causing additional lipase activity degradation. A recent study by Kapturowska et
93
Chapter 5 al, 2013, demonstrated the negative effects of extended sonication time on lipase
activity from Yarrowia lipolytica KKP 379 (Kapturowska et al., 2012).
00.10.20.30.40.50.60.70.8
0
20
40
60
80
100
120
140
30 50 70
Spec
ific
activ
ity
Lip
ae a
ctiv
ity/p
rote
in c
onc
Sonication power (%)
Lipase activity (IU/ml) Protein conc (μg/ml) Specific activity (IU/μg)
00.20.40.60.811.21.41.61.82
0
20
40
60
80
100
120
140
3 6 9
Spec
ific
activ
ity
Lip
ase
activ
ity/P
rote
in C
onc
Sonication Time (min)
Lipase activity (IU/ml) Protein conc (mg/ml) Specific activity (IU/mg)
a
b
94
Chapter 5 Figure. 5.5. Cell disruption optimisation of Schizochytrium S31 a) as a function of
sonication power, b) time duration. (Sonication power 70% for 10 mins was subjected
to 100 mg biomass for the lipase extraction).
5.3.4. Hydrolysis of p-Nitrophenyl esters
Understanding and controlling lipase fatty acid selectivity is important in the
modification of fats and oils for high value products and in the production of structured
lipids (Qin et al., 2014). The substrate selectivity of S31 crude lipase activity was
determined using the standard assay in the presence of 8 mM of the different p-NP
esters of various chain lengths: p-NP acetate (C2); p-NP butyrate (C4); p-NP caprylate
(C8); p-NP decanoate (C10), p-NP laurate (C12), p-NP myristate (C14), p-NP
palmitate (C16), p-NP stearate (C18), p-NP-EPA and p-NP-DHA. Lipase extracted
from Schizochytrium S31 showed a broad range of substrate activity (Fig. 6).
Maximum activity was observed towards short chain length fatty acids, followed by
medium chain length fatty acids. An unusual selectivity was observed towards p-NP-
DHA, when compared with p-NP-EPA. This type of unusual fatty acid selectivity was
previous observed for a fish digestive lipase, although this lipase was never
characterised (Halldorsson et al., 2004). However, a study investigated the fatty acid
selectivity of five commercial lipases using p-NP esters found that all lipase showed
preference to EPA over DHA (Nalder et al., 2014). This unusual selectivity might be
due to the requirement of the Schizochytrium to process DHA more than EPA, since
EPA is present in only trace amounts in the oil from this strain, whereas DHA levels
are relatively high, in contrast to most microalgae.
95
Chapter 5
Figure. 5.6. Substrate specificity of lipase from Schizochytrium S31 (p-nitrophenyl
esters of different chain lengths were incubated with crude enzyme).
5.4. Conclusion
Schizochytrium S31 was grown in a bioreactor using Tween 80 as the carbon
source, which resulted in high biomass and oleic acid levels. Lipase extraction by
sonication was optimised and the fatty acid selectivity of the extracted lipase
investigated using p-NP fatty acid esters. The lipase extract preferentially hydrolysed
shorter chain fatty acids, however, DHA was preferentially hydrolysed over EPA. This
preferential hydrolysis of DHA is unusual and may be industrially useful. This
selectivity may be a result of this Schizochytrium species having much higher levels
of DHA then EPA, and so DHA processing is more important for the organism growth
and function.
0
20
40
60
80
100
120
Rel
ativ
e ac
tivity
(%)
p-NP acyl esters
96
Chapter 6
Chapter 6: Bead milling for lipid recovery from
thraustochytrids cells
97
Chapter 6 6.1. Introduction
Marine fish and microalgal oils are rich sources of omega-3 fatty acids. The
health benefits of omega-3 oil supplements have been documented in numerous studies
(Ellulu et al., 2015). Methods available for the concentration of omega-3 fatty acids
from fish and microbial oils include urea complexation, chromatography, distillation,
low temperature crystallisation, supercritical fluid extraction and enzymatic methods
(Rubio-Rodríguez et al., 2010). The primary advantages of enzymatic methods are
operation under mild reaction condition and less or no use of chemicals which can
degrade the oxidatively sensitive structures.
Omega-3 fatty acid production from microalgae is potentially useful due to
their ability to accumulate up to 30-70% of their dry weight as lipid (Adarme-Vega et
al., 2014; Martins et al., 2013). Microalgae can be used to produce bulk ingredients for
applications in pharmaceuticals, cosmetics, nutraceuticals, animal feeds and functional
foods (Yen et al., 2013). Thraustochytrids are marine heterotrophic protist commonly
found in marine environments, and are classified under Labyrinthulomycetes (Gupta
et al., 2012). They have rapid growth rates and are capable of producing significant
amounts of PUFA, including DHA (Gupta et al., 2013). Thraustochytrids have recently
been identified as a potential microalgae for omega-3 fatty acid and carotenoids
production (Singh et al., 2015a). A range of studies have also demonstrated the
suitability of thraustochytrids for biodiesel production (Chen et al., 2015c; Gupta et
al., 2015; Gupta et al., 2013).
Specific compounds produced from microbial sources, such as lipids, proteins,
carotenoids and carbohydrates, must be extracted and purified using downstream
processes prior to their respective applications (Wijffels et al., 2010). Production of
intracellular metabolites from microalgae involves extensive processing steps
including microalgae cultivation, harvesting, extraction and product recovery. The
selection of suitable technologies depends on the characteristics of the algae and the
stability of the product (Guldhe et al., 2014; Pahl et al., 2013). Research is ongoing
into the development of improved downstream processing techniques for microalgae
lipid recovery, a step which is a substantial contributor to total costs for algal
production systems (Delrue et al., 2012). Following harvesting, cell disruption is an
important step for the extraction of microalgae produced compounds. Different cell
disruption methods have been applied to a range of microalgae, including
98
Chapter 6 ultrasonication, enzymatic treatment, high-pressure homogenisation, alkali treatment,
microwaves and pulsed electric field (Grimi et al., 2014; Safi et al., 2014a), however,
some of these methods have resulted in low extraction yields of the products of
interest.
Bead mill, a homogeniser used for the grinding of ceramics and paints, has
been used for the mild disruption of microorganisms for the release of proteins
(Schwenzfeier et al., 2011). Recent advancements in temperature control in microalgae
cell suspensions make bead milling a promising technique for microalgae component
recovery (Doucha & Lívanský, 2008; Günerken et al., 2015). The operating principle
of bead milling is based on rapid grinding of thick microbial cell suspensions in the
presence of beads. The operating mode can be either batch (recirculation) or
continuous (single passage through milling chamber) depending on the feedstock used.
Parameters which influence cell disruption efficiency include feedstock residence time
(suspension feed rate in continuous operation, or process time in batch operation)
agitator speed, agitator design, milling chamber design, biomass concentration, bead
diameter, bead density and bead filling (Doucha & Lívanský, 2008). It is hypothesized
that the suspended cells are disrupted in the bead collision zones by compaction or
shear forces with energy transfer from the beads to cells (Gunerken et al., 2015). Bead
milling has been successfully applied in Saccharomyces sp and Chlorella sp cell
disruption for bioactive recovery (Balasundaram & Pandit, 2001; Balasundaram et al.,
2012).
In this study, we optimised thraustochytrid cell disruption using bead mill for
maximising lipid extraction yields. The cells of Schizochytrium were spherical and
filled with numerous lipid bodies. The process was optimised for lipid yield, energy
consumption and processing time. The extracted oil was characterised and further
employed for enzymatic hydrolysis for the concentration of omega-3 fatty acids,
particularly DHA.
6.2. Materials and Methods
6.2.1. Chemicals
All chemicals used in this study were of analytical grade. Medium components
such as glucose, yeast extract, mycological peptone (Sigma-Aldrich, USA) and sea salt
(Instant Ocean, USA) were used for biomass production. Schizochytrium S31 (ATCC
20888) was procured from the American Type Culture Collection (ATCC) and used 99
Chapter 6 as a standard culture. Solvents such as chloroform, methanol, hexane (from Merck),
acetic acid and diethyl ether (from Fischer and Honeywell, Australia) were used in the
lipid class analysis and gas chromatography. TLC standards (from Nu-Chek Prep,
USA) were used to identify each lipid class.
6.2.2. Thraustochytrids biomass cultivation
Thraustochytrids used in this study were maintained on GYP medium
consisting of glucose 5, yeast extract 2, mycological peptone 2, agar 10 g L 1 and
artificial seawater 50%, at 25 °C and sub-cultured after 15 days. Two strains,
Schizochytrium sp. S31 (ATCC) and Schizochytrium sp. DT3 (an in-house isolate),
grown in glycerol media, showed high biomass and lipid productivity (Gupta et al.,
2013). Seed medium (1% glucose, 0.2% yeast extract, 0.2% mycological peptone,
ASW 50%, pH 6.5) and production medium (4% Glycerol, 1% yeast extract, 0.1%
mycological peptone, ASW 50%, pH 6.5) were autoclaved at 121 °C for 20 min. Seed
medium (50 ml) was inoculated from agar plates and grown for 2 days in a shake flask
at 25 °C, at 150 rpm. Inoculum (5%) was used to inoculate production medium and
incubated under the same conditions for 5 days. Biomass was harvested by
centrifugation (10,000 × g, 15 min) and washed twice with distilled water, then freeze-
dried for 24-48 h. Freeze-dried biomass was stored at -20 °C for further use.
6.2.3. Cell disruption by bead milling
The Dyno-Mill (bead mill; Model No. 130857) from Klausen Pvt. Ltd.,
Germany was used for cell disruption. This mill is well suited to applications in
research and development. The Dyno-mill grinding chamber is positioned horizontally
with a chamber volume 79.6 ml. Algal suspension is fed through a funnel mounted on
the grinding chamber. The beads are accelerated by a motor connected via a central
shaft, causing the grinding effect. The bead mill contained an integrated cooling
system to prevent overheating. A cooling jacket and cooling coil are integrated into
the grinding chamber and supplied by an external water bath to extract heat, and
maintain a constant temperature below 20 oC throughout processing. The cell
suspension residence time in the chamber is controlled via the pump speed. The beads
and product are separated at the end of the chamber by a sieve. Beads used in this study
were (0.4-0.6 and 0.8-1.0 mm) zirconium oxide, which are reported to exhibit
maximum efficiency in cell disruption (Lee et al., 2012a). As per manufacturer advice
100
Chapter 6 the beads were filled to 54% v/v in the chamber. Keeping these parameter stable, the
effect of biomass concentration, bead size, RPM and processing time were
investigated. In total, four different cell concentrations in distilled water were used;
1%, 3%, 5%, and 7%. Samples were collected at 2 min intervals up to 8 mins and
analysed for total lipid yield and fatty acid profiles.
6.2.4. Microscopic observation
In addition to using lipid yield as a parameter for validating the effectiveness
of bead mill treatment on thraustochytrid cells, the direct effect on the cells was
observed under a differential interference contrast microscope (Axio-imager, Zeiss,
Germany). Microbial smears were prepared on glass slides, and air dried prior to
observation.
6.2.5. Lipid extraction
Fatty acid extraction was performed in accordance with Gupta and co-workers
(Gupta et al., 2012c) with some modifications. 10 mg of freeze dried biomass was
blended with 0.6 ml of solvent. Biomass and solvent were vortexed and then
centrifuged at 4000 rpm for 15 min. Supernatant was place in pre-weighed glass vials
and solvent was evaporated under nitrogen. Lipid content (% dry weight basis) was
determined gravimetrically.
6.2.6. Lipase catalysed hydrolysis of thraustochytrid oil
Lipase-catalysed hydrolysis of oil was carried out using a modification
literature method (Kahveci & Xu, 2011). 500 mg of thraustochytrid oil, a known
amount of lipase (3000 U, EC 232-619-9, Candida rugosa; Sigma-Aldrich, St. Louis,
US) and 1.8 ml of Tris-HCl buffer (50 mM, pH 7.0) were taken in 25 ml round bottom
flasks and heated to 45 oC at 300 rpm stirring. Substrates were stirred for 30 min and
lipase was added to initiate hydrolysis. Samples form the reaction mixture were
withdrawn at specific time intervals over the course of the 1-4 h reaction. The enzyme
hydrolysis reaction was scaled up with 2 g of thraustochytrid oil with optimised
enzyme concentration. All experiments were conducted in triplicates with the mean
values reported.
101
Chapter 6
6.2.7. Thin Layer Chromatography for lipid class analysis
Both the hydrolysed and unhydrolysed portions of thraustochytrid oil were
analysed by capillary chromatography with a flame ionization detector (Iatroscan
MK5, Iatron Laboratories Inc., Tokyo, Japan). The Iatroscan settings were: air flow
rate; 200 ml/min, hydrogen flow rate; 160 ml/min and scan speed of 30 s/scan. Under
these conditions, the chromarods were cleaned by scanning twice before applying
samples. 1 μL of each lipid fraction in hexane was spotted onto the rods with the aid
of an auto pipette along the line of origin on the rod holder and developed for 22 min
in a solvent tank containing hexane/diethyl ether/acetic acid (60:17:0.2, vol/vol/vol).
TLC standards were used to identify each lipid class.
6.2.8. Gas Chromatography analysis for fatty acid composition
Hydrolysed and unhydrolysed thraustochytrid oil compositions were analysed
by gas chromatography (Agilent 6890) with a flame ionisation detector (FID),
equipped with a Supelcowax 10 capillary column (30 m, 0.25 mm i.d., 0.25
thickness; Supelco). Fatty acids were converted to methyl esters by acid-catalysed
trans-esterification with minor modifications (Christie & Han, 2010). 1 ml toluene was
nonadecanoate (C19
methanol (2 ml) was also added to the tube and kept for overnight incubation at 50 °C.
Fatty acid methyl esters (FAMEs) were extracted into hexane. The hexane layer was
removed and dried over sodium sulphate. FAMEs were concentrated using nitrogen
gas. Helium was used as the carrier gas at a flow rate of 1.5 ml min-1. The injector was
identified on comparison of retention time data with internal standard (Sigma-Aldrich,
St. Louis, MO, USA) and corrected using theoretical relative FID response factors
(Ackman, 2002). The peaks were quantified with chemstation chromatography
software (Agilent Technologies, Santa Clara, CA, USA).
6.3. Results and discussion
6.3.1. Thraustochytrid cultivation
102
Chapter 6
Thraustochytrids are marine microalge known for their ability to produce large
quantities of lipid, including a high ratio of omega-3 fatty acids, particularly DHA.
The carbon source used in the culture medium has a significant effect on biomass and
lipid productivity (Abedini Najafabadi et al., 2015). In this study, we used production
medium with glycerol as a carbon source for thraustochytrids cultivation, since
glycerol is a low cost carbon source that thraustochytrids grow well on (Gupta et al.,
2013; Li et al., 2015). In this study the highest biomass and lipid production achieved
were 12.23 ± 0.4 gL-1 and 29% ±1.9 for Schizochytrium sp. S31 and 14.65 ± 0.54 g-L-
1 and 35% ± 2.1 for Schizochytrium sp. DT3.
6.3.2. Cell disruption by bead mill
Bead milling has been utilised for cell disruption and extraction of intracellular
metabolites from microorganisms such as yeasts (Middelberg, 1995), bacteria (Schütte
et al., 1983), filamentous fungi (Baldwin & Moo-Young, 1991) and even microalgae
such as Chlorella vulgaris (Postma et al., 2015). Nevertheless, the use of bead milling
in lipid extraction from marine microalgae thraustochytrids has not been previously
reported. Important process parameters such as biomass concentration, bead size, RPM
and process time were optimised in the present study to improve the lipid extraction
from thraustochytrids specifically. The experimental process from thraustochytrid cell
disruption to enzyme concentration used in the present study is depicted in Fig. 6.1.
103
Chapter 6
Figure. 6.1. Schematic diagram of experimental process followed from thraustochytrid
cell disruption to enzymatic omega-3 fatty acids concentration.
The extent of cell disuprtion depends on residence time, shear forces, type of
microroganism, and cell conentration etc. During experimental stage, biomass
concentration, processing time, bead size (0.4-0.6 and 0.8-1.0 mm zirconium oxide)
and processing speed were optimised and the bead load (54%) was kept constant based
on manufactures advice. Various biomass concentrations (1%, 3%, 5% and 7%) and
processing times (in minutes 2, 4, 6 and 8) were investigated. Thraustochytrids cells
disrupted by bead mill were observed under a microscope to validate the cell
disruption. Fig. 6.2 shows typical microscopic images of disrupted cells at optimised
bead mill conditions.
Figure. 6.2. Thraustochytrid cell disruption (40 X magnification) employing bead
mill at 5% biomass concentration, bead size 0.4-0.6 mm, 4500 rpm and 4 min
processing time.
The rate of cell disurption will increase with increasing stirring speed,
however, simultaneous increase in biomass load may decrease this effect. The degree
of cell disruption increased with increasing bead filling volume in the grinding
chamber, up to about 70% where interference among the grinding elements prevents
the establishment of effective cell disruption (Doucha and Livansky, 2008). As shown
in Fig. 6.3, lipid recovery increased up to biomass concentrations of 5% and then
104
Chapter 6 decreased, probably due to increased viscosity above 5% (Oh et al., 2009). Our results
are in aggrement with a previous study in which the high cell density of
Chlorochoocum was found not to increase cell disruption (Doucha & Lívanský, 2008),
but are inconsistent with a study showing that yeast mass concentration during bead
milling had no significant effect on cell disruption (Baldwin & Moo-Young, 1991;
Mogren et al., 1974).
Figure. 6.3. Optimisation of lipid extraction by bead mill as function of biomass
concentration. Four different biomass concentrations include 1 g, 3 g, 5 g and 7 g
were used.
Lipid yield increased between 1500 rpm and 4500 rpm while further increase
in rpm had no effect on lipid yield (Fig. 6.4). Doucha & Lívanský, 2008) found that
the agitator speed (rpm) between 4500 rpm and 5500 rpm resulted in maximal cell
disintegration in Chlorococum cells. Increasing milling time increased lipid recovery
at all biomass concentrations. Maximum lipid recovery was achieved with 5%
biomass, 4500 rpm and 4 min processing time, with no increase in lipid recovery after
4 minutes since thraustochytrid cells were completely disrupted at that time (Fig. 6.2).
A recent study using bead milling to extract Chlorella cells resulted in 90-95% cell
disruption in 4 min of bead milling (Postma et al., 2015).
20
25
30
35
40
45
0 2 4 6 8
Lipi
d yi
eld
(%)
Time (min)
1 g3 g5 g7 g
105
Chapter 6
Figure. 6.4. Effect of bead mill agitator speed (rpm) on lipid yield from thraustochytrid
biomass (5% concnetration) at 4 min processing time
In order to understand the effect of bead size on thraustochytrid cell disruption
and lipid yield, we investigated two diffent bead sizes, 0.4-0.6 mm and 0.8-1.0 mm.
The maximum lipid yield (40.5% ± 0.9) was achieved with 0.4-0.6 mm beads
compared to 38.6 ± 1.2 with 0.8-1.0 mm beads. The optimum bead sizes was
previously indicated to be 0.5-0.7 mm for yeast, 0.25-0.5 mm for bacteria and 0.3-0.7
mm for C. vulgaris (Schütte et al., 1983) (Doucha & Lívanský, 2008; Lee et al.,
2012a).
At low biomass concentration 10 Kg m-3 and 8 min processing time, the highest
energy consumption was 528 MJ Kg-1 biomass and lipid yield was 31.45%. Maximum
lipid recovery was observed with 50 Kg m-3 and 4 min processing time, yielding
40.12%, with an estimated energy consumption of 264 MJ Kg-1. The disruption
efficiency was quantified as a measure of lipid recovery and had the highest efficiency
at 50 Kg m-3 and 4 min processing time with low energy consumption compared to
other parameters (Table 6.1). Lee et al, 2010, reported the energy consumption of bead
milling on three different microalgae Botryococcus, Chlorella and Scenedesmus is
equivalent to 504 MJ kg-1 of the dry mass.
Table 6.1. Energy consumption analysis for different mass concentrations and lipid
yields. Schizochytrium sp. 31 biomass was used in different mass concentration and
energy was calculated where the maximum lipid yield was achieved.
05
1015202530354045
1500 2500 3500 4500 5500
Lipi
d yi
eld
(%)
RPM
106
Chapter 6 Mass
concentration
(Kg m-3)
Energy
consumption (MJ
Kg-1 mass)
Lipid yields (%) Process time
(min)
10 528 31.45 8
30 396 35.87 6
50 264 40.12 4
70 396 38.4 6
6.3.3. Lipid profile analysis
The optimised bead mill processing parameters were applied to lipid extraction
from two thraustochytrid species, resulting in 49.6% lipid as a percentage of dry
weight, with 39.8% DHA in the extracted lipid, for Schizochytrium sp. DT3. Extraction
of Schizochytrium sp. S31 resulted in lipid (39.06%) as a percentage of dry weight,
with 31.4% DHA in the extracted lipid (Fig. 6.5). Our results are in agreement with a
recent study on lipid and DHA production from Aurantiochytrium limacinum SR21,
which achieved maximum lipid and DHA accumulation using glycerol as the carbon
source (Li et al., 2015). A similar study conducted by Gupta et al, (2015) investigated
the suitability of Schizochytrium sp. DT3 for lipid production from enzyme sachrified
hemp, and reported DHA accumulation of 38% (Gupta et al., 2015). Schizochytrium
sp. DT3 oil contained more DHA than did strain S31 and so was investigated further
for the lipase concentration of DHA.
107
Chapter 6
Figure. 6.5. Fatty acid composition of lipid extracted from thraustochytrids S31 and
DT3.
6.3.4. Selective hydrolysis of Schizochytrium DT3 oil using Candida rugosa lipase
Before hydrolysis, oil extracted from the DT3 strain was confirmed to be
essentially in the triacylglycerol form (Fig. 6.6a). Candida rugosa lipase (CRL) was
used for DHA concentration since it is known to preferentially hydrolyse saturated
(SFAs), then mono-unsaturated fatty acids (MUFAs), with low activity towards
polyunsaturated fatty acids (PUFAs) from TAGs (Kahveci et al., 2010; McNeill et al.,
1996). Enzyme concentrations from 1000 U-4000 U were tested with 500 mg of
thraustochytrid oil where oil was incubated with different enzyme concentrations and
samples were collected at specific time interval and analysed for hydrolysis. Fig. 6.7
shows that the percentage of oil hydrolysed increased with in increase in enzyme
concnetration, particularly with 3000 and 4000 U enzyme resulting in more than 50%
of hydrolysis after 3 hrs. Enzyme concnetrations of more than 3000 U did not increase
the level of thraustochytrid oil hydrolysis, possibly because of substrate limitation. A
similar trend was observed in the hydrolysis of crude palm oil by Candida rugusa
lipase (You & Baharin, 2006). Enzymatic hydrolysis with Candida rugosa lipase
05
101520253035404550
% T
otal
fatty
aci
ds
Major fatty acids
S31 DT3 isolate
108
Chapter 6 resulted in free fatty acid increase over time (Fig. 6.8), with corresponding increases
in diacyl and monoacyl glycerolds (Fig. 6.6b). Fifty percent hydrolysis of TG occurred
in 3 hours. Candida rugosa lipase preferentially hydrolysed myristic acid (C14:0) and
plamitic acid (C16:0), with DHA and DPA being only slightly hydrolysed (Fig. 6.9).
a
b
Figure. 6.6. Capillary chromatography (iatrascan) profile of thraustochytrid oil a) before and
b) after hydrolysis by Candida rugosa lipase at 40 C.
-2
0
2
4
6
8
10
12
14
16
0 0.1 0.2 0.3 0.4 0.5 0.6
Sign
al (m
V)
Time (min)
TAG
-2
0
2
4
6
8
10
0 0.1 0.2 0.3 0.4 0.5 0.6
FFA
DA
MAG
TAG
Time (min)
Sign
al (m
v)
109
Chapter 6
Figure. 6.7. Optimisation of enzyme concetrtation. 500 mg of thraustochytrid oil was
incubated with varying enzyme concentrations and samples were withdrawn at
different time intervals.
Figure. 6.8. Lipid class analysis of thraustochytrid oil after hydrolysis with Candida
rugosa lipase at different time intervals.
0
10
20
30
40
50
60
70
0 1 2 3 4 5
Hyd
roly
sis
(%)
Time (h)
1000 (U)2000 (U)3000 (U)4000 (U)
0
20
40
60
80
100
120
0 1 2 3 4
Lipi
d cl
ass (
%)
Time (h)
TAG FFA
110
Chapter 6
Figure. 6.9. AG and FFA are separated hydrolysed oil and analysed by gas
chromatography. Short chain fatty acids were preferentially hydrolysed form TAG
and DPA and DHA significantly concentatred at glyceride portion.
In the triacylglycerol portion myristic acid was reduced from 10.2% to 4.3% and
palmitic acid from 35.3% to 10.24%. DPA in the triacylglyercol portion increased
from 19.2% to 29.14%, with DHA increasing from 39.8% to 55.45%. CRL has been
shown previously to preferably hydrolyse short and medium chain fatty acids while
increasing omega-3 levels in the remaining triacylglyercides (Kahveci & Xu, 2011
(Nalder et al., 2014). This fatty acids selectivyt is due to the to the deep tunnel like
active site which is buried inside the protein structure. Under optimised conditions in
this study the omega-3 content of thraustochytrid oil was enriched to 88.7%, 1.4 folds
above initial levels. The fatty acid profiles obtained were as expected, from fatty acid
chain length selectivity of Candida rugosa lipase.
6.4. Conclusion
Schizochytrium sp. DT3 is a renewable source of omega-3 fatty acids,
particularly DHA. We optimised a bead mill based cell disruption method for
maximising lipid extraction with lower energy consumption. We proposed a simple
enzymatic process for the concentration of DHA from marine microalgae
0
10
20
30
40
50
60
70
C14:0Myristic
acid
C16:0palmitic
acid
C18:0stearicacid
C18:1Oleic acid
C20:5n3EPA
C22:5n6DPA
C22:6n3DHA
Fatt
y ac
id c
ompo
sitio
n (%
)
Major fatty acids
Native oil AG FFA
111
Chapter 6 Schizochytrium sp. DT3 oil using Candida rugosa lipase. This lipase at 3000 U
hydrolysed 50% of thraustochytird oil in 3 h, increasing DHA levels by 1.4 fold.
112
Chapter 7
Chapter 7: Phospholipase A1 as a catalyst to enrich omega-
3 fatty acids in anchovy and algal oil
113
Chapter 7 7.1. Introduction
Consumption of fish oils has been shown to have health benefits, mainly in
cardiovascular events, based on several prospective and retrospective studies (Lavie et
al., 2009; Molfino et al., 2014). Most of the benefits of fish oil were attributed to the
polyunsaturated omega-3 ( -3) fatty acids eicosapentaenoic acid (EPA) and
docosahexaenoic acid (DHA) (Udani & Ritz, 2013). The American Heart Association
recommends consumption of 1g/day of omega- fatty acids to patients with coronary
heart diseases (Kralovec et al., 2012). Omega-3 fatty acids constitute approximately
30% of the total fatty acids in natural fish oils. Concentrated esters of omega-3 fatty
acids have been formulated to deliver higher amounts of EPA and DHA per dose,
particularly in pharmaceutical products where 80% EPA and DHA concentrates are
used (Molfino et al., 2014). However, conversion to ethyl esters followed by fractional
distillation and urea concentration is damaging to these oxidatively sensitive omega-3
fatty acids. Also, re-esterification to triacylglycerides requires further damaging
processing of these oils, and also results in statistical distribution of fatty acids on the
glycerol backbone (Catal et al., 2013).
To overcome heat induced degradation, lipases were employed to selectively
hydrolyse omega-3 fatty acids. Triglycerides from natural fish oils are complex in
composition (Miller et al., 2008), besides containing several kinds of fatty acids,
mainly saturated, monounsaturated and polyunsaturated, the fatty acids are also non-
uniformly distributed on the glycerol backbone. 13C NMR spectral studies on anchovy
fish oils show that DHA in more abundant at sn-2 than at sn-1 and -3 positions, while
EPA is nearly statistically distributed across all three positions (Akanbi et al., 2013).
Five lipases were tested for specificity in the hydrolysis of fish oil, using fatty acid
esters as a model system. Discrimination for EPA and DHA was observed with fatty
acid esters but not with fish oils (Lyberg & Adlercreutz, 2008). Another study on lipase
mediated fish oil hydrolysis suggested that hydrolysis is biased towards the chemical
nature of the fatty acid rather than to their positional abundance (Akanbi et al., 2013).
Statement of Contribution to Research for this Chapter: The research outlined in the chapter was carried
out jointly with Mr. Tushar Ranjan Moharana (PhD student) and Dr Madhusudhana Rao at CCMB,
Hyderabad, India, with funding from an Australia-India strategic research grant. The experiments data
shown in Figures 7.2B, 7.3B, 7.5B, 7.6 and 7.7 were performed by Mr. Moharana and the data shown
in Figures 7.1,7.2A, 7.3A, 7.4, 7.5A, 7.8, 7.9 and Table 1 were performed by me.
114
Chapter 7
Lipases have been extensively used for selective concentration of -3 fatty acids from
fish oils either by fish oil hydrolysis or selective esterification (Akanbi et al., 2013;
Kosugi & Azuma, 1994; Kralovec et al., 2010; Shimada et al., 1997). Lipases possess
some important properties such as selectivity towards chain length and position of fatty
acid in a triglyceride and also discriminate between fatty acids with single and multiple
double bonds. These properties of lipases make them as suitable candidates for
enzymatic concentration of omega-3 fatty acids (Shahidi & Wanasundara, 1998). Most
lipases preferentially hydrolyses saturated and mono-unsaturated fatty acids from
triacylglycerides and strongly discriminate against omega-3 fatty acids, due to the
presence of double bonds that enhance the steric hindrance at lipase active site (Casas-
Godoy et al., 2014; Halldorsson et al., 2004). Most lipases preferentially hydrolyse
EPA over DHA, probably due to the presence of an additional double bond located
closer to the ester bond in DHA (Akanbi et al., 2013; Shimada et al., 1998). In another
study, pancreatic lipase was observed to preferably hydrolyse docosapentaenoic acid
(DPA) over EPA and DHA (Akanbi et al., 2014).
Phospholipase A1 specifically hydrolyses the fatty esters at the sn-1 position
and release 2-acyl lysophospholipid and a fatty acid. Functions of PLA1 are not clearly
established and some PLA1 were reported to show lipase-like activity (Mishra et al.,
2009; Richmond & Smith, 2011). Lipases that show phospholipase activity have been
extensively investigated, but not the reverse (Aloulou et al., 2014). The catalytic
mechanism between lipases and phospholipases is identical; however, the specificity
emerges from the active site properties. Studies have indicated that lipases with a short
lid and a short 9 loop are more suitable to accommodate polar phospholipids (Carrière
et al., 1998). PLA1 was tested for its specificity on triglycerides in this study and the
activity profiles were compared with those of a lipase from Bacillus subtilis. To
investigate the ability of a phospholipase in regioselective hydrolysis of
triacylglycerols, we choose PLA1 since it is likely to be selective for the sn-1, 3
position in triacylglycerols.
Phospholipase A1 is a recombinant enzyme manufactured by Novozymes by
the fusion of a lipase gene from Thermomyce lanuginosus and a phospholipase gene
from Fusarium oxysporum. This enzyme was observed to hydrolyse both phospholipid
and triglycerides with a single active site (Fernandez-Lorente et al., 2008; Mishra et
al., 2009). The phospholipase activity of this enzyme is applied in the commercial
115
Chapter 7 degumming of vegetable oils and for modifying phospholipids (Sheelu et al., 2008;
Yang et al., 2008). The lipase activity of PLA1 has been applied in organic synthesis
and in the synthesis of structured lipids (Sheelu et al., 2008). PLA1 also successfully
immobilised on different nanoparticles to further improve its catalytic properties (Liu
et al., 2012; Mishra et al., 2009; Wang et al., 2010). Therefore, in this study, we
attempted to use phospholipase A1 for the concentration of omega-3 fatty acids from
anchovy fish oil. We investigated the chain length selectivity of phospholipase A1
using p-NP esters. The ability of Phospholipase A1 to concentrate omega-3 fatty acids
into the glyceride portion was investigated by examining the preferential fatty acid
hydrolysis using gas chromatography and 13C NMR spectroscopy.
7.2. Material and Methods
7.2.1. Chemicals
Bleached Anchovy oil was supplied by Ocean Nutrition Canada (Canada). The
major fatty acid composition of native oil was determined by Gas chromatography
analysis (Akanbi et al., 2013). Phospholipase A1 (Lecitase Ultra®) and gas
chromatography standards were procured from Sigma-Aldrich (Castle Hill, Australia)
and hexane (Merck). All the buffers were made using analytical grade chemicals. All
solvents used were either analytical grade or higher. Gum Arabic and phospholipase
A1 (PLA1) and methyl nonadecanoate were purchased from Sigma Aldrich. TLC
plates were purchased from Merck.
7.2.2. Hydrolysis of anchovy oil
Reaction mixture containing 50 mM Tris, 25 mM CaCl2, 5% (w/v) gum Arabic
at pH 8.00 and 5% (v/v) anchovy oil were micellised by sonication. 100 unit of PLA1
was added and the reaction was carried at 37 C at constant stirring under nitrogen.
itrating
against 1 M NaOH. 2 ml of sample were withdrawn at every time point and free fatty
acid was separated from glyceride by solvent extraction and the purity of both the
fractions were analysed by TLC. Solvent was evaporated under nitrogen gas and
fractions stored in an airtight polypropylene tube at -20 C for further analysis. Oxygen
was removed by blowing nitrogen whenever the sample was exposed to air.
116
Chapter 7 7.2.3. Thraustochytrid oil hydrolysis by phospholipase A1
Thraustochytrid lipid extraction and analysis was described in section 6.2.5,
6.2.7 and 6.2.8. Phospholipase hydrolysis was performed based on the method
described in section 6.2.6 with some modifications. 500 mg of thraustochytrid oil and
1.8 ml of Tris HCL buffer (50 mM, pH 7.0) were taken in 25 ml round bottom flask
and heated up to 45 oC and 300 rpm stirring. Substrates were stirred for 30 min and
Phospholipase A1 (2500 U/g oil) added to initiate the hydrolysis. Samples were
collected periodically and analysed.
7.2.4. IATROSCAN analysis
Both the unhydrolysed and hydrolysed portions of the fish oil were analysed
by capillary chromatography with flame ionization detector (Iatroscan MK5, Iatron
Laboratories Inc., Tokyo, Japan). The Iatroscan settings were: air flow rate; 200
ml/min, hydrogen flow rate; 160 ml/min and scan speed of 30s/scan. Under these
conditions, the chromarods were cleaned by scanning twice before applying samples.
One microliter of each lipid fraction in hexane was spotted onto the rods with the aid
of an auto pipette along the line of origin on the rod holder and developed for 22
minutes in a solvent tank containing hexane/diethyl ether/acetic acid (60:17:0.2,
vol/vol/vol). TLC standards purchased from Nu-Chek Prep were used to identify each
lipid class (Akanbi et al., 2013).
7.2.5. Methylation and GC analysis
5 μL (both free fatty acid and glyceride) of sample was methylated by using
acetyl chloride in methanol as described by Christie et al 2010 (Christie & Han, 2010).
Hydrolysed and unhydrolysed fish oil was analysed using gas chromatography
(Agilent 6890) with flame ionisation detector (FID), equipped with a Supelcowax 10
. Fatty acids
were converted to methyl esters by acid-catalysed esterification with minor
modifications. 1 ml toluene was added to the methylation tubes followed by the
(5mM) of internal standard, methyl nonadecanoate (C19:0) and 200
(1mM) of butylated hydroxytoluene (BHT). 2 ml acidic methanol (prepared by
adding 10% acetyl chloride in methanol drop wise on ice bath) was also added to the
tube and kept for overnight incubation at 50°C. Fatty acid methyl esters (FAMEs) were
117
Chapter 7 extracted into hexane. The hexane layer was removed and dried over sodium sulphate.
FAMEs were concentrated using nitrogen gas and analysed by gas chromatography.
Helium was used as the carrier gas at a flow rate of 1.5 ml min-1. The injector was
identified on comparison of retention time data with external standards (Sigma-
Aldrich, St. Louis, MO, USA) and corrected using theoretical relative FID response
factors (Ackman, 2002). Peaks were quantified with Chemstation chromatography
software (Agilent Technologies, Santa Clara, CA, USA).
7.2.6. Analysis of positional distribution of fatty acid by NMR
300 μl of glyceride were dissolved in 1 ml of CDCl3 (99.8% pure). NMR
spectra were collected by using Bruker 600MHz NMR machine. Peak corresponding
to different fatty acid at different position were assigned as described by (Suárez et al.,
2010). Relative quantification was done by comparing the area under each peak.
Quantitative 13C NMR spectra of the unhydrolysed oils were recorded under
continuous 1H decoupling at 24°C.In order to quantify the residue of each fatty acid at
different positions, peak area ratios were analysed by integration and presented in
percentages (Akanbi et al., 2013).
7.3. Results and discussion
7.3.1. Lipase activity of Phospholipase A1
PLA1 used in this study was isolated from Thermomyces lanuginosus
(LecitaseTM) and is a recombinant enzyme preparation. This enzyme has been explored
primarily for the degumming of vegetable oils (Yang et al., 2006). Some studies
include the use of PLA1 lipase activity for the resolution of mandelate esters and the
regioselective hydrolysis of carbohydrate esters (Mishra et al., 2009). Lipase-like
activity of PLA1 was reported earlier. However its ability to hydrolyse triglycerides is
only marginal in the vegetable oil degumming processes. PLA1 was previously
characterised for substrate selectivity and it was observed that this enzyme does
hydrolyse C2-esters (Mishra et al., 2009). Initially the lipase activity of PLA1 was
tested using p-nitrophenyl esters of various chain length fatty acids. Some of the p-NP
esters were synthesised based on published methods (Nalder et al., 2014; Neises &
Steglich, 1978). Fig. 7.1 shows that PLA1 was able to hydrolyse esters of fatty acid
118
Chapter 7 chain lengths from C4–C20 with comparable efficiency. The activity was highest for
C10 and lowest with C4 ester (40% of C10). The presence of unsaturation, single or
multiple double bonds, did not impact the activity appreciably. However, esters of
DHA were not efficiently hydrolysed. A similar chain length study was performed for
lipase from Bacillus subtilis (BSL) and it was found that C8 chains were most
efficiently hydrolysed (Dartois et al., 1992).
Figure.7.1 Phospholipase A1 activity towards p-NP esters with different acyl
chains. Activity of PLA1 on p-NP-decanoate was taken as 100%.
Lipase activity of phospholipases and vice versa was studied in few cases. The
ratio of lipase to phospholipase activity of lipases or phospholipases varies widely and
the ratio is not only related to the structure of the lipase but also to the reaction system
(Yang et al., 2008). For example, PLA1 from Thermomyces lanuginosus used in this
study shows hydrolytic activity against both triacylglcyerides and phospholipids and
is used to degum oils (Sheelu et al., 2008). PLA1 shows predominately phospholipase
activity at reaction temperatures above 40 oC. Pancreatic phospholipases hydrolyse
phospholipids in the aqueous phase, while lipase from Fusarium oxysporum
hydrolyses phospholipids in the oil phase (Yang et al., 2008).
7.3.2. Hydrolysis of Anchovy oil by PLA1
Initially hydrolysis of anchovy oil was tested in an oil-in-water emulsion and
the extent of hydrolysis was monitored using Iatroscan. Fig.7.2A shows the scan
119
Chapter 7 before and after hydrolysis of the oil by PLA1. To obtain more quantitative kinetics of
the hydrolysis, anchovy oil hydrolysis was monitored by a pH Stat, while maintaining
a constant pH, using 5% Gum Arabic as emulsifier. Initially we measured the rate of
hydrolysis of anchovy oil by PLA1 and BSL (Fig.7.2B). Both enzymes were able to
hydrolyse the oil to a comparable extent. At 37 oC the reaction reached 45% hydrolysis
and then plateaued.
Figure. 7.2. Time course of hydrolysis of anchovy fish oil by PLA1 and BSL
A, Iatroscan of unhydrolysed (light trace) and hydrolysed (darker trace) fraction of
anchovy oil by PLA1. B, pH stat based determination of rate of hydrolysis of anchovy
oil in the presence of PLA1 (closed circle) and BSL (open circle).
The fatty acid fraction was separated from the glyceride fraction, methylated
and analysed by GC. Fig.7.3A shows the time course of release of various fatty acids
120
Chapter 7 from anchovy oil by PLA1. Both saturated and monounsaturated fatty acids were
extensively hydrolysed (approximately 40% hydrolysis), while the hydrolysis of both
the omega-3 fatty acids EPA and DHA was relatively poor 15% and 4%, respectively.
Identical experiments were also performed with BSL (Fig.3B). BSL also hydrolysed
oil with similar efficiency. However, the extents of hydrolysis of various fatty acids
were similar. While the extents of hydrolysis of unsaturated, monounsaturated and
EPA were equal (50%), hydrolysis of DHA was marginally less at 30%. The fatty acid
products obtained after 3 h hydrolysis are shown in Fig.7.4. The data suggests that
PLA1 discriminates against EPA and DHA, while BSL shows marginal discrimination
against DHA. A significant five-fold higher discrimination of DHA by PLA1
compared to BSL was observed.
121
Chapter 7 Figure. 7.3. Time course of hydrolysis of anchovy oil and fatty acid distribution in the
hydrolysate. Anchovy oil was subjected to hydrolysis by PLA1 (A) and BSL (B). The
reaction product at various times was subjected to methylation and GC analysis.
Percent hydrolysis of each fatty acid was calculated based on its hydrolysis at tx as a
fraction of to.
PLA1 specifically hydrolyses sn-1 acyl esters from phospholipids producing
FFAs and lysophospholipase. PLA1 enzymes normally exhibit very little
lysophospholipase or lipase activity. However, some PLA1 enzymes exhibit some
neutral lipase activity. PLA1 enzymes may be descendants of neutral lipases, and
several PLA1 sequences show substantial sequence similarity to the well characterised
pancreatic, hepatic and endothelial lipases (Aoki et al., 2007; Richmond & Smith,
2011). Although PLA1 enzymes are found in a wide variety of cells and tissues, only
a small number of PLA1 enzymes have been cloned and their substrate preferences
have not been well studied (Richmond & Smith, 2011). Several structural and protein
engineering studies on lipases have focused on the architecture of the active site and
its substrate preferences. The hydrophobicity and 9
loop and the lid domain of lipases play a selective role in preferring a triglyceride or a
phospholipid (Aoki et al., 2007; Tiesinga et al., 2007). The lid domain of guinea pig
pancreatic lipase related protein 2(GPLRP2) is reduced in size and consequent
exposure of hydrophilic residues enables the active site to accept phospholipids and
larger galacto lipids (Carrière et al., 1998). Human pancreatic lipase that does not show
phospholipase activity was not able to accommodate the phospholipid molecule.
Sequence alignment of porcine pancreatic lipases indicates that Val260 is critical for
interaction with lipids (Van Kampen et al., 1998).
PLA1 from Thermomyces lanuginosus, used in this study, discriminated
against DHA and partly against EPA when hydrolysing anchovy oil. In similar
experiments using a lipase from Bacillus subtilis there was no observed selectivity
against EPA and DHA. Anchovy oil has DHA preferentially located at the sn-2
position (60%), compared to the sn-1, 3 positions (40%). If PLA1 preferentially
hydrolyse at the sn-1, 3 position in TG oils then more DHA than EPA should be present
in the unhydrolysed fraction. This was observed in our experiments (Fig.7.4). In our
study PLA1 poorly hydrolysed DHA at all positions, and preferably hydrolysed EPA
122
Chapter 7 at the sn-1, 3 positions. BSL preferentially hydrolysed DHA at the sn-2 position, while
preferentially hydrolysing EPA at sn-1, 3.
7.3.3. 13C NMR studies of the anchovy oil hydrolysis by PLA1 and BSL
13CNMR spectra of complex triglycerides, such as fish oils, can provide
positional information on the various fatty acids in the oil. This information enables
the study of the positional specificity of hydrolysis by PLA1 and BSL and also to
obtain quantitative information on the extent of hydrolysis of fatty acids at each
position. Initially we acquired the positional information of various fatty acids in
anchovy oil. Table 7.1 provides the details of positional distribution of fatty acids in
Anchovy oil. The ratio of abundance of various fatty acids at sn-1, 3 and sn-2 positions
was observed to be saturated 1.64, monounsaturated 3.06, EPA 4.2 and DHA 0.6. A
ratio of 2 suggests the fatty acids are equally distributed. A ratio of 0.6 for DHA
indicates it is predominantly present at the sn-2 position. The ratio is high (4.2) for
EPA indicating its preferential distribution at sn-1, 3 position, compared with the
distribution of DHA.
Table 7.1. Positional distribution of various fatty acids in anchovy oil.
Fatty acid
Distribution (%)
sn-1,3 sn-2
Saturated 22.9 14.0
Mono unsaturated 18.7 6.1
Stearidonic acid (STA) 6.2 2.7
EPA 14.9 3.5
DHA 3.9 6.8
Anchovy oil hydrolysis by PLA1 and BSL was allowed to proceed until 30%
of the oil was hydrolysed and then the unhydrolysed portion of the oil was extracted
and 13C NMR spectra were obtained. This was repeated with unhydrolysed anchovy
oil. Figure 7.5 shows the 13C NMR spectra of hydrolysed and unhydrolysed anchovy
oil subjected to PLA1 (Fig. 7.5A) and BSL (Fig. 7.5B) treatment. The overlay of
hydrolysed and unhydrolysed spectra for both PLA1 and BSL clearly demonstrates
that BSL preferentially hydrolyses EPA and DHA more than does PLA1. Fatty acids
123
Chapter 7 remaining in the unhydrolysed portion of the oils with PLA1 and BSL are shown in
Fig. 7.4. Saturated and monounsaturated fatty acids were preferentially hydrolysed by
both the enzymes while omega-3 fatty acids were discriminated against. BSL also
preferentially hydrolysed EPA over DHA.
Figure. 7.4. Percent hydrolysis of various fatty acids (from Fig.7.3) at 32% hydrolysis
of anchovy oil by PLA1 (light) and BSL (dark). The relative proportions at other time
points were also similar.
124
Chapter 7
Figure. 7.5. 13C NMR spectra of the unhydrolysed oil fraction before (blue) and after
(red) 30% hydrolysis of anchovy oil by PLA1 (A) and BSL (B). Fatty acids and their
positions were marked. Sat (saturated), mono (monounsaturated), STA (stearidonic
acid), EPA (eicosapentaenoic acid) and DHA (docosahexaenoic acid).
The percent accumulation data shown in Fig. 7.6A indicates that EPA at sn-1
was more efficiently hydrolysed by BSL than by PLA1. BSL hydrolysed EPA at sn-1
but not DHA at sn-1or at sn-2, while PLA1 had no preference for EPA over DHA.
These results indicate that the discrimination against omega-3 fatty acids by PLA1 is
a result of the chemical nature of the fatty acid and not its position on the TG. To verify
this observation, we have performed hydrolysis of a structured triglyceride, 1, 3-
dioleoyl-2-hexadecanoic glyceride, using PLA1 and BSL and estimated the fatty acid
products by GC (Fig.7.7). If PLA1 shows absolute preference for fatty acids at the sn-
Sn-1,3 Sat
Sn-1,3 Mono unsat
Sn-1,3STA
M
Sn-1,3EPA
Sn-2 Sat
Sn-2 Mono Unsat
Sn-1,3 STA
Sn-2 DHA
Sn-1,3DHASn-
Sn-2 EPA
A
B
125
Chapter 7 1, 3 position, the hydrolysis of hexadecanoic acid at sn-2 should be much lower than
that of oleic acid. The difference between the specificity of hydrolysis by PLA1 and
BSL was also negligible, indicating that PLA1 did not show any specificity towards
the sn-1,3positions. However, there is a distinct preference against DHA in anchovy
oil, indicating the fatty acid specificity of this enzyme.
Figure. 7.6. Percent accumulation of different fatty acid classes in TG fraction after
30% hydrolysis of anchovy oil by PLA1 (dark) and BSL (light). A, Percent
accumulation of each fatty acid is calculated as (UHFAx-TotalFAx)/TotalFAx) X 100).
Each of the fatty acid fractions were calculated based on peak area in the NMR spectra.
B, Percent accumulation data replotted with positional information.
126
Chapter 7
Hexadecanoic acid 9-Octadecenoic acid0
10
20
Fatty
aci
d re
leas
ed (
mol
es)
Figure. 7.7. Lipase hydrolysis of structured triglyceride1, 3-dioeoyl-2-myristic
triglyceride was subjected to hydrolysis by PLA1 (Light) and BSL (Dark) and the
released fatty acids were quantified by GC analysis.
To assess the positional specificity of these hydrolases we have employed a
structured glyceride with fixed positional distribution of fatty acids. Using 3-dioleoyl-
2-myristic glyceride, a structured TG, neither PLA1 nor BSL exhibited any position
specific hydrolysis under our reaction conditions, with both fatty acids being equally
hydrolysed. However, with anchovy oil the hydrolysis by PLA1 clearly shows bias
against DHA, indicating fatty acid selectivity against polyunsaturated fatty acids. In a
previous study five lipases were tested for their specificity by using fish oil and methyl
esters of EPA, DHA and palmitic acid. All lipases discriminated against EPA and
DHA when presented as methyl esters. However, lipase from Thermomyces has shown
discrimination against DHA, particularly in the early stage of the hydrolysis reaction.
A study observed that Candida rugosa lipase was most efficient in the enrichment of
DHA in the glyceride fraction (Lyberg & Adlercreutz, 2008). In another study lipase
from Candida rugosa discriminated omega-3 fatty acids from other fatty acids during
hydrolysis of sardine oil (Okada & Morrissey, 2007). These studies indicate that the
specificity in hydrolysis of natural oils by lipases or phospholipases is strongly
dependent on the temperature and duration. Also, positional selectivity can found fatty
127
Chapter 7 acid selectivity analyses in natural triacylglycerides, since positional distribution is not
always analysed in selectivity studies.
Our study and similar studies with PLA1 lipase from Thermomyces
lanuginosus suggests that hydrolysis is due to fatty acid selectivity more than to
regioselectivity (Akanbi et al., 2013). A related previous study showed that substrate
binding was dependent on the relative volume of the substrate to the volume of the
active site (Kamal et al., 2015). Thirty eight lipases were investigated in silico, for
their affinity to structured TGs with different chemical and positional compositions.
This study indicated that binding affinity differences between various substrates with
lipases is complex, but helped to identify active site positions that are critical to
binding. This information could be useful for designing site saturation mutagenesis to
identify amino acid substitutions that would enhance hydrolysis of specific fatty acids.
Few studies have successfully identified amino acid positions that are important in
binding TG and phospholipids (Carrière et al., 1998). Further studies are required to
enable the design of enzymes that can specifically hydrolyse omega-3 fatty acids.
7.3.4. Thraustochytrid oil hydrolysis by phospholipase A1
Phospholipase A1 is an engineered carboxyl ester hydrolase, produced from
Thermomyces lanuginosus/Fusarium oxysporum by genetically modified Aspergillus
oryzae. Phospholipase A1 possess activity towards both phospholipids and
triglycerides. (Mishra et al., 2009). Research has focused on its phospholipase activity
and little information is available on the production of structured glycerides using
phospholipase A1 from oils (Liu et al., 2012; Liu et al., 2011). DT3 oil was hydrolysed
to determine phospholipase A1 activity towards concentration of omega-3 fatty acids.
Hydrolysis was gradually increased with incubation time and reached 50% at 150 min.
Fig. 7.8 illustrates the percentage of hydrolysis of DT3 oil by phospholipase A1.
128
Chapter 7
Figure. 7.8. Lipid class analysis of thraustochytrid oil after hydrolysis with
phospholipase at different time intervals.
Fatty acids profiles of glycerol and free fatty acids of hydrolysed DT3 oil were
analysed by gas chromatography and results are shown in Fig. 7.9. After 150 min of
phospholipase A1 hydrolysis, a significant amount of short chain fatty acids and
monounsaturated fatty acids were cleaved from the glycerol back bone and
concentrated in the free fatty acids portion. In the triacylglycerol portion, the major
saturated fatty acids were reduced, myristic acid was reduced from 10.2% to 5.3% and
palmitic acid from 35.2% to 16.24%. EPA was equally distributed in the free fatty acid
and glyceride portions and DHA was significantly enriched in the glyceride portion,
indicating a strong selectivity for EPA over DHA (Fig. 7.9). The DHA concentration
was enriched by 1.3 fold compared to native oil. These selectivity is similar to those
observed for hydrolysis of fish oil using lipase TL100 (Akanbi et al., 2013).
0
20
40
60
80
100
120
0 30 60 90 120 150 180 210
Hyd
roly
sis
(%)
Time (min)
TAG FFA
129
Chapter 7
Figure. 7.9. Fatty acid profiles of hydrolysed and unhydrolysed DT3 oil.
7.4. Conclusion
The position selective hydrolysis of PLA1 was tested for the hydrolysis of
anchovy oil. Fatty acid chain length selectivity of PLA1 was investigated using p-NP
esters of chain length C2 to C22, including the long chain PUFAs, EPA and DHA. For
anchovy oil hydrolysis SFAs and MUFAs were preferentially hydrolysed by PLA1
and omega-3 fatty acids were discriminated against. Of the omega-3 fatty acids, EPA
was more preferentially hydrolysed than was DHA. Lipase from Bacillus subtilis did
not show discrimination of fatty acids to the same extent as PLA1. Hydrolysis of the
structured TG, 1, 3–dioleoyl-2-hexadecanoic glyceride, supported that the
discrimination property of PLA1 was primarily due to the chemical nature of the fatty
acid rather than its position in the triacylglyceride. PLA1 is a potential catalyst for
selective enrichment of omega-3 fatty acids in triacylglycerides.
Phospholipase A1 was used to catalyse the hydrolysis of thraustochytrid DT3
oil to enrich DHA in glyceride portion, with short chain fatty acids being preferentially
cleaved from the triacylglyceride. This results show that phospholipase A1 can be used
to concentrate DHA from triacylglycerides.
0
10
20
30
40
50
60
C14:0Myristic
acid
C16:0palmitic
acid
C18:0stearicacid
C18:1Oleic acid
C20:5n3EPA
C22:5n6DPA
C22:6n3DHA
Com
posi
tion
(%)
Major fatty acids
Native oil AG FFA
130
Chapter 8
Chapter 8. Summary and future aspects
131
Chapter 8
The marine ecosystem is a large relatively unexplored pool for the isolation of
potential compounds and materials for industrial and biotechnological applications.
Marine microorganisms remain a major source for scientific discovery. The number
of microorganisms isolated and characterised for their potential metabolites is
increasing each year. Researchers have isolated bioactive compounds (lipids,
enzymes, polysaccharides, proteins and pigments) from a range of marine inhabitants.
The demand for novel enzymes is also increasing, particularly for the production of
chemically and structurally important molecules with industrial applicability.
Naturally marine environments provide unique genetic diversity and life habitats such
as nutrient rich regions, nutrient limited regions, high salinity, high pressure, low
temperature and light conditions. Microalgae lipid production is an area that is being
explored, although the cost of production is high, partly due to downstream processing
costs. Thus, cost-effective downstream processing methods play an important role in
achieving commercial scale production of microalgae derived metabolites.
The main objective of this study is the isolation and selection of suitable
thraustochytrid strains for lipid production, the development of lipid extraction and
omega-3 enzymatic concentration methods applicable to the production of
thraustochytrid derived oil. One aim was to develop a rapid lipid quantification method
to screen newly isolated thraustochytrids. Another aim was to optimise lipid and lipase
extraction methods. A third aim was to concentrate omega-3 fatty acids from
thraustochytrids oil using lipases.
The first aim of the study was to develop a rapid colorimetric method for lipid
quantification from thraustochytrid cells. The conventional lipid quantification
methods are laborious and time consuming. Initially, to evaluate the colorimetric
method, different lipid samples (plant oils, pure triolein, DHA and EPA) were
quantified. A straight line showed the range of linearity of the method with different
samples. In addition, the false positives from non-lipid components (glucose, glycerol
and proteins) did not interfere with lipid quantification. In this study, four different
thraustochytrids strains, two of them new isolates, were quantified for lipid content
and the results compared with those obtained using a conventional gravimetric method.
The results obtained from both methods were similar within experimental error.
132
Chapter 8
Chapter 3 describes the effect of different solvents and cell disruption methods
on lipid extraction from thraustochytrids. A total of nine solvents; chloroform,
dichloromethane, diethyl ether, ethanol, heptane, hexane, isopropanol, methanol and
toluene and their combinations, were investigated for lipid extraction. Maximum lipid
extraction for single solvents were hexane (12.5%), heptane (11%) and chloroform
(9.7%). However, a solvent mixture of chloroform and methanol (2:1) showed
maximum lipid extraction of 22%, followed by chloroform and hexane (2:1) with
13.4%. Furthermore, different cell disruption methods were evaluated to improve lipid
extraction from Schizochytrium S31 and Thraustochytrium AMCQS5-5. Cell
disruption resulted in increased lipid extraction from both Schizochytrium S31 and
Thraustochytrium AMCQS5-5, and maximal lipid extraction was obtained using
osmotic shock, with 48.7% lipid extracted from Schizochytrium S31 and 29.1% lipid
from Thraustochytrium AMCQS5-5. The other methods, grinding (44.6%), sonication
(31%) and shake mill (30.5%) extracted higher lipids than the control method (22%).
The fatty acid profiles of the lipids extracted from thraustochytrids were investigated
using GC. GC analysis revealed the suitability of S31 fatty acid profiles for biodiesel
production and AMCQS5-5 profiles for nutraceutical applications. In addition, in
silico analysis of fatty acid profiles confirms the potential of S31 lipid for biodiesel
production. The energy consumption was evaluated for the methods investigated in
this study to aid in determining their potential use in large scale processing. The
osmotic shock method was found to be the most economical method compared to other
methods for industrially scaled-up.
Thraustochytrids have been documented for polyunsaturated fatty acids
production, particularly DHA, but little information is available about their enzymatic
activities. The first part of this study (Chapter 4) describes screening of enzymatic
activities from thraustochytrid strains using the API ZYM kit. The isolates investigated
in this study were collected form two different collection trips. The investigated
thraustochytrid strains exhibited high esterase, lipase, acid and alkaline phosphatase
activities and the maximum lipase activity was observed in AMCQS1-9 followed by
AMCQS-4b27. In addition, the effect of different cell disruption methods on protein
release and lipase activity was evaluated from AMCQS1-9 and AMCQS-4b27 strains.
In this study, four different cell disruption methods, bead vortexing, grinding with
liquid nitrogen, sonication and homogenisation, were compared for impact on protein
and lipase recovery from thraustochytrids. Strain AMCQ-4b27 cells were more
133
Chapter 8 sensitive to protein release than those from AMCQS1-9 and sonication was found to
be the suitable method for lipase yield from AMCQ-4b27 (0.903 IU/μg) and
AMCQS1-9 (1.44 IU/μg). Lipase activity extracted from strain AMCQS1-9 was
approximately 1.5 fold more than that from AMCQ-4b27.
Lipase production is influenced by the culture conditions. Chapter 5
demonstrated the optimisation of Schizochytrium S31 culture conditions for enhancing
lipase production in a stirred tank reactor. In addition, sonication parameters were
optimised to improve lipase extraction yields. Initially, Tween 80 concentration was
optimised and a Tween 80 concentration of 1% gave the maximum biomass and lipase
productivity. The optimum temperature was 25 oC and the optimum pH was 6. The
process was scaled-up in a 2 L stirred tank reactor to monitor biomass, lipid and lipase
productions and utilisation of substrate and change in pH. To quantify lipid content,
fatty acid composition and lipase activity, biomass was collected and analysed at
different time intervals up to 9 days. Biomass and lipase production reached a
maximum after 5 days of incubation. Based on the results obtained from Chapter 4,
sonication was resulted in the highest level of lipase extraction from thraustochytrids.
In this study, we optimised sonication parameters to enhance lipase extraction.
Optimised sonication conditions resulted in a 5 fold increase in lipase activity
extraction. Novel Schizochytrium S31 lipase was employed to hydrolyse different
chain length p-NP esters. The extracted lipase preferentially hydrolysed shorter chain
fatty acids, however, DHA was preferentially hydrolysed over EPA.
Chapter 6 describes the use of bead milling in thraustochytrid lipid extraction
and enzymatic concentration of DHA from extracted lipids. In this study,
thraustochytrids were grown in glycerol, a low cost carbon source for biomass and
lipid production. The highest level of biomass and lipid produced were 12.23 ± 0.4 g
L-1 and 29% ± 1.9 for Schizochytrium sp. S31 and 14.65 ± 0.54 g L-1 and 35% ± 2.1
for Schizochytrium sp. DT3, respectively. Schizochytrium S31 biomass was used for
optimising lipid extraction using a bead mill. Important process parameters such as
biomass concentration, bead size, RPM and process time were optimised. The highest
lipid yield (40.5%) was obtained with biomass concentration (5%), bead size (0.4-0.6
mm), RPM (4500) and 4 min processing time. Fatty acid profiles of extracted lipid
from the both strains was compared and DHA accumulation was observed more in
strain DT3 (39.8%) than S31 (31.4%). Candida rugosa lipase was used to concentrate
DHA in the glyceride portion. Initially, the lipase concentration was optimised and it 134
Chapter 8 was found that 3000 U resulted in 50% oil hydrolysis in 3 hours. The presence of lipid
classes FFA, DAG and MAG were observed by TLC analysis. Under optimised
conditions the omega-3 fatty acid content of thraustochytrid oil was enriched to 88.7%,
a 1.4 fold increase on initial levels.
We also used phospholipase A1, a carboxyl ester hydrolase, for concentrating
omega-3 fatty acids from Schizochytrium DT3 oil extracted from optimised bead
milling process. At 2.5 h of incubation, oil hydrolysis reached 50%, and the fatty acid
profiles were analysed using GC. EPA was equally distributed in the free fatty acid
and glyceride portions, while DHA was significantly enriched in glyceride portion,
indicating a strong selectivity for EPA over DHA.
Chapter 7 demonstrated the application of phospholipase as a catalyst for
enriching omega-3 fatty acids from anchovy fish oil. Initially we investigated the
substrate selectivity of PLA1 using p-NP esters chain lengths of C2 to C22, including
the long chain PUFAs, EPA and DHA. PLA1 showed preferential hydrolysis of EPA
and discriminated against DHA. In anchovy oil hydrolysis, the saturated and mono-
unsaturated fatty acids were hydrolysed from the glyceride portion. Of the omega-3
fatty acids, EPA was preferentially hydrolysed over DHA. We compared fish oil
hydrolysis with the lipase from Bacillus subtilis, which did not discriminate between
fatty acids to the same extent as PLA1. Furthermore, hydrolysis of the structured lipid
1, 3-dioleoyl-2-hexadecanoic confirmed that the observed selectivity of PLA1 was due
to fatty acid selectivity and not positional selectivity.
Microalgae derived biofuels and nutraceuticals are an alternative approach for
sustainable production. However, there are several challenges to be addressed. The
large scale production of lipids from microalgae is only possible with the isolation and
selection of microalgae with high biomass and lipid productivities. This can be
achieved by using robust techniques in isolation and selection of microalgae from
varied habitats for commercial scale production. Although the lipid extraction methods
explored in this thesis were optimised using dry biomass, it is necessary to explore
extraction of wet biomass, since biomass drying is an expensive process. The
development of enzyme assisted and nano particle based lipid extraction and the use
of natural solvents such as terpenes and organic-nanoclays reduce the environmental
impacts and waste generation during processing. Thraustochytrids appears to have
135
Chapter 8 some potentially useful lipases, including lipases with DHA versus DPA selectivity,
although the structures of these lipases still remain to be identified.
136
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