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Microalgal Lipids, Lipases and Lipase
Screening Methods
by
Tim D. Nalder
BSc (Hons), MSc
Submitted in fulfilment of the requirements for the degree of
Doctor of Philosophy
Deakin University
March 2014
Acknowledgments
I would firstly like to extend a sincere thank you to my supervisors Prof. Colin
Barrow and Dr. Susan Marshall. The time, support and red pen ink you have given
me has been greatly appreciated. Furthermore, your guidance and willingness to not
only listen to my ideas, but allow me to follow them through has enabled me to grow
as a researcher. I look forward to the continuation of our work in the future.
To my darling wife Lynda, a few words here don’t do justice to how much your
unconditional love and support means to me. Without you I doubt this would have
been possible. To the rest of my whānau, you guys are simply the best and your
ongoing support makes life all the easier. I know you’ll be looking forward to being
able to say your son/brother is something other than a student!
My genuine thanks also go out to others who have aided me in my research. A
special thank you must be given to Dr. Ivan Kurtovic and Dr. Matt Miller who have
provided sound advice on lipase and lipid related work. Nicky Roughton, Cara
McGreggor and Dr. Mike Packer (Cawthron Institute) have been wonderful to deal
with and instrumental in the culture of microalgae. Help from Dr. Fred Pfeffer, Gail
Dyson, Dr. Pimm Vongsvivut and Dr. Jacqui Adcock was invaluable to me in the
chemical synthesis and characterisation of pNP-esters.
A big thank you must also be given to the other students and postdocs who have
worked alongside me. To those who helped me along the way, whether it was advice
given or just a laugh, I am grateful. I would particularly like to thank the late Kathrin
Stähler. I still can’t fathom your early passing, but I have only fond memories of the
time spent working with you and the algae (more you than the algae).
To the Plant and Food Research group, thank you for making me feel so welcome. I
made some great friendships during my stay at PFR, many of which I’m sure will
continue in the future. Likewise, to the group of misfit chemists upstairs at Deakin,
thanks for the all social banter and numerous events known simply as ‘slurrrpage’.
Finally I would like to thank both Deakin University and Plant and Food Research.
The facilities provided have allowed me to take the next step in my career. I must
also thank you for the funding provided by the MSI programme (C02X0806) (New
Zealand) and various internal funding streams from Deakin University (Australia).
i
Published documents that have resulted from this work
Miller, M. R., Quek, S. Y., Staehler, K., Nalder, T. D. & Packer, M. A. (2012)
Changes in oil content, lipid class and fatty acid composition of the microalga
Chaetoceros calcitrans over different phases of batch culture. Aquaculture Research.
DOI: 10.1111/are.12107
Nalder, T. D., Marshall, S., Pfeffer, F. P. & Barrow, C. J. (2014) Characterisation of
lipase fatty acid selectivity using novel omega-3 pNP-acyl esters. Journal of
Functional Foods. vol. 6, pg. 259-269
ii
Abstract
The long chain omega-3 polyunsaturated fatty acids (LC n-3 PUFAs)
eicosapentaenoic acid (EPA) and docosahexaenoic acid (DHA) are physiologically
important lipids that are vital to good health. EPA and DHA in their natural form and
as concentrates are of commercial interest and generate a multi-billion dollar market
annually. Conventional concentration methods are often costly and use procedures
that may damage LC n-3 PUFAs. Alternative methods using lipases offer several
advantages, such as milder processing conditions and limited solvent/chemical usage.
Lipases are used extensively as versatile biocatalysts in many industrial applications,
offering a ‘green’ approach to chemical processes. Because of these applications, the
discovery of novel lipases is commercially important.
The microalgae Chaetoceros calcitrans, Isochrysis sp. (T.ISO) and Pavlova lutheri
were investigated as sources of lipases with potentially useful activities. Microalgae
were batch cultured, their growth profiles determined and the lipid content and
composition of each species analysed. The analyses provide a more in-depth
investigation of lipid composition in these species than previously reported. Neutral
lipid was the major lipid class in C. calcitrans and Isochrysis sp., while glycolipid
was most prevalent in P. lutheri. LC n-3 PUFA content of the three species was
highest in early culture, between lag and logarithmic growth. For lipase isolation, C.
calcitrans, Isochrysis sp. and P. lutheri were grown for approximately 7, 9 and 19
days, respectively.
Protein homogenates from Isochrysis sp. and P. lutheri were found to contain lipase
activity which was calcium-dependent. Multiple lipolytic enzymes were observed in
each species. Purification of the lipases from the homogenates by ammonium
sulphate precipitation and ion-exchange chromatography was not achieved.
Hydrophobic interaction chromatography using Sepharose and Toyopearl resins was
found to be the most promising isolation technique, with lipases adsorbed
specifically. Preliminary biochemical characterisation showed that the pH and
temperature optima of the crude lipase preparations from Isochrysis sp. and P. lutheri
were 7.0/55 C and 5.0-6.0/30 C, respectively. The fatty acid selectivity of the lipases
from these organisms was relatively broad, with optimal hydrolysis observed against
pNP-myristate (C14:0) when profiled with an extended pNP-acyl ester library.
iii
Simple, high-throughput assays are required in order to screen lipases for desirable
activities. Synthetic substrates, such as pNP-acyl esters can be used for this purpose.
However, a comprehensive range of substrates with differing chain lengths and bond
saturation is not available commercially. Therefore, we synthesised six olefinic pNP-
acyl esters (derivatives of C16:1 n-7, C18:1 n-9, C18:2 n-6, C18:3 n-3, C20:5 n-3
(EPA) and C22:6 n-3 (DHA)). These were used in conjunction with commercial
esters and provided a substrate library for the characterisation of lipase FA
selectivity. The modified Steglich esterification reactions used for the synthesis of
substrates gave yields >85%. The synthesised compounds were chemically
characterised and found to be stable under both assay and storage conditions. Five
commercial lipases were profiled against the pNP-acyl esters. Lipase B from
Candida antarctica and Aspergillus niger lipase were the least selective, and
Thermomyces lanuginosus and Rhizomucor miehei lipases were inefficient at
hydrolysing olefinic substrates. Candida rugosa lipase was found to be the most
selective towards EPA and DHA. Lipase profiles were consistent with respect to
active site structure-function relationships.
Previously unreported, calcium-stimulated activity was observed for two of the five
commercial lipases tested. Lipases with cleft-type (but not funnel- or tunnel-type)
active site conformations (Thermomyces lanuginosus and Rhizomucor miehei
lipases) were stimulated by calcium. However, calcium-stimulation was only
observed against certain emulsion compositions, namely triacylglycerol and
phospholipid mixtures. In the presence of calcium, the activity of the Thermomyces
lanuginosus lipase against tributyrin and lecithin increased by 170 and 46 fold,
respectively. Calcium was found to modify the physicochemical properties of the
emulsions by reducing surface charge and increasing micelle size distribution.
The scope of the work carried out in this project is relatively broad. As a whole it
provides new insights into the lipids and lipases in microalgae. The development of a
pNP substrate screening library is a useful tool for the selection of promising new
lipases. The demonstration of previously undescribed calcium-stimulated lipase
activities in some commercial lipases indicates that it might be possible to
manipulate and/or selectively enhance the hydrolysis of different lipid substrates.
iv
Table of Contents
Published documents that have resulted from this work ........................................ i
Abstract ....................................................................................................................... ii
Table of Contents ....................................................................................................... iv
List of figures ............................................................................................................. ix
List of tables ............................................................................................................... xi
List of abbreviations ................................................................................................ xiii
CHAPTER I General introduction and research objectives ............................... 1
1.0 Introduction .................................................................................................... 1
1.1 Long chain omega-3 polyunsaturated fatty acids ........................................... 2
1.1.1 LC n-3 PUFAs in human health ........................................................... 4
1.1.2 Sources of LC n-3 PUFAs .................................................................... 6
1.1.3 LC n-3 PUFA processing and concentration ........................................ 7
1.2 The potential of algae ..................................................................................... 9
1.2.1 Microalgal applications and biotechnology ........................................ 10
1.2.2 Microalgae and LC n-3 PUFA metabolism ........................................ 11
1.3 Lipases .......................................................................................................... 14
1.3.1 Lipase structure and function .............................................................. 14
1.3.2 Lipase sources ..................................................................................... 19
1.3.3 Lipase isolation and purification ........................................................ 21
1.3.4 Lipase-based omega-3 concentration ................................................. 22
1.4 Methods for lipase detection and characterisation ....................................... 24
1.4.1 Thin layer chromatography/Iatroscan ................................................. 24
1.4.2 Chromatography ................................................................................. 25
1.4.3 Titrimetric assays ................................................................................ 26
1.4.4 Colorimetric and fluorometric assays ................................................. 26
1.5 p-Nitrophenyl assay ..................................................................................... 28
1.6 Research objectives ...................................................................................... 30
CHAPTER II Materials and methods ................................................................. 31
2.0 Materials ....................................................................................................... 31
2.0.1 Chemicals ........................................................................................... 31
v
2.0.2 Proteins and standards ........................................................................ 32
2.0.3 Lipids and substrates ........................................................................... 32
2.0.4 Buffers, reagents, solvent systems and emulsions .............................. 33
2.1 Methods ........................................................................................................ 36
2.1.1 Culture of microalgae ......................................................................... 36
2.1.2 Algal lipid extraction and determination of total lipids ...................... 37
2.1.3 Separation and determination of lipid classes .................................... 37
2.1.4 Lipid methylation and determination by gas chromatography ........... 38
2.1.5 Algal cell lysis and protein extraction ................................................ 39
2.1.5.1 Bead-beating ....................................................................................... 39
2.1.5.2 Homogenisation by Ultra-Turrax ....................................................... 40
2.1.6 Freeze-drying of algal protein homogenates ...................................... 40
2.1.7 Lowry protein concentration assay ..................................................... 40
2.1.8 p-Nitrophenyl activity assay ............................................................... 41
2.1.9 Titrimetric activity assay .................................................................... 42
2.1.10 4-Methylumbelliferyl butyrate activity assay ..................................... 43
2.1.11 1D SDS-PAGE ................................................................................... 44
2.1.12 Non-denaturing PAGE (for proteins pI ≤ 7.0) .................................... 45
2.1.13 Acidic non-denaturing PAGE (for proteins pI 7.0) ......................... 45
2.1.14 Gel-overlay activity assay ................................................................... 46
2.1.15 Ammonium sulfate precipitation ........................................................ 46
2.1.16 Anion-exchange chromatography ....................................................... 47
2.1.17 Cation-exchange chromatography ...................................................... 47
2.1.18 Hydrophobic interaction chromatography .......................................... 48
2.1.19 p-Nitrophenyl-acyl ester synthesis ..................................................... 48
2.1.20 Thin layer chromatography of pNP-acyl esters .................................. 49
2.1.21 Silica gel chromatography of pNP-acyl esters .................................... 49
2.1.22 Yield calculations of pNP-acyl esters ................................................. 50
2.1.23 Mass spectrometry of pNP-acyl esters ............................................... 50
2.1.24 UV spectroscopy of pNP-acyl esters .................................................. 50
2.1.25 FTIR of pNP-acyl esters ..................................................................... 51
2.1.26 1H and 13C NMR of pNP-acyl esters .................................................. 51
2.1.27 Reverse-phase high performance liquid chromatography of pNP-acyl esters ................................................................................................... 52
vi
2.1.28 Iatroscan analysis of lecithin and lecithin hydrolysis products .......... 52
2.1.29 Analysis of micelle surface charge ..................................................... 53
2.1.30 Analysis of micelle size distribution ................................................... 53
CHAPTER III Growth and lipid composition of Chaetoceros calcitrans, Isochrysis sp. and Pavlova lutheri ................................................ 54
3.0 Introduction .................................................................................................. 54
3.1 Growth of Chaetoceros calcitrans, Isochrysis sp. and Pavlova lutheri ....... 56
3.2 Lipid content and class in Chaetoceros calcitrans, Isochrysis sp. and Pavlova lutheri ............................................................................................. 58
3.2.1 Lipid content and classes of Chaetoceros calcitrans ......................... 60
3.2.2 Lipid content and classes of Isochrysis sp. ......................................... 62
3.2.3 Lipid content and classes of Pavlova lutheri ...................................... 64
3.3 Fatty acid composition of Chaetoceros calcitrans, Isochrysis sp. and Pavlova lutheri ............................................................................................. 66
3.3.1 Fatty acid composition of Chaetoceros calcitrans ............................. 66
3.3.2 Fatty acid composition of Isochrysis sp. ............................................. 68
3.3.3 Fatty acid composition of Pavlova lutheri .......................................... 70
3.4 LC n-3 PUFA content in the neutral lipid of Chaetoceros calcitrans, Isochrysis sp. and Pavlova lutheri ............................................................... 72
3.5 Summary ...................................................................................................... 78
CHAPTER IV Investigation of lipases from Isochrysis sp. and Pavlova lutheri ........................................................................................................ 79
4.0 Introduction .................................................................................................. 79
4.1 Detection of lipases in microalgae ............................................................... 80
4.2 Algal cell lysis methods ............................................................................... 83
4.3 Calcium-dependence of lipase activity ........................................................ 85
4.4 Storage of algal protein homogenates .......................................................... 86
4.5 Lipase purification by precipitation and ion-exchange chromatography ..... 88
4.5.1 Ammonium sulfate precipitation ........................................................ 88
4.5.2 Anion-exchange chromatography ....................................................... 89
4.5.3 Acidic native-PAGE analysis of P. lutheri extract ............................. 92
4.5.4 Cation-exchange chromatography ...................................................... 93
4.6 Lipase purification by hydrophobic interaction ........................................... 95
4.6.1 Hydrophobic interaction with Sepharose® resins ............................... 96
4.6.2 Hydrophobic interaction with Toyopearl resin ................................. 98
vii
4.7 Enzymatic analysis of lipase-containing crude material ............................ 100
4.7.1 Molecular weight analysis by multi-PAGE and mass spectrometry 100
4.7.2 Effect of temperature on lipase hydrolysis ....................................... 102
4.7.3 Effect of pH on lipase hydrolysis ..................................................... 104
4.7.4 Fatty acid selectivity of algal homogenates ...................................... 105
4.8 Summary .................................................................................................... 107
CHAPTER V Novel omega-3 pNP-acyl esters for the characterisation of lipase fatty acid selectivity .................................................................... 109
5.0 Introduction ................................................................................................ 109
5.1 pNP-acyl ester synthesis protocol development and optimisation ............. 111
5.2 Synthesis of novel olefinic pNP-acyl esters ............................................... 112
5.3 Characterisation of synthesised pNP-acyl esters ........................................ 115
5.3.1 Mass spectrometry and UV spectroscopy ......................................... 117
5.3.2 Fourier transform infrared spectroscopy .......................................... 117
5.3.3 1H and 13C NMR ............................................................................... 118
5.4 Stability analyses of synthesised pNP-acyl esters ...................................... 120
5.4.1 Storage stability analyses .................................................................. 121
5.4.2 Assay stability analyses .................................................................... 124
5.5 Screening of lipase fatty acid selectivity using pNP-acyl esters ................ 125
5.5.1 Fatty acid selectivity profiles of Candida antarctica lipase B and Aspergillus niger lipase .................................................................... 126
5.5.2 Fatty acid selectivity profile of Candida rugosa lipase .................... 128
5.5.3 Fatty acid selectivity profiles of Thermomyces lanuginosus and Rhizomucor miehei lipases ............................................................... 129
5.6 Observations from lipase activity profiles ................................................. 132
5.7 Summary .................................................................................................... 134
CHAPTER VI Calcium-stimulated activity in fungal lipases with cleft-type active sites ................................................................................... 135
6.0 Introduction ................................................................................................ 135
6.1 Increased pNP-acyl ester hydrolysis in the presence of calcium ............... 137
6.2 Calcium alters hydrolysis rate and not specificity ..................................... 139
6.3 Effect of calcium on hydrolysis of TAG substrates by lipases .................. 142
6.3.1 Effect of calcium on tributyrin hydrolysis ........................................ 142
6.3.2 Effect of calcium on hydrolysis of fish oil by lipases ...................... 143
6.4 Effect of calcium on hydrolysis of lecithin by lipases ............................... 144
viii
6.5 Analysis of lecithin and lecithin hydrolysis products ................................ 146
6.7 Analysis of emulsion properties ................................................................. 148
6.8 Summary .................................................................................................... 151
CHAPTER VII Conclusions and future directions .......................................... 152
BIBLIOGRAPHY .................................................................................................. 158
VIII APPENDICES ............................................................................................ 178
Appendix 8.1 Fatty acid profiles of Chaetoceros calcitrans, Isochrysis sp. and Pavlova lutheri and their lipid classes ......................................... 178
Appendix 8.2 Solvents trialled for gel-overlay assays with 4-methylumbelliferyl palmitate ....................................................................................... 190
Appendix 8.3 Ion-exchange chromatography of Isochrysis sp. protein homogenate .................................................................................. 190
Appendix 8.4 Acidic and basic native-PAGE band analysis .............................. 193
Appendix 8.5 Sequences identified by MALDI TOF/TOF ................................ 194
Appendix 8.6 Extinction coefficients of p-nitrophenol for defining lipase pH optima........................................................................................... 194
Appendix 8.7 13C NMR of fatty acids used in pNP-acyl ester synthesis............ 195
Appendix 8.8 pNP-acyl ester profiles for CalB, TlL and RmL without CaCl2 .. 196
Appendix 8.9 Titrimetric assay trace of TlL against lecithin ............................. 197
ix
List of figures
Figure 1.1 Chemical structures of EPA (C20:5 n-3) and DHA (C22:6 n-3). .... 3
Figure 1.2.2 Biosynthesis of LC n-3 PUFAs in eukaryotic microalgae via
enzymatic elongation and desaturation. ......................................... 12
Figure 1.3.1 Secondary structure of the ‘canonical’ /β hydrolase fold. ........... 16
Figure 1.3.2 Basic representation of lipase lid conformational change and
interaction with hydrophobic surfaces. .......................................... 17
Figure 1.3.3 Active site topology of lipase B from C. antarctica, R. miehei
lipase and C. rugosa lipase. ........................................................... 18
Figure 3.1 Growth curves of C. calcitrans, Isochrysis sp. and P. lutheri ....... 57
Figure 3.2 Chemical structures of triacylglycerol (A), phosphatidylcholine (B)
and monogalactosyldiacylglycerol (C). ......................................... 59
Figure 3.2.1 Lipid class composition over the growth of C. calcitrans. ............ 61
Figure 3.2.2 Lipid class composition over the growth of Isochrysis sp.. ........... 63
Figure 3.2.3 Lipid class composition over the growth of P. lutheri. .................. 65
Figure 3.4.1 EPA and DHA content in the total lipid and neutral lipid of C.
calcitrans over the duration of growth. .......................................... 73
Figure 3.4.2 EPA and DHA content in the total lipid and neutral lipid of
Isochrysis sp. over the duration of growth. .................................... 74
Figure 3.4.3 EPA and DHA content in the total lipid and neutral lipid of P.
lutheri over the duration of growth. ............................................... 76
Figure 4.1 4-Methylumbelliferyl butyrate gel-overlay assay performed on a
basic native-PAGE ......................................................................... 82
Figure 4.2.1 P. lutheri cells before and after bead-beating. ............................... 83
Figure 4.4 Relative activities against pNPP for P. lutheri protein homogenate
before and after freeze-drying. ....................................................... 87
Figure 4.5.2 Anion-exchange chromatography of the protein homogenate from
P. lutheri......................................................................................... 90
Figure 4.5.3 4-Methylumbelliferyl butyrate gel-overlay assay on an acidic
native-PAGE of P. lutheri protein homogenate. ............................ 92
Figure 4.5.4 Cation-exchange chromatography of the protein homogenate from
P. lutheri......................................................................................... 94
Figure 4.6.1 Relative and specific lipase activities of the Sepharose adsorption
fractions against pNP-acyl esters. .................................................. 97
x
Figure 4.6.2 Relative and specific lipase activities of the fractions from
Toyopearl adsorption against pNP-acyl esters. .............................. 99
Figure 4.7.1 SDS-PAGE of fluorescent bands cut from native-PAGE of P.
lutheri protein homogenate. ......................................................... 101
Figure 4.7.2 Effect of temperature on the activity of protein homogenates from
Isochrysis sp. and P. lutheri. ........................................................ 103
Figure 4.7.3 Effect of pH on the activity of protein homogenates from Isochrysis
sp. and P. lutheri. ......................................................................... 104
Figure 4.7.4 Activity profiles of protein homogenates from Isochrysis sp. and P.
lutheri against a range of pNP-acyl esters. .................................. 106
Figure 5.2.1 Reaction mechanism of the Steglich esterification with EDCI. ... 113
Figure 5.2.2 Synthesis, complete structures and yields of pNP-acyl esters. .... 114
Figure 5.3.2 ATR-FTIR analyses of the synthesised pNP-acyl esters. ............ 118
Figure 5.4.1.1 Reverse phase-HPLC of mono-/di-unsaturated pNP-acyl esters. 122
Figure 5.4.1.2 Reverse phase-HPLC of omega-3 pNP-acyl esters. ..................... 123
Figure 5.5.1 Activity profiles depicting the relative activity of CalB and AnL
against a range of pNP-acyl esters. .............................................. 127
Figure 5.5.2 Activity profile depicting the relative activity of CrL against a
range of pNP-acyl esters. ............................................................. 129
Figure 5.5.3 Activity profiles depicting the relative activity of TlL and RmL
against a range of pNP-acyl esters. .............................................. 131
Figure 6.1 Activity of commercial lipases against pNP-acyl esters of medium
chain-length in the presence and absence of CaCl2 and NaCl. .... 138
Figure 6.2 Activity profiles depicting the relative activity of CalB, TlL and
RmL against a range of pNP-acyl esters with and without CaCl2.
...................................................................................................... 141
Figure 6.7 Weighted-mean of micelle size distribution in lecithin emulsions
with and without CaCl2. ............................................................... 150
Figure 8.3.1 Anion-exchange chromatography of the protein homogenate from
Isochrysis sp.. ............................................................................... 191
Figure 8.3.2 Cation-exchange chromatography of the protein homogenate from
Isochrysis sp.. ............................................................................... 192
Figure 8.4 MUB gel-overlay assay of P. lutheri protein homogenate separated
by acidic and basic native-PAGE................................................. 193
xi
Figure 8.8 Activity profiles of CalB, TlL and RmL against a range of pNP-
acyl esters in the absence of CaCl2. ............................................. 196
Figure 8.9 STAT-pH trace from a titrimetric assay of TlL against a lecithin
emulsion with CaCl2 added after assay initiation. ....................... 197
List of tables
Table 1.1 Common n-3 PUFAs found in nature .............................................. 3
Table 4.2.2 Lipase activity and protein concentration of homogenate obtained
via homogenisation and bead-beating of P. lutheri cells. .............. 84
Table 4.3 Lipase activity against pNPP from microalgal protein homogenates
with different assay buffers. ........................................................... 86
Table 4.5.1 Specific activity of ammonium sulfate precipitated fractions from
Isochrysis sp. protein homogenate. ................................................ 89
Table 5.3.3 The observed 13C NMR chemical shifts (δ) of synthesised pNP-acyl
esters............................................................................................. 119
Table 6.3.1 Activity of lipases on tributyrin in the presence and absence of
CaCl2. ........................................................................................... 143
Table 6.3.2 Activity of lipases on tuna oil in the presence and absence of
CaCl2. ........................................................................................... 144
Table 6.4 Activity of lipases on lecithin in the presence and absence of CaCl2.
...................................................................................................... 145
Table 6.5 Composition (%) of lecithin before and after reaction with lipases.
...................................................................................................... 147
Table 8.1.1 Changes in total lipid fatty acid composition (%) of Chaetoceros
calcitrans over growth. ................................................................ 178
Table 8.1.2 Changes in neutral lipid fatty acid composition (%) of Chaetoceros
calcitrans over growth. ................................................................ 179
Table 8.1.3 Changes in glycolipid fatty acid composition (%) of Chaetoceros
calcitrans over growth. ................................................................ 180
Table 8.1.4 Changes in polar lipid fatty acid composition (%) of Chaetoceros
clacitrans over growth. ................................................................ 181
xii
Table 8.1.5 Changes in total lipid fatty acid composition (%) of Isochrysis sp.
over growth. ................................................................................. 182
Table 8.1.6 Changes in neutral lipid fatty acid composition (%) of Isochrysis
sp. over growth. ............................................................................ 183
Table 8.1.7 Changes in glycolipid fatty acid composition (%) of Isochrysis sp.
over growth. ................................................................................. 184
Table 8.1.8 Changes in polar lipid fatty acid composition (%) of Isochrysis sp.
over growth. ................................................................................. 185
Table 8.1.9 Changes in total fatty acid composition (%) of Pavlova lutheri over
growth. ......................................................................................... 186
Table 8.1.10 Changes in neutral lipid fatty acid composition (%) of Pavlova
lutheri over growth....................................................................... 187
Table 8.1.11 Changes in glycolipid fatty acid composition (%) of Pavlova
lutheri over growth....................................................................... 188
Table 8.1.12 Changes in polar lipid fatty acid composition (%) of Pavlova
lutheri over growth....................................................................... 189
Table 8.2 Solvents tested for use in 4-methylumbelliferyl palmitate gel-
overlay assay ................................................................................ 190
Table 8.6 Extinction coefficients of p-nitrophenol at different pH. ............. 194
Table 8.7 The observed 13C NMR chemical shifts (δ) of free fatty acids. ... 195
xiii
List of abbreviations
1D one dimensional
ALA -linolenic acid
AnL Aspergillus niger lipase
AQIS Australian Quarantine and Inspection Service
ATP adenosine triphosphate
ATR Attenuated total reflectance
BHT butylated hydroxytolulene
BSA bovine serum albumin
CalB Candida antactica lipase B
CPR combined protein reagent
CrL Candida rugosa lipase
CSIRO Commonwealth Scientific and Industrial Research Organisation
Da Daltons
DAG diacylglycerol
DCC N,N'-dicyclohexylcarbodiimide
DCM dichloromethane
DEAE diethylaminoethanol
DHA docosahexaenoic acid
DMAP 4-dimethylaminopyridine
DPA docosapentaenoic acid
DTT dithiothreitol
EDCI 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide
EDTA ethylenediaminetetraacetic acid
EPA eicosapentaenoic acid
FA fatty acid
FAME fatty acid methyl ester
FF fast flow
FFA free fatty acid
FID flame ionisation detector
FPLC fast performance liquid chromatography
FPS Folin-phenol solution
FSW filtered seawater
FTIR Fourier transform infrared
xiv
GC gas chromatography
GC-MS gas chromatography-mass spectrometry
GL glycolipid
GOED Global Organisation for EPA and DHA Omega-3
HPLC high performance liquid chromatography
LA linoleic acid
LC long chain
MAG monoacylglycerol
MALDI matrix assisted laser desorption/ionization
MES 2-(N-morpholino) ethanesulfonic acid
MQ Milli Q
MS mass spectrometry
MUB 4-methylumbelliferyl butyrate
MUFA monounsaturated fatty acid
MUP 4-methylumbelliferyl palmitate
MWCO molecular weight cut-off
n.d not detected
n-3 omega-3
n-6 omega-6
NL neutral lipid
NMR nuclear magnetic resonance
OA oleic acid
PAGE polyacrylamide gel electrophoresis
pI isoelectric point
PKS polyketide synthase
PL polar lipid
pNP para-nitrophenol
pNPB para-nitrophenyl butyrate
pNPM para-nitrophenyl myristate
pNPP para-nitrophenyl palmitate
ppm parts per million
PUFA polyunsaturated fatty acid
Q quaternary amine
RmL Rhizomucor miehei lipase
xv
RuBisCO ribulose-1,5-bisphosphate carboxylase/oxygenase
SCO single cell oil
SDA stearidonic acid
SDS sodium dodecyl sulfate
sn stereospecificity number
SP sulphopropyl
SPE solid phase extraction
TAG triacylglycerol
TEMED tetramethylethylenediamine
TFA trifluoroacetic acid
T.ISO Tahitian isolate
TL total lipid
TLC thin layer chromatography
TlL Thermomyces lanuginosus lipase
TMS tetramethylsilane
TOF time of flight
UV ultra violet
CHAPTER I
1
CHAPTER I
General introduction and research objectives
1.0 Introduction
The omega-3 fatty acids eicosapentaenoic acid (EPA) and docosahexaenoic acid
(DHA) are amongst the most widely researched lipids today. In 2012 the estimated
global consumer spending on supplements, drug and food products containing these
compounds was US$25.4 billion (Ismail 2013). The expansion of this market is
largely due to evidence supporting the various health benefits associated with the
consumption of long chain omega-3 polyunsaturated fatty acids (LC n-3 PUFAs).
Demand for these compounds will only increase in the future as the world’s
population continues to grow. Several factors need to be taken into account regarding
the methods utilised for the concentration of LC n-3 PUFAs. Most important is the
efficacy and the quality of products produced by different methods. A range of
conventional methods are available. However, many have significant drawbacks with
respect to high cost, chemical/solvent usage and process conditions that are
detrimental to the LC n-3 PUFAs themselves. A method that has been gaining
interest is the use of lipases. Lipases offer several advantages such as, allowing the
process to be carried out under milder conditions, limiting solvent and chemical
waste products and being more specific for the targeted concentration of EPA and
DHA. Previously characterised lipases have already been shown to have omega-3
concentrating abilities. However, there is huge potential for the discovery of novel
lipases with more specific activities that may be better suited for the concentration of
these highly desirable lipids.
Marine microalgae represent a source of lipases with potentially useful activities
towards LC n-3 PUFAs. Many microalgal species are capable of biosynthesising
large quantities of LC n-3 PUFAs; particularly EPA and DHA. It is therefore
possible that lipases in these organisms have evolved activities relevant to the
hydrolysis of LC n-3 PUFAs. Microalgae are also an interesting source of lipases
because although the presence of the enzymes is known, to date no lipases have been
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isolated and characterised from native sources. Thus they represent a relatively
unexplored enzyme resource.
To aid in the isolation of new lipases, the development/improvement of methods for
characterising them is also important. The detection and characterisation of lipases
suitable for the concentration of LC n-3 PUFAs is slowed by the conventional
methods used to analyse activity. While providing comprehensive analyses,
conventional methods are time consuming and can require substantial amounts of
enzyme. Therefore, the development of more sensitive and rapid assays for the
screening of LC n-3 PUFA-specific activity is advantageous. Various
spectrophotometric methods allow high-throughput screening, but they do not
include the analysis of omega-3 specific activities. The use of these assay platforms
with new LC n-3 PUFA substrates would provide a useful tool.
1.1 Long chain omega-3 polyunsaturated fatty acids
The LC n-3 PUFAs EPA and DHA are of biological significance to both the
organisms that synthesise them and to those that rely on them through dietary intake.
The structure and properties associated with LC n-3 PUFA make them vital in
numerous important biological functions and as such they have gained popular
acceptance for their role in cellular function and health. For these reasons, EPA and
DHA are the two most widely studied LC n-3 PUFA. The term ‘omega-3’ refers to
the presence of a double bond that includes the third carbon atom from the methyl, or
‘omega’ terminus, of the acyl chain. Naturally occurring omega-3 FAs are PUFAs
with methylene-interrupted double bonds, all of which are in the cis conformation.
Table 1.1 presents the most common omega-3 FAs found in nature. The high degree
of unsaturation in EPA and DHA (Figure 1.1) make them more prone to chemical
alteration and degradation than shorter, more saturated FAs. Chemical alterations
such as oxidation, geometric isomerisation and polymerisation can take place and are
damaging to the beneficial properties associated with omega-3s (Shahidi &
Wanasundara 1998); for example, DHA’s role in modifying membrane structure and
function (Stillwell & Wassall 2003). These unfavourable reactions are of particular
importance when EPA and DHA are processed in commercial settings and are
discussed further in Section 1.1.3.
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Table 1.1 Common n-3 PUFAs found in nature
Common name
(trivial nomenclature)
Lipid number
(shorthand designation)
-Linolenic acid (ALA) 18:3 n-3
Stearidonic acid (SDA) 18:4 n-3
Eicosatrienoic acid 20:3 n-3
Eicosatetraenoic acid 20:4 n-3
Eicosapentaenoic acid (EPA) 20:5 n-3
Docosapentaenoic acid (DPA) 22:5 n-3
Docosahexaenoic acid (DHA) 22:6 n-3
Figure 1.1 Chemical structures of EPA (C20:5 n-3) and DHA (C22:6 n-3).
(1) EPA and (2) DHA.
Due to the physical nature of these FAs, membranes containing them are more fluid
than those composed of more saturated FAs (Stillwell et al. 2003). Membrane
fluidity is thought to be one of the major functions of LC n-3 PUFAs in organisms
which produce them, such as microalgae. Another quality of LC n-3 PUFAs is
flexibility. The acyl chains of LC PUFA are disordered and become increasingly so
as unsaturation increases. It has been shown that DHA may be present in up to 100
conformations at any given time and furthermore, that the acyl chain may rapidly
change between different conformations (Feller et al. 2002; Gawrisch et al. 2003).
This ability is also associated with membrane dynamics and fluidity and influences
the function of membrane-bound proteins (Feller & Gawrisch 2005; Feller et al.
2002; Hyvonen & Kovanen 2005). The presence of DHA in phospholipid
membranes has also been linked to membrane permeability (Armstrong et al. 2003).
1
2
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4
All of the above properties demonstrate that LC n-3 PUFAs, such as DHA, are
highly functional compounds that are important for proper cellular health. Examples
of LC n-3 PUFA functions are diverse and are mostly researched in higher organisms
(that obtain the compounds from the diet), with much research has taking place in
humans.
1.1.1 LC n-3 PUFAs in human health
The notion that EPA and DHA play a role in health was first suggested after research
on Greenlandic Inuit (Bang & Dyerberg 1972; Bang et al. 1971). This work reported
that despite the consumption of a high fat diet, similar in volume to that of Western
society, the incidence of health problems such as cardiovascular disease were almost
non-existent in the Inuit population. This seemingly contradictory evidence
suggested that LC n-3 PUFAs had some protective effect (Dyerberg et al. 1975).
Since these early findings an overwhelming body of evidence now supports the
positive health benefits associated with these compounds. One of the most notable
observations in those initial studies was that the ratio of omega-6 to omega-3 FA
differed between populations, with Western diets consisting of more omega-6
(Simopoulos 2002b). Estimates of the Western diet have put the ratio of omega-6 FA
to omega-3 at 15-16.7:1 indicating a major deficiency in omega-3 FA, which is
thought to promote the pathogenesis of many chronic diseases found in Western
society today (Simopoulos 2002a). Research has since demonstrated that to a large
extent, the ratio of the two PUFA families is important (Simopoulos 2002a, 2008),
with LC n-3 PUFA providing a suppressive effect of disease pathogeneses that
would otherwise occur.
The most widely documented influence that omega-3 FAs have on human health is
the multitude of ways in which they improve cardiovascular function. This includes,
but is not limited to, reducing the incidence of arrhythmias (Chrysohoou et al. 2007;
Dallongeville et al. 2003; Geelen et al. 2005) decreasing blood pressure (Appel et al.
1993; Bonaa & Thelle 1991; Toft et al. 1995), decreasing platelet aggregation
(Goodnight et al. 1981; Tremoli et al. 1995), lowering plasma triacylglycerol (TAG)
levels (Herrmann et al. 1995; Lovegrove et al. 2004; Sanders et al. 2006), increasing
high density lipoprotein cholesterol (Chan et al. 2006; Herrmann et al. 1995;
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5
Lovegrove et al. 2004) and improved vasodilation (Fleischhauer et al. 1993; Morgan
et al. 2006). All of the above factors may be linked directly or indirectly to reducing
cardiac related death.
Numerous other examples of health benefits are also apparent, including effects on
neurological and visual function. It has been demonstrated that the risk of poor
neural development in infants is increased with insufficient DHA in the diet
(Dunstan et al. 2008; Helland et al. 2003; Hibbeln et al. 2007). Likewise insufficient
DHA intake has been associated with various neurological disorders in older
individuals, including conditions such as dementia, Alzheimer’s disease (Cole et al.
2009; Schaefer et al. 2006) and other neuronal disorders like Parkinson’s disease
(Bousquet et al. 2011). DHA has also been implicated in visual function and is an
essential component of the retina (SanGiovanni & Chew 2005). Further examples of
the benefits of LC n-3 PUFAs in eye health can be seen in the meta-analysis by
Chong et al. (2008), where DHA consumption resulted in a 38% reduction in the on-
set of age-related macular degeneration.
Research on the downstream products, of which EPA and DHA are precursors, is
also increasing. Perhaps the most prominent example of this is in the role LC PUFAs
play in inflammation and autoimmune disorders. Inflammation is relevant to a
number of medical conditions, a prime example being rheumatoid arthritis. Research
has shown that joint inflammation is reduced when dietary intake of LC n-3 PUFAs
is increased (Calder 2008; Proudman et al. 2008). EPA, DHA and the omega-6
PUFA arachidonic acid are converted into molecules known as eicosanoids and
cytokines. These go on to form various different compounds such as prostaglandins,
prostacyclins, thromboxanes, leukotrienes and lipoxins (Serhan et al. 2008). Omega-
3 derived molecules play important roles in the resolution of inflammation, while
molecules originating from omega-6 FA are pro-inflammatory (Serhan et al. 2008).
Arachidonic acid and EPA directly compete for access to lipoxygenase and
cyclooxygenase enzymes (Simopoulos 2002b), thus the ratio of omega-6 to omega-3
LC PUFAs is of great importance. This further supports the need for a balanced diet
in regard to omega-3 and omega-6 intake.
The evidence above supports the beneficial effects of dietary intake of LC n-3
PUFA. With demand for these compounds rising, commercial interest in them has
also increased. The omega-6 (n-6) PUFA-rich nature of Western food, coupled with
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6
a lack of sufficient seafood consumption, means that omega-3 intake is often
achieved through supplements. To increase their potency these products usually
contain LC n-3 PUFA concentrates.
1.1.2 Sources of LC n-3 PUFAs
The most common sources of omega-3 PUFAs are oils derived from marine and
plant sources. Currently the most widely used and abundant commercial source of
EPA and DHA is fish oil. Other common marine sources include squid, krill and
microalgae (Nichols et al. 2010). Oil from marine mammals such as seals is also rich
in LC n-3 PUFA, including the omega-3 docosapentaenoic acid (DPA) (Conquer et
al. 1999). Fish, like most other higher organisms, have a limited ability to synthesise
LC n-3 PUFA themselves. Therefore EPA and DHA are accessed primarily through
dietary intake (Dunstan et al. 1993; Kanazawa et al. 1979). Microalgae form the base
of the food-web and are the primary producers of these compounds. Accordingly,
nearly all LC n-3 PUFAs bioaccumulated in fish originate from these marine
microbes. A major problem faced when sourcing EPA and DHA from wild fisheries
is the finite nature of fish stocks. Unsustainable over-harvesting of these resources
leads to the collapse of fisheries (Worm et al. 2006), further compounding supply
and demand issues. Development of viable alternative sources of EPA and DHA are
required to help reduce pressure on wild fish stocks while meeting demands of
nutraceutical and aquaculture industries (Turchini et al. 2012).
Microalgal oils, sometimes referred to as ‘single cell oils’ (SCO), are one emerging
source of LC n-3 PUFAs. They represent a renewable resource that could potentially
provide a sustainable source of EPA and DHA. Development and growth in the area
is set to rise, with microbial SCOs currently holding approximately 3-5% of the
omega-3 market (Turchini et al. 2012). Krill is another emerging source of LC n-3
PUFAs and provides a phospholipid-rich oil with high concentrations of EPA and
DHA (Virtue et al. 1995). However, krill faces the same issues as other oils sourced
from wild fisheries and must be managed in a sustainable fashion (Miller et al.
2008).
The omega-3 PUFA derived from plants is usually -linolenic acid (ALA, C18:3 n-
3), with common sources including flaxseed, canola and walnut (Gebauer et al.
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7
2006). Relative to EPA and DHA, there are currently very few health benefits
directly attributed to ALA (Anderson & Ma 2009). However, as demonstrated in
early research by Burr and Burr (1930), ALA is an essential FA in humans. This is
because humans (and other mammals) cannot synthesise LC n-3 PUFAs (EPA and
DHA) de novo, requiring ALA to bypass the limiting step (Das 2006). Plants lack the
desaturase and elongase enzymes required to synthesise long chain ( C20) LC n-3
PUFA. However, in the last decade significant inroads have been made to
incorporate the genes encoding these enzymes into plants via genetic engineering
(Petrie et al. 2012; Qi et al. 2004). Technologies such as this are set to alleviate the
reliance on wild fish stocks for EPA and DHA, and will aid in supplying
nutraceutical and aquaculture demands (Petrie et al. 2012).
Regardless of the source, efficient methodologies are required for the processing and
concentration of LC n-3 PUFA oils to be used in nutraceuticals and fortified foods.
The various approaches and techniques employed to achieve concentration are
predominantly physical and or chemical in nature.
1.1.3 LC n-3 PUFA processing and concentration
The demand for oils with higher proportions of EPA and DHA by food and
nutraceutical markets means that methods for producing high quality concentrates
are of industrial importance. Concentration of these FAs is desired as it increases the
potency of products containing them, while reducing the overall saturated fat and
cholesterol intake (Davidson et al. 1991). It has been found that consumption of
biologically-effective quantities of some LC n-3 PUFA-containing oils poses a risk
of vitamin A and D overdose (Davidson et al. 1991). Likewise, the consumption of
large amounts of fish (usually large pelagic species) to obtain LC n-3 PUFAs carries
the risk of heavy metal exposure (Castro-Gonzalez & Mendez-Armenta 2008).
Concentrates allow consumption of a lower amount of oil whilst still obtaining
sufficient EPA and DHA. Methods for achieving LC n-3 PUFA concentration are
traditionally physical and chemical methods, with some of the most common
protocols outlined below.
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8
Distillation is one of the most widely applied methods for the separation of lipid
mixtures. The various distillation methods exploit differences in the volatility and
molecular weight of lipid species to affect their separation under reduced pressure
(Breivik 2007). A significant draw back to the use of distillation is the requirement of
high temperatures (250 C), which can cause LC n-3 PUFAs to undergo hydrolysis,
oxidation, polymerisation and isomerisation (Shahidi et al. 1998). Different forms of
distillation are used, such as short-path (or molecular) distillation. This variation
employs milder temperatures with short heating intervals and can be highly effective.
However, for lipid mixtures containing high molecular weight species, such as LC n-
3 PUFAs, the fraction resolution can be poor (Shahidi et al. 1998).
Winterisation (low-temperature crystallisation) is also widely used in the processing
of commercial omega-3 oils. The method separates different lipid species based on
their solubility under low temperatures. Generally the melting point of FAs and FA-
containing lipids (such as TAG) changes dramatically with the degree of unsaturation
present. The separation of mixtures is achieved at low temperatures, because long
chain saturated FAs (higher melting points) crystallise while PUFA remain in their
liquid form (Haraldsson 1984). Some winterisation protocols require the use of
organic solvents which must be dealt with subsequently.
Urea complexation works on the basis that urea forms solid complexes with straight
chain compounds, including FAs, causing crystallisation. Compounds with increased
unsaturation interact with the urea less readily and therefore most remain in the
liquid fraction of the reaction (Breivik 2007). This method does not concentrate all
PUFA to the same degree and residual amounts of LC n-3 PUFAs will remain in the
crystallised fraction. Another disadvantage of this method is the requirement for the
oil to be saponified into FA or ester components, as it is not applicable for TAG
(Breivik 2007). There is also the requirement to deal with the chemical effluent that
is produced.
More recently, techniques such as super critical fluid extraction have been
developed. This technique uses various solvents/gases (most commonly carbon
dioxide), which are fluid at a temperature and pressure above their critical point. This
generally requires very high pressures while temperature remains relatively low. In
this way, it bypasses some of the problems encountered in methods such as
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9
distillation (Shahidi 2009). A more in-depth description of this method, in the
concentration of LC n-3 PUFAs can be found in a review by Mishra et al. (1993).
Currently the major disadvantage of this method is the high capital and running
costs, which limits its use to larger processing companies (Shahidi et al. 1998).
Another relatively new approach for concentrating LC n-3 PUFAs utilises lipases.
Lipases are lipid hydrolysing enzymes which specifically cleave different FAs at
different rates. This approach offers significant advantages over conventional
concentration methods and will be discussed in more detail in Section 1.3.4. For this
method to be improved, new lipases are needed with activities suitable for the
concentration of LC n-3 PUFAs. Microalgae represent a source from which lipases
with these types of activities may be discovered.
1.2 The potential of algae
The term ‘algae’ has no formal taxonomic meaning and may refer to organisms
belonging to four different kingdoms; Bacteria, Plantae, Chromista or Protozoa
(Guiry 2012). Thus the modern taxonomic definition of algae can be confusing, as
documents may discuss them as eukaryotes only or both eukaryotes and prokaryotes.
A common example of this arises when cyanobacteria are placed interchangeably
between being bacteria or algae. For the purpose of this document the assumption
will be made that all algae referred to are eukaryotes. Algae represent a large and
very diverse group of organisms that are typically photosynthetic autotrophs. They
are similar to higher plants in many aspects, however they are considered ‘simple’ as
they lack many of the distinct cellular structures and properties synonymous with
plants (Barsanti & Gualtieri 2014). The diversity of species is vast with estimates of
the total number of species varying greatly (from 30,000 to over 1,000,000). A recent
estimate by Guiry (2012) puts the figure at a conservative 72,500 species, but notes
that deducing an exact number is difficult as the definition of what an alga is, is not
clear cut. Algal species are present in both macroscopic and microscopic forms,
which are denoted macro- or micro-algae. Macroalgae are more commonly and
loosely referred to as seaweed, involving the arrangement of groups of organisms
into large, sometimes semi-complex structures (relative to higher plants). The term
‘microalgae’ refers to algae that exist as single celled organisms or in small
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undifferentiated groups/chains. Microalgae can also exist as endosymbionts,
examples of which include lichens, mosses, corals and sponges (Andersen 1992). As
the primary focus of this research is microalgae all subsequent mention of algae will
be in reference to microalgae unless otherwise stated.
The classification of microalgae species is based primarily on their pigment
components and there are nine divisions. The majority of species are divided into
three main phyla, namely Chlorophyceae (green algae), Rhodophyceae (red algae)
and Heterokontophyceae; which includes Phaeophyceae (brown algae) and
Bacillariophyceae (diatoms) (Guiry 2012; Harwood & Guschina 2009).
Heterokontophyta are predominantly marine based and include a huge diversity of
organisms. Diatoms are by far the most common of the heterokonts and perhaps all
algae, with some estimates suggesting >200,000 species may exist (Mann & Droop
1996), while more recent estimates put the number at ~20,000 (Guiry 2012).
Rhodophyta are also found primarily in marine environments of which macroscopic
forms are common. The opposite is true for Chlorophyta which are primarily
microscopic in form (Lewis & McCourt 2004). This phylum includes a huge
diversity of organisms that exist in the most variable range of environments of all the
phyla. They are widely accepted to be the ancestors from which higher plants
emerged (Lewis et al. 2004; Palmer et al. 2004). The topic of phycology is largely
outside the scope of this project and as such an in-depth discussion of algal
taxonomy is not presented. However, the brief introduction above demonstrates the
vast number of species that exist and the potential for their exploitation in
biotechnological applications.
1.2.1 Microalgal applications and biotechnology
Of the approximately 44,000 described algal species (Guiry 2012), only a relatively
small number have been investigated beyond taxonomic identification. Only a
fraction of those investigated beyond taxonomy have been investigated in depth for
biotechnological and industrial purposes. Furthermore, there are vast numbers of
undiscovered organisms. The sheer variety of species means that organisms may
contain different biological compounds and provide a potentially significant resource
for various applications. These include the isolation of novel compounds, such as
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11
polysaccharides, lipids, peptides, carotenoids and toxins (Cardozo et al. 2007).
According to Cardozo et al. (2007), over 15,000 novel compounds have been
described from algae, however, commercially produced high-value compounds
currently on the market are largely restricted to carotenoids and algal oils (Olaizola
2003). Prominent examples include astaxanthin from Haematococcus, DHA from
Crypthecodinium and β-carotene from Dunaliella (Olaizola 2003). The low number
of algal-derived products available reflects the significant cost of microalgal culture.
Other applications include the use of microalgal species with high oil content, as they
are convenient producers of hydrocarbons and FAs that can be converted into
biodiesel. The need for renewable sources of biodiesel is being driven by the
increasing cost of petroleum and ongoing concerns surrounding the continued
burning of fossil fuels and its impact on climate change (Gavrilescu & Chisti 2005).
Another area of microalgal lipid research focuses on the ability of numerous species
to produce LC n-3 PUFAs. Aquaculture utilises this ability to provide nutrients for
larval and juvenile molluscs, and indirectly for crustacean and fish species (Fidalgo
et al. 1998), of which many examples have been published (Gordon et al. 2006;
Pettersen et al. 2010; Utting & Millican 1997). The lipid composition and quantity of
microalgae is of particular interest (Delaunay et al. 1993; Pettersen et al. 2010; Ragg
et al. 2010), so that nutrient delivery can be maximised. The versatility of microalgae
as a nutrient source is also apparent, as it has been documented that lipid content,
fatty acid and lipid class compositions of microalgae can change during their growth
(Alonso et al. 2000). Microalgal oils containing high EPA and DHA content are also
being utilised for human consumption, as mentioned in Section 1.1.2. To allow for
better utilisation of these organisms in biotechnological applications, a fundamental
understanding of lipid metabolism in microalgae must be obtained.
1.2.2 Microalgae and LC n-3 PUFA metabolism
In nature, the synthesis of EPA and DHA is carried out almost exclusively by marine
microalgae (Ryckebosch et al. 2012). Microalgal lipid metabolism and the
mechanisms that control it are still relatively poorly understood (Khozin-Goldberg &
Cohen 2011). However, recent efforts using sequence data and in silico analyses are
providing a better understanding of the regulation of lipid metabolism in these
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12
organisms. Much of the understanding derived from this work is based on the use of
previously defined homologous genes/proteins in higher plants. These approaches
have been advanced by the increasing availability of complete microalgal genomes
(Khozin-Goldberg et al. 2011). The production of LC n-3 PUFAs in microalgae can
be achieved by a number of pathways. Figure 1.2.2 displays the most common
pathways characterised for EPA and DHA biosynthesis. This synthesis requires
successive desaturation and elongation steps by the appropriate desaturase and
elongase enzymes. However, alternative pathways for DHA synthesis, such as the
polyketide synthase pathway (PKS), have been defined in other marine eukaryotes of
the Thraustochytriaceae family (Metz et al. 2001). Thus, further research is required
to gain a full understanding of LC n-3 PUFA synthesis in microalgae.
Figure 1.2.2 Biosynthesis of LC n-3 PUFAs in eukaryotic microalgae via enzymatic elongation
and desaturation. Desaturases and elongases are denoted by D and E, respectively. Modified from
Harwood et al. (2009)
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Microalgae appear to synthesise LC n-3 PUFAs primarily for the structural and
functional properties they impart in membranes. While these FAs can also be used
for energy production, the processing of LC PUFAs for β-oxidation requires
additional enzymatic steps (e.g. saturases to remove double bonds) (Adarme-Vega et
al. 2012). The incorporation of n-3 PUFAs into various membranes at both the
cellular and organellular levels is well documented. For example, Sato et al. (2003)
showed that limiting carbon dioxide caused changes in chloroplastic membrane
compositions. Numerous studies have demonstrated that the percentage of LC n-3
PUFA in the phospholipids of algae increase as temperature decreases. This indicates
that the increased fluidity provided by the PUFAs in the membranes allows algae to
adapt and survive fluctuations in temperature (Araujo & Garcia 2005; Jiang & Gao
2004; Tatsuzawa & Takizawa 1995; Zhang et al. 2011). Microalgal lipid composition
can also be changed by altering growth conditions such as light, carbon/nitrogen
source and salinity (Fidalgo et al. 1998; Guedes et al. 2010). Under certain
conditions (usually growth limiting), various microalgal species have been shown to
accumulate large amounts of LC n-3 PUFAs (Dunstan et al. 1993).
The shuttling of FAs, particularly PUFAs, between lipid pools in microalgae
suggests that they possess numerous metabolic mechanisms for controlling their lipid
biochemistry. The regulation and control of these processes requires a range of
enzymes. While in silico approaches have identified the presence of numerous
enzymes in microalgae, relatively few have been characterised biochemically. In
species containing high concentrations of LC n-3 PUFAs it would be expected that a
subset of enzymes exist that are capable of modifying these compounds. Enzymes
involved in the modification/catabolism of TAG in microalgae are of potential
utility. Lipases represent one example of enzymes with a great deal of commercial
interest, owing to their versatile catalytic properties (Jaeger & Reetz 1998). Lipases
with specific activities for LC n-3 PUFA hydrolysis, or that select for FAs other than
PUFAs are industrially important, as they can enable concentration of omega-3 FA
for nutritional and pharmaceutical applications. To date, the isolation and
characterisation of a lipase from microalgae has not been described.
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1.3 Lipases
Lipases have been studied for over 150 years since it was first noted that mammalian
pancreatic juices could degrade fats such as butter (Bernard 1856). Lipases are water
soluble proteins that catalyse the hydrolysis of ester bonds in water-insoluble
substrates, such as triacylglycerides (TAG), phospholipids and cholesteryl esters
(Wong & Schotz 2002). In addition to hydrolysis they can exhibit an array of
activities under different conditions. These include esterification, interesterification,
acidolysis, alcoholysis and aminolysis (Joseph et al. 2008). Therefore, lipases are
used extensively as biocatalysts in many commercial and industrial applications,
including pharmaceuticals, cosmetics, detergents, oil and biodiesel processing
(Sharma et al. 2001). They offer a ‘green’ way to perform various chemical
processes and carryout reactions under relatively mild conditions with high
selectivity (Gotor-Fernandez et al. 2006). General research areas include the
structural characterisation, elucidation of catalytic mechanisms and the isolation of
lipases via both molecular biological and native approaches.
1.3.1 Lipase structure and function
Lipases belong to a large family of enzymes known as /β-hydrolases, of which all
members possess a /β-hydrolase fold. Other enzymes also placed in this family
include esterases, proteases, peroxidases, dehalogenases and epoxide hydrolases
(Nardini & Dijkstra 1999). Often referred to as ‘true lipases’, triacylglycerol lipases
(EC 3.1.1.3) are distinct from other lipolytic enzymes (for example phospholipases,
sterol esterases, carboxyl esterases) due to their specificity for TAG. In the past
enzyme classification has been largely based on the activity of enzymes for
substrates of differing solubilities (Anthonsen et al. 1995). From this definition
lipases are esterases that act primarily on insoluble substrates, while esterases react
with water soluble substrates. In this sense lipases are considered a special subset of
esterases (Verger 1997). More recently the classification of lipolytic enzymes has
increasingly been distinguished by analysis of amino acid and nucleotide sequence
data (Wong et al. 2002). This has led to an explosion in the detection and
classification of lipases from sequence-based approaches. However, limitations in the
predictive capabilities of chemical theories means that sequence data alone cannot be
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15
used to accurately determine enzyme activity/specificity (Reymond & Wahler 2002).
Thus, the use of both sequence data and empirical data in combination is the most
comprehensive approach.
Since the first lipase structure was solved by Brady et al. (1990) and the presence of
a catalytic triad was defined, research investigating structure-function relationships
has advanced significantly. The core structure of lipases is the /β hydrolase fold,
which forms the central scaffold of the tertiary structure (Ollis et al. 1992). The
structure of this fold is characterised by a particular -helix and β-sheet architecture,
with the central primarily parallel β-sheet displaying a left-handed super helical
twist. Most of the β-strands are connected by -helices in the central sheet (Nardini
et al. 1999). This topology provides a stable scaffold for the positioning of the
catalytic triad active site, of which the spatial positioning of side chains is
remarkably well conserved (Holmquist 2000). A diagram of the ‘canonical’ or basic
/β hydrolase fold is shown in Figure 1.3.1.
The catalytic triad in lipases is composed of a catalytic nucleophile residue (serine
(Ser)), an acid residue (Aspartic or Glutamic acid (Asp/Glu)) and a histidine (His)
residue (Nardini et al. 1999). The Ser residue resides in a tight turn between sheet β5
and helix C, in what is often referred to as the ‘nucleophile elbow’ (Ollis et al.
1992). The serine active site is generally identified by a sequence motif denoted as
PS00120 [LIV]-[KG]-[LIVFY]-[LIVMST]-G-[HYWV]-S-[YAG]-G-[GSTAC] (S =
serine nucleophile) on the PROSITE database (Hulo et al. 2004). It is the geometry
of the nucleophile elbow that supports the formation of the ‘oxyanion hole’, which
helps stabilise the tetrahedral intermediate formed during hydrolysis (Nardini et al.
1999). The acidic residue may be present as either an Asp or Glu residue. The His
residue is the most conserved of the triad positions and all currently known enzymes
that utilise this active site structure contain a His residue at this position (Nardini et
al. 1999).
Another structural element unique to lipases is the presence of a flexible domain
referred to as a ‘flap’ or ‘lid’. The lid is usually composed of an -helix flanked by
loop regions and is amphipathic in nature. The loop regions act as hinges and are
flexible, allowing for movement of the lid and conformational change to take place.
In the closed form the lid covers the active site preventing catalysis, while in the
CHAPTER I
16
open form the lid swings away revealing the active site, allowing substrate
interaction (Derewenda et al. 1992). This domain is not common to all true lipases
(Hjorth et al. 1993; Martinez et al. 1992; WithersMartinez et al. 1996), however the
majority possess a lid. Furthermore the lid can vary greatly between enzymes with
respect to size and residue composition. For example, lipase B from Candida
antarctica and Candida rugosa lipase have lids containing 5 and 26 residues,
respectively (Grochulski et al. 1994b; Uppenberg et al. 1994). The lid plays an
important role in defining the hydrolytic activity and selectivity of lipases (Brocca et
al. 2003). It also provides a structural explanation for another property of lipases
known as ‘interfacial activation’. Interfacial activation refers to the enhanced activity
of lipases (but not esterases) observed upon contact with a lipid-water interface,
which is thought to trigger the conformational change of the lid (Nardini et al. 1999).
In the interfacially activated conformation, the hydrophobic residues surrounding the
active site and the underside of the lid interact with the lipid interface and hydrolysis
occurs (Schmid & Verger 1998). Figure 1.3.2 depicts this process. This property can
be exploited for lipase purification and immobilisation purposes, which is discussed
in Section 1.3.3.
Figure 1.3.1 Secondary structure of the ‘canonical’ /β hydrolase fold. -Helices and β-sheets
are represented by cylinders and arrows, respectively. Black dots represent the location of residues in
the catalytic triad. Modified from Nardini et al. (1999).
CHAPTER I
17
The activity of lipases can be selective in several ways. The most basic form is
substrate selectivity, whereby a lipase distinguishes between different glyceride
substrates (e.g. TAG, diacylglycerol (DAG), monoacylglycerol (MAG) and
phospholipid). Fatty acid selectivity is also exhibited by lipases and they will
preferentially hydrolyse or discriminate between certain acyl chains (e.g. short-chain
FAs, PUFAs). Lipases also display regioselectivity (positional specificity). This is
defined by which FA position (sn-1, sn-2 or sn-3) on the glycerol backbone the lipase
preferentially hydrolyses. Most lipases are sn-1,3 specific; however a number of
lipases will show no selectivity and hydrolyse all positions readily. There are very
few lipases that display sn-2 selectivity (Kurtovic et al. 2009). Lastly, lipases are
enantioselective, differentiating substrates based on the chirality of the molecules,
which is important for the production of fine chemicals (Berglund 2001).
Figure 1.3.2 Basic representation of lipase lid conformational change and interaction with
hydrophobic surfaces. Modified from Hanefeld et al. (2009).
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18
Lipase selectivity is also determined by the tertiary structure of the active site. Lipase
active sites can vary greatly in their dimensions. From structural analyses a number
of different topologies have been identified (three examples are shown in Figure
1.3.3). The active site topologies depicted are for lipase B from Candida antarctica
(CalB), Rhizomucor miehei lipase (RmL) and Candida rugosa lipase (CrL). CalB
possesses a funnel-type active site, RmL has a cleft-type active site and CrL has a
tunnel-type active site (Pleiss et al. 1998).
Figure 1.3.3 Active site topology of lipase B from C. antarctica, R. miehei lipase and C. rugosa
lipase. The direction of the cross-section view is indicated by an arrow. Modified from Pleiss et al.
(1998).
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19
The physical dimensions of the different active sites determine the rate of reaction on
FA substrates differing in chain-length and degree of unsaturation (Gutierrez-Ayesta
et al. 2007). FA selectivity is the most important consideration to address when
lipases are used for concentrating LC n-3 PUFAs. Most lipases catalyse the
hydrolysis of LC n-3 PUFAs relatively poorly due to both the long acyl chain-length
and the high level of unsaturation. Further hypotheses have been formed regarding
the poor hydrolysis of LC n-3 PUFAs. Firstly, that the unsaturation twists the acyl
chain in a manner that brings the methyl terminus in close proximity to the ester
bond. This results in steric hindrance of the lipase’s access to the ester bond.
Secondly, PUFAs with a double bond in an even numbered position (e.g. cis-4, cis-6
or cis-8) closest to the carboxyl end are discriminated against more severely (Bottino
et al. 1967; Mukherjee et al. 1993). These properties can be useful in the
concentration of EPA and DHA, as specificity towards other FA means they are
effectively concentrated in the glyceride fraction.
1.3.2 Lipase sources
Lipases are ubiquitous throughout the natural world, where they are utilised by
organisms for the mobilisation of FAs from TAG. They have evolved into a diverse
group of enzymes with an array of activities. Lipases have been isolated from and
studied in numerous organisms. Early lipase research mostly focussed on lipases
from mammals. Most lipases were isolated from the digestive tract, such as the
pancreatic lipases, and aided in understanding lipid digestion in humans (Garner &
Smith 1972; Gidez 1968; Vandenbo et al. 1973; Winkler et al. 1990). This research
continues today with particular interest in the role lipases play in lipid metabolism
and their relevance to modern medicine (Zechner et al. 2012; Zimmermann et al.
2009). Other vertebrate sources of lipases include fish (Aryee et al. 2007; Kurtovic et
al. 2010; Nayak et al. 2003). Aquatic organisms offer potentially useful activities,
such as low-temperature activity optima and unique FA selectivity in regard to
PUFAs (Kurtovic et al. 2009). Lipases isolated from vertebrate sources generally
require the presence of bile salts and/or various metal ions to achieve optimal
activity. Plant lipases have also been researched and are mostly isolated from seeds
and fruits of oleaginous plants, such as those belonging to the family Brassicaceae.
Research in this area is important for understanding lipid metabolism in seeds during
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20
germination (Hassanien & Mukherjee 1986). It is common for lipases isolated from
plants to require the presence of divalent cations such as Ca2+ or Mg2+ for optimal
activity. Some of the plant sources lipases have been isolated from are summarised
by Barros et al. (2010).
In recent years the most widely utilised sources for lipase isolation and
characterisation have been microbial. Bacterial (Kennedy & Lennarz 1979; Koritala
et al. 1987; Mitsuda & Nabeshima 1991; Mitsuda et al. 1988; Schuepp et al. 1997)
and fungal (Nagaoka & Yamada 1973; Namboodiri & Chattopadhyaya 2000; Omar
et al. 1987; Sugihara et al. 1988; Tombs & Blake 1982; Tomizuka et al. 1966;
Veeraragavan et al. 1990) lipases have been the subject of intense research for a
number of reasons; (1) the convenience of their growth and production (some
produce lipase extracellularly), (2) availability of established genetic modification
systems and (3) the variety of biochemical properties and specificities these lipases
possess (Hasan et al. 2006). Fungal lipases represent the group of lipases most
widely utilised in industry at this time (Singh & Mukhopadhyay 2012). More
recently there has been interest in isolating lipases from marine microbes such as
algae and thraustochytrids (Kanchana et al. 2011), for their potentially unique and
useful activities. Research in this area has largely been driven by the search for
lipases with LC n-3 PUFA-related activities. However, thus far no lipases from
microalgal sources have been isolated and their activity towards LC n-3 PUFA
substrates characterised. Currently, the example that may be closest to microalgae is
a lipase isolated from the cyanobacteria Spirulina platensis (Demir & Tukel 2010).
More recently a group in France has cloned and expressed lipase-like (Godet et al.
2010) and thioesterase/carboxylesterase (Kerviel et al. 2014) enzymes from the
microalga Isochrysis galbana. Work has also been performed on recombinantly-
expressed lipid hydrolases (galactoglycerolipid lipase, DAG lipase) from
Chlamydomonas reinhardtii to further the understanding of lipid metabolism in this
organism (Li et al. 2012a; Li et al. 2012b). None of these studies have described the
analysis of LC n-3 PUFA specificity of the enzymes, but their work demonstrates the
emerging interest in microalgae as a source of lipases and other lipid hydrolases.
CHAPTER I
21
1.3.3 Lipase isolation and purification
There is no set purification scheme for the isolation of lipases and as such numerous
methods have been developed to purify these enzymes from a range of organisms.
The level of purification required depends on the application. For example, most
commercial preparations do not require purification to homogeneity, while it is
necessary for laboratory-scale preparations for structural analyses (Nagarajan 2012).
Extensive reviews have been presented by Taipa et al. (1992), Saxena et al. (2003)
and Nagarajan (2012) on this area of lipase research, thus a short summary of
purification techniques is given here. The most commonly used methods are non-
specific in nature, relying on general protein properties such as charge, solubility and
molecular weight. These methods include various salt or solvent precipitations, ion-
exchange chromatography and gel filtration chromatography. Saxena et al. (2003)
estimate that approximately 80% of all microbial lipase purification schemes have
used precipitation, with ammonium sulfate precipitation most common.
Precipitations are favoured as they can be used with large quantities of material and
deliver relatively high protein yields (Saxena et al. 2003). Chromatographic step(s)
are usually used after the precipitation step. Ion-exchange chromatography is the
most common chromatography performed for lipase isolation, with the anion-
exchanger diethylaminoethyl (DEAE) the most broadly utilised (Saxena et al. 2003).
Gel filtration chromatography is also widely used, with more than half of all
purification schemes employing this technique. Gel filtration is usually used as a
final step in purification of already partially purified samples (relatively clean) and
provides the additional advantages of desalting and/or buffer-exchanging samples.
Non-specific approaches can be troublesome and generally result in relatively low
yields (Gupta et al. 2004; Saxena et al. 2003).
More specific methods, such as hydrophobic interaction and affinity chromatography
have also been used. Hydrophobic interaction chromatography has been shown to be
highly effective, to the point where one-step purifications have been possible (Gupta
et al. 2005). This method can be lipase-specific and exploits the interfacial activation
these enzymes naturally undergo in the presence of hydrophobic surfaces. It can also
be used in lipase immobilisation (Bastida et al. 1998; Fernandez-Lafuente et al.
1998). Various affinity chromatography protocols have been developed which allow
for increased specificity and therefore faster purifications. Immunopurification is one
such example, where highly specific antibodies are raised against lipases, providing
CHAPTER I
22
an extremely efficient purification. However, despite the efficiency of this method,
the relatively high cost of antibody production means its use is limited (Gupta et al.
2004). Perhaps the newest novel example of affinity chromatography is the use of
lipase-lipase interactions, where covalently immobilised lipases are used to
specifically isolate lipases from crude preparations (Volpato et al. 2010).
Other less widely used methods include aqueous two-phase and reversed micellar
systems. These methods provide alternatives for purification when more
conventional approaches are not working. They are not as selective as the methods
mentioned above and are not stand-alone methods, requiring subsequent processing
to achieve homogeneity (Nagarajan 2012). But the scalability of the methods means
that they may be useful for larger commercial preparations (Gupta et al. 2004).
Regardless of the purification scheme used, the need for the isolation of novel lipases
and their application in biotechnology is apparent.
1.3.4 Lipase-based omega-3 concentration
Lipases provide an attractive platform for the concentration of LC n-3 PUFAs such
as EPA and DHA. They offer a number of advantages over the conventional methods
outlined in Section 1.1.3. The most notable is their ability to carry out hydrolysis
under relatively mild conditions with respect to temperature and pH (Wanasundara &
Shahidi 1998). As EPA and DHA are relatively unstable, the use of mild reaction
conditions reduces the chance of undesirable chemical modifications such as
oxidation, polymerisation or isomerisation of the FAs. The use of enzyme catalysts
also removes the need for organic solvents and chemicals, therefore lessening the
amount of chemical effluent produced. Finally, due to the positional and FA
selectivity of lipases the method can potentially offer a more specific and targeted
approach (Wanasundara et al. 1998). As DHA is found primarily in the sn-2 position
in fish oil, lipases with sn-1,3 selectivity and FA selectivity against LC n-3 PUFAs
are desirable.
In practice lipases are used to concentrate LC n-3 PUFAs by hydrolysing non-LC n-3
PUFA species off the glycerol backbone of TAG. The FFAs are then removed by
physical methods and the LC n-3 PUFAs are left in the acylglycerol fraction, thus the
omega-3 content is enriched (Xu et al. 2007). Owing to the different specificities of
CHAPTER I
23
various lipases this approach can be tailored to different oil substrates, based on their
FA composition and positional distribution. For example, DHA is found primarily in
the sn-2 position in fish oil (while it is predominantly at sn-1/3 in marine mammals),
therefore a lipase with a preference for sn-1/3 positions and FA selectivity for non-
PUFAs would be desirable (Xu et al. 2007).
Currently LC n-3 PUFA concentration by lipases does have some disadvantages.
Firstly, the lipase catalysed process is relatively slow compared to other methods.
Secondly, the cost of producing lipases is quite high. Further research is required to
address these issues. This includes optimising lipase production and catalysis
efficiency as well as the development of improved immobilisation technologies. The
later allows the removal of lipase from the hydrolysate and the recycling of lipases
which is beneficial in commercial settings (Christensen et al. 2003).
Numerous examples of LC n-3 PUFA concentration by lipase have been published.
Work carried out in the early 1990’s demonstrated that the concentration of DHA in
glyceride hydrolysis products increased by two to three times after reaction with
lipase from Candida cylindracea (Hoshino et al. 1990; Tanaka et al. 1992). Since
then a significant amount of research has reported on the concentration of EPA,
DHA, or both, using a variety of lipases (Ko et al. 2006; Kralovec et al. 2010;
Wanasundara et al. 1998). The vast majority of work has focussed on the
concentration of EPA and DHA in the glyceride fraction (i.e. selective hydrolysis of
FAs other than PUFAs). However, it is possible that lipases could be used for the
targeted hydrolysis of these PUFAs from the glyceride, which can be used
subsequently to produce high-concentration reconstituted TAGs. A lipase that
preferentially hydrolyses LC n-3 PUFAs from TAG has not yet been characterised
in-depth. However, some studies of fish digestive juices suggest that lipases with
these properties exist (Halldorsson & Haraldsson 2004a; Lie & Lambertsen 1985).
Halldorsson et al. (2004a) found that LC n-3 PUFAs were preferentially hydrolysed
from astaxanthin diesters by lipolytic enzymes from salmon and rainbow trout
intestine. The same group observed the same specificity with herring oil
(Halldorsson et al. 2004b). These works demonstrate the potential of lipase-based
approaches. Further research is needed for this process to become commercially
viable, requiring the development of already known lipases and/or isolation of new
lipases with more specific activities.
CHAPTER I
24
1.4 Methods for lipase detection and characterisation
As previously discussed, lipases exhibit an array of activities such as esterification,
interesterification, acidolysis, alcoholysis and aminolysis in addition to the standard
hydrolysis (Joseph et al. 2008). An important area of lipase research is the
characterisation of lipase activity. After all, it is the various activities and
specificities that make these enzymes such versatile biocatalysts. Assays that are
sufficiently sensitive and comprehensive yet reliable and convenient are of high
priority for lipase detection and characterisation. Numerous methods have been
developed to fulfil these requirements, with method choice depending on the user’s
requirements for assessing lipase activity. The sophistication of assays can vary
greatly and usually increases as the differential objectives of, screening for activity
(qualitative and semi-quantitative), quantifying activity and characterizing kinetic
patterns of selectivity are attained (Pinsirodom & Parkin 2001). The rate of lipase-
catalysed reactions can be measured a number of ways. The most common methods
measure the rate of substrate disappearance (TAG), the rate of product production
(FA), release of a bound chromophore or fluorophore, rate of emulsion clarification
or changes to viscosity (Beisson et al. 2000). Such methods can be utilised to analyse
lipase activity on substrates in either a quantitative (i.e. rate of hydrolysis) or
qualitative means (FA and regio-selectivity), or both. Methods can be used as stand-
alone techniques or in conjunction with other analytical methods that can increase
the sensitivity and comprehensiveness of sample analysis. An extensive review of
various methods has been presented by (Beisson et al. 2000) and some of the more
common methods are outlined below.
1.4.1 Thin layer chromatography/Iatroscan
Thin layer chromatography (TLC) may be used to separate hydrolysate species into
the various lipid classes. In recent years this system has been modified and
automated with the inclusion of a flame ionisation detector (FID) such as the
Iatroscan TLC-FID models (NTS International, USA). Using this equipment and
methodology TAG, DAG, MAG, free fatty acids (FFA) and various other lipid
compounds can be separated and the relative peak area ratios analysed. This method
can be used to gain quantitative information on hydrolysis rates and semi-qualitative
CHAPTER I
25
data regarding regiospecicificty. A major advantage of this method is the relatively
short time it takes to perform and while superseded in many aspects by newer
methods, it still has its place in many laboratories. A recent review on TLC by Fuchs
et al. (2011) gives a more extensive review of the method.
1.4.2 Chromatography
Reverse-phase high performance liquid chromatography (HPLC) can be used to
analyse lipase hydrolysates. It is a relatively rapid method that requires little sample
preparation, as the FAs do not need to be modified before loading onto the column
(Ergan & Andre 1989). However, the determination of lipid species is restricted by
available sensors, which are limited to refractive index and UV detectors. Gas
chromatography (GC) has largely superseded HPLC as the preferred method of FA
detection and analysis as it provides more sensitive detection, employing a flame
ionisation detector (FID) (Thomson et al. 1999). It is commonly used to separate,
identify and quantify hydrolytic products released by lipase digestion. Hydrolysed
FAs are first modified by methylation yielding fatty acid methyl esters (FAMEs).
This modification increases the volatility of the FA, enabling better migration of the
compounds through the column. Various physicochemical properties of FA species
cause differences in elution time, thereby effecting separation. Eluted compounds are
then detected by FID. The sensitivity of this method can be further improved by the
inclusion of a mass spectrometer (GC-MS), allowing for increased resolution. GC-
MS is considered the gold standard in lipid analyses, providing a comprehensive
means for analysing mixtures of FA both quantitatively and qualitatively. Periodic
sampling of the hydrolysate and analysis of FA release allows hydrolysis to be
followed. While GC (or GC-MS) is a very comprehensive method it does have
drawbacks, namely the time required to perform experiments. Sample hydrolysis, the
requirement of FA methylation (Thomson et al. 1999), combined with column run
time means that the method takes significantly longer than others. The
instrumentation required for this analysis is also expensive.
CHAPTER I
26
1.4.3 Titrimetric assays
The measurement of the hydrolysis rate of a lipase against a specific substrate for
kinetic purposes is more commonly performed by measurement of FA release itself.
Titrimetric methods can be used with glyceride substrates. Measurement of
hydrolysis is usually performed by monitoring pH, whereby the resultant hydrolysed
carboxylic acid is assayed by titration with a suitable base, usually sodium
hydroxide. The pH-stat method is usually employed as a simple means for
monitoring the rate of FA hydrolysis from glyceride substrates. This takes place in an
enclosed reaction vessel where the emulsion (containing natural or synthetic TAG) is
mechanically stirred. Rate is determined by the amount of base added to maintain a
stable end point pH (i.e. the higher the lipolytic activity the more base is added).
From this, kinetic data can be obtained for the characterisation of lipase activity.
Tributyrin is commonly used as a substrate for these assays. However, other natural
substrates such as ghee, olive oil and fish oil may also be used (Kurtovic et al. 2010;
Pinsirodom et al. 2001). Comprehensive titrimetric assay methods use commercial
systems, such as those offered by Metrohm Ltd. (Herisau, Switzerland). The method
can be relatively time intensive meaning it is not ideal for the large-scale screening
of lipase activity (Hou & Johnston 1992).
1.4.4 Colorimetric and fluorometric assays
Lipase activity can be investigated using synthetic substrates. This approach involves
a FA (or other lipid substrate) being attached to a suitable chromophore/fluorophore
through the carboxylic terminus via an ester bond. The release of the
coloured/fluorescent component (which is relative to hydrolytic activity) enables the
continuous monitoring of reaction kinetics and specific lipase activity for a particular
substrate (Pinsirodom et al. 2001). Other advantages of using these methods include
the simplicity and rapidity of the assays, as well as the ability for them to be applied
in a relatively high-throughput fashion using standard laboratory equipment (i.e. a
spectrophotometer or fluorimeter) (Pinsirodom et al. 2001). Common compounds
used as coloured substrates include esters of 2-naphthol (Ramsey 1957), 2,4-
dinitrophenol (Mosmuller et al. 1992) and p-nitrophenol (pNP) (Winkler &
Stuckmann 1979), whereby the absorbance of the released alcohol is read. However,
pNP is by far the most common of these substrates and will be discussed in further
CHAPTER I
27
detail in Section 1.5. Fluorogenic substrates share many of the advantages associated
with colorimetric methods, but offer an additional advantage of increased sensitivity
(Laborde de Monpezat et al. 1990). Commonly used fluorogenic substrates include
rhodamine B and esters of 4-methumbelliferone. The method using rhodamine B was
first used by Kouker and Jaeger (1987) for the screening of lipases from bacterial
cultures on agar plates. Rhodamine B reacts with FAs hydrolysed from TAG that is
present in the agar plate and can be visualised under UV-light. It provides an easy
means of quantifying lipase activity in different strains and colonies. Whilst not as
accurate for reaction rate kinetics as other methods, it provides a very useful early-
stage screening method. Esters of 4-methylumbelliferone are also used for the
analysis of lipase activity and the assay was first performed by Jacks and Kircher
(1967). The assay was further developed for the rapid measurement of lipase activity
using 4-methumbelliferyl butyrate by Roberts (1985). Protocols using 4-
methumbelliferyl acyl esters are carried out in a similar manner to those of
colorimetric substrates, with hydrolysis of the ester bond releasing 4-
methumbelliferone which is subsequently measured at excitation and emission
settings of 365 nm and 450 nm, respectively (Roberts 1985). This substrate is also
used for the detection of lipolytic protein bands in non-denaturing polyacrylamide
gels, as performed by Diaz et al. (1999).
Synthetic compounds can be used both to detect lipase activity and to measure lipase
affinity for certain FA substrates. The latter is particularly true when a group/library
of these substrates is applied for profiling lipase activity, giving both quantitative and
qualitative data (Reymond et al. 2002). One disadvantage of these methods is that the
substrates are unnatural and therefore subsequent analysis with natural substrates
(e.g. TAG) should be performed (reasons discussed in Section 1.5) (Thomson et al.
1999). However, these methods are still highly useful for screening and profiling
purposes.
In addition to the methods outlined above that utilise colorimetric/fluorescent esters,
several other assay substrates that use these detection methods exist. These assays
use natural or labelled TAGs, DAGs and MAGs for screening lipase activities.
Examples using TAG include the use of the spectrophotometric assay for measuring
lipase activity using long-chain TAGs from Aleurites fordii seeds (or tung oil)
(Pencreac'h et al. 2002), naturally fluorescent TAGs from Parinari glaberrimum
(Beisson et al. 1999) and fluorescent pyrene-labeled triacylglycerols (Negre et al.
CHAPTER I
28
1988). An example of a non-TAG substrate that has been developed is p-
nitrobenzofurazan-labeled monoacylglycerol, which has been used to detect low
levels of activity (Petry et al. 2005).
1.5 p-Nitrophenyl assay
FA derivatives of p-nitrophenol (pNP) represent the most extensively used
chromogenic compounds for the measurement of lipase activity. The method was
established in the1970s (Chapus et al. 1976; Sémériva et al. 1974) and involves the
appearance of yellow colour (pNP) as the ester bond is hydrolysed. This is monitored
by reading the absorbance at 400-410 nm (Winkler et al. 1979). Commercially
available pNP-acyl esters are primarily saturated FA derivatives, ranging in chain-
lengths from acetate (C2:0) to stearate (C18:0). Many other derivatives (mainly
esterase substrates) have been synthesised (Qian et al. 2011). Several studies have
also used the synthesised mono-/di-unsaturated (oleic and linoleic acid derivatives,
respectively) acyl groups (Acharya & Rao 2002; Nourooz-Zadeh et al. 1992). The
most widely used derivatives are those of palmitic and butyric acids, which are often
used as stand-alone methods for detection and measurement of lipase and esterase
activity, respectively (Gupta et al. 2003).
This assay is often used for defining chain-length specificity in lipases, by using a
range of substrates and measuring the relative hydrolysis rates. Chain-length data can
be assembled into enzyme activity profiles or ‘fingerprints’ and provide useful
information, particularly in mutagenesis studies (Yang et al. 2002). The assay is
versatile and has been modified for numerous purposes, such as enzymatic catalysis
in solvent systems (Teng & Xu 2007). A limitation of the assay is that it can only be
used at pH >6.0, as pNP absorbance at 410 nm is not present in acidic conditions
(Biggs 1954). In solution, pNP dissociates into the anion p-nitrophenolate or p-
nitrophenoxide and a proton. At acidic pH the colourless p-nitrophenoxide
(maximum absorbance at 317 nm) is favoured and measurement at 410 nm is
ineffective (Hriscu et al. 2013). Hriscu et al. (2013) recently suggested that this
limitation may be addressed by utilising an alternate isosbestic point of p-
nitrophenolate. However, currently most protocols quantify hydrolysis at 410 nm and
accordingly the extinction coefficient of pNP at different pH must be defined.
CHAPTER I
29
Caution should also be taken when attempting to make direct comparisons between
observed pNP activity and lipase activity against natural substrates such as TAG. It
has been suggested that the lipid structure (e.g. TAG or mono-FA substrates such as
methyl esters) influences the enzyme’s affinity for a particular FA (Lyberg &
Adlercreutz 2008). Furthermore, studies have shown that the presence of various FAs
in TAG, such as DHA, may provide inherent protection from hydrolysis by certain
lipases, not only for DHA itself but all FAs in the molecule, referred to as
‘triglyceride specificity’ (Tanaka et al. 1993). These influences cannot be addressed
using pNP-acyl esters or other mono-FA substrates. Consequently, other analyses
using TAG compatible methods, such as those mentioned in previous sections, are
required for more in depth characterisation.
There are also other restrictions to the assay that must be addressed with respect to
non-specific hydrolysis of the esters, particularly when working with short-chain
substrates. For example serum albumin is capable of hydrolysing pNP esters (Sakurai
et al. 2004). This pseudo-esterase activity of albumin has been shown to be the result
of irreversible acetylation of 82 residues and is not the result of catalytic turnover
(Lockridge et al. 2008). Pseudo activity against short-chain pNP esters has also been
demonstrated in pancreatic lipase (Decaro et al. 1986). Decaro et al. (1986)
demonstrated that the isolated C-terminal domain obtained by proteolytic cleavage,
had the same apparent activity on pNP esters as the entire enzyme, while it was
shown later that the enzyme catalytic domain was located at the N-terminus (Winkler
et al. 1990). Again, the pseudo activity of pancreatic lipase on pNP esters was due to
the acylation of a lysine residue (373) (Decaro et al. 1988). For these reasons short-
chain pNP esters must be used with caution. Regardless of the above mentioned
limitations, the relative simplicity and versatility of the pNP assay means that it is
still frequently used, as demonstrated by recent publications such as that by Qian et
al. (2011).
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30
1.6 Research objectives
This research aimed to investigate lipases from microalgal sources for their
suitability in the concentration of LC n-3 PUFAs. The specific aims were:
1. To investigate the growth, lipid class and FA composition of Chaetoceros
calcitrans, Isochrysis sp. and Pavlova lutheri. Then use this data to determine
when lipase activity was present and if it was LC n-3 PUFA-specific.
2. To demonstrate the presence of lipases in various microalgal species and isolate
the enzymes from the native sources.
3. To characterise the molecular weight, activity optima and FA selectivity of the
microalgal lipases through enzymatic analyses.
4. To develop high-throughput sensitive methods for the analysis of lipase FA
selectivity towards LC n-3 PUFAs. (1) To establish a method for synthesising
olefinic pNP-acyl esters in satisfactory (>80%) yields. (2) To incorporate the
novel substrates into the assay platform for examination of lipase FA selectivity.
CHAPTER II
31
CHAPTER II
Materials and methods
2.0 Materials
Product information is listed according to manufacture source at time of usage. All
chemicals were of analytical grade unless otherwise specified.
2.0.1 Chemicals
1-Ethyl-3-(3-dimethylaminopropyl) carbodiimide (EDCI), 4-dimethylaminopyridine
(DMAP), para-nitrophenol (pNP), ammonium chloride, benzamidine hydrochloride
hydrate, β-alanine, bromophenol blue, n-butanol, butyl-Sepharose 4 Fast Flow,
Toyopearl butyl-650C (Supelco®), butylated hydroxytolulene (BHT), calcium
chloride, calcium sulphate, chloroform-d, copper sulphate (pentahydrate), 1,4-
dioxane, ethylenediaminetetraacetic acid (EDTA), Folin & Ciocalteu’s phenol
reagent, glycerol, glycine, gum arabic, iodine crystals, 2-methoxyethanol, 2-(N-
morpholino) ethanesulfonic acid (MES), methyl green, octyl-Sepharose CL-4B,
potassium acetate, potassium iodide, potassium tartrate, sodium carbonate, sodium
chloride, sodium cholate, sodium hydroxide (NaOH), sodium methoxide,
trifluoroacetic acid (TFA), tris (hydroxymethyl) aminomethane (Tris), Triton X-100
(Sigma-Aldrich, St. Louis, MO, USA)
Acrylamide/bis-acrylamide (30%/0.8%, w/v), ammonium persulfate, Bio-safe
Coomassie stain, dithiothreitol (DTT), Silver stain plus kit, sodium dodecyl sulphate
(SDS), tetramethylethylenediamine (TEMED) (Bio-Rad, Hercules, CA, USA)
Acetic acid, acetone, ethyl acetate, hexane, isopropanol, petroleum spirits (60-80 C)
(Chem-Supply, Adelaide, SA, Australia)
Acetonitrile, silica gel 60 (70-230 mesh ASTM) (Scharlab S.L., Barcelona, Spain)
Chloroform, dichloromethane, diethyl ether, methanol (Merck, Darmstadt, Germany)
Hydrochloric acid (HCl), sulphuric acid (BHD Laboratory supplies, Poole, England)
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32
2.0.2 Proteins and standards
Bovine serum albumin (BSA), broad range molecular weight ladder, Aspergillus
niger lipase (AnL), Candida rugosa lipase (CrL) (Sigma-Aldrich, St. Louis, MO,
USA)
Lipase B from Candida antarctica (CalB), Rhizomucor miehei lipase (RmL),
Thermomyces lanuginosus (formerly Humicola) lipase (TlL) (Novozymes,
Bagsvaerd, Denmark)
Sodium caseinate (Westland Milk Products, Hokitika, New Zealand)
2.0.3 Lipids and substrates
Docosahexaenoic acid (DHA), eicosapentaenoic acid (EPA), linoleic acid, -
linolenic acid, palmitoleic acid, oleic acid (Nu-Chek Inc., Waterville, MN, USA)
1,2-dipalmitoyl-rac-glycerol, L-α-phosphatidylcholine, methyl stearidonate, octanoic
acid, palmitic acid, p-nitrophenyl acetate, p-nitrophenyl butyrate, p-nitrophenyl
decanoate, p-nitrophenyl myristate, p-nitrophenyl octanoate, p-nitrophenyl palmitate,
p-nitrophenyl stearate, stearic acid, 37 Component FAME Mix (Supelco®),
tributyrin, tridecanoic acid, tripalmitin (Sigma-Aldrich, St. Louis, MO, USA)
4-Methylumbelliferyl butyrate, 4-methylumbelliferyl palmitate (Gold-Bio, St. Louis,
MO, USA)
Soya lecithin (Archer Daniels Midland Company, Decatur, IL, USA)
Tuna oil (Nu-Mega Ingredients Pty Ltd, Melbourne, VIC, Australia)
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33
2.0.4 Buffers, reagents, solvent systems and emulsions
Algal protein extraction buffer
- 50 mM Tris HCl pH 7.4
- 1 mM EDTA
- 50 mM NaCl
- 2 mM benzamidine hydrochloride hydrate
Lugol’s reagent
- 10 g potassium iodide
- 5 g iodine crystals
- Made to 100 mL in MQ H2O
- 10 mL glacial acetic acid added subsequently
pNP assay buffer
- 20 mM Tris HCl pH 8.0
- 20 mM CaCl2
- 0.01% gum arabic
Lowry reagent A
- 25 g Sodium carbonate
- Dissolved in 250 mL of 0.5 M NaOH
Lowry reagent B
- 0.2 g copper sulphate (pentahydrate)
- 20 mL MQ H2O
Lowry reagent C
- 0.4 g potassium tartrate
- 20 mL MQ H2O
SDS-PAGE running buffer
- 25 mM Tris HCl
- 200 mM glycine
- 0.1% SDS (w/v)
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34
SDS-PAGE sample buffer (5X)
- 200 mM Tris HCl pH 6.8
- 10% SDS (w/v)
- 300 mM DTT
- 20% glycerol (v/v)
- 0.05% bromophenol blue (w/v)
Non-denaturing PAGE running buffer (proteins pI ≤ 7)
- 25 mM Tris-HCl 8.8
- 200 mM glycine
Non-denaturing PAGE sample buffer (proteins pI ≤ 7) (5X)
- 312.5 mM Tris HCl pH 6.8
- 50% glycerol (v/v)
- 0.05% bromophenol blue (w/v)
Acidic non-denaturing PAGE running buffer (proteins pI ≥ 7)
- 350 mM β-alanine
- 140 mM acetic acid
- Adjust with NaOH to pH 4.3
Acidic non-denaturing PAGE sample buffer (proteins pI ≥ 7) (5X)
- 250 mM acetate-KOH pH 6.8
- 50% glycerol (v/v)
- 0.02% Methyl green (w/v)
TLC mobile phase for pNP-acyl ester separation
- 85% petroleum spirits 60-80 C
- 15% ethyl acetate
Column chromatography mobile phase for pNP-acyl ester separation
- 50% petroleum spirits 60-80 C
- 50% dichloromethane
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35
Tributyrin emulsion
- 2.4 g sodium caseinate
- 0.2 g soya lecithin
- 2 mL tributyrin
- 400 mL MQ H2O
- 220 mg CaCl2 (5 mM)
Tuna oil emulsion
- 5% tuna oil (w/v)
- 5% gum arabic (w/v)
- 400 mL MQ H2O
- 220 mg CaCl2 (5 mM)
Lecithin emulsion
- 2.4 g sodium caseinate
- 1.2 g soya lecithin
- 400 mL MQ H2O
- 220 mg CaCl2 (5 mM)
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36
2.1 Methods
2.1.1 Culture of microalgae
Three microalgal species, Chaetoceros calcitrans (CS178), Isochrysis sp. (T.ISO)
(CS177) and Pavlova lutheri (CS182) were grown at The Cawthron Institute, with
cultures originally being obtained from the Commonwealth Scientific and Industrial
Research Organisation (CSIRO), Australia. Batch cultures were grown in triplicate,
under the same environmental conditions. The cells were cultured in 20 L plastic
carboys of modified Conway’s media (36 ppt salt water, phosphate (20 g/L) pH 7.5,
vitamins and other micro-nutrients) (Tannock 2006). The C. calcitrans medium
differed from the others with the inclusion of silicate (30 g/L). Each culture was
initiated with an inoculation density of approximately 6% and grown with exposure
to 24 hr light (~44.8 μE.m-2.sec-1), at 19-20°C and constant aeration with 1% carbon
dioxide (~12.5 mL air/min/carboy). Cell growth was monitored daily by taking a 10
mL aliquot from each carboy, from which a 0.5 mL aliquot was subsequently taken.
To that aliquot, 4 μL of Lugol’s reagent (Karayanni et al. 2004) was added and the
cells were counted using a hemocytometer on the same or following day. Cell
counting took place until the cultures of each species appeared to be entering the
regression phase.
‘Large’ aliquots of approximately 7-10 L were taken periodically throughout the
growth phases for the purposes of lipid isolation and analyses. C. calcitrans and
Isochrysis sp. were sampled four times during their respective logarithmic phases
and three times during stationary phase. Due to its longer growth period P. lutheri
was sampled five times during its logarithmic phase and twice during stationary
phase. The cells present in the larger aliquots were concentrated by centrifugation at
approximately 2,500 rpm (Laboratory separator LA PX 202, Alfa-Laval, Lund,
Sweden). The resulting cell concentrate was then further centrifuged at 2,500 x g at
4 C for 10 min using a Beckman Avanti® J-25 I centrifuge fitted with a J-lite® JLA-
10.500 fixed angle rotor (Beckman Coulter, Brea, CA, USA). Two centrifugations
were used due to the large volumes being handled. The concentrated cells were used
immediately for bulk oil extraction or stored for no longer than 48 h at -80°C.
Concurrently, at each sampling point a ‘small’ 48 mL aliquot of culture was taken for
the determination of the lipid content per cell, which was carried out on the same
day.
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37
2.1.2 Algal lipid extraction and determination of total lipids
Oil was extracted according to Bligh and Dyer (1959). For oil extraction from the
‘large’ sample, the concentrated cells were resuspended with MQ H2O to a volume
of 48 mL. Each 48 mL sample (large and small) was added to 60 mL of chloroform
and 120 mL methanol in a separation funnel, mixed by shaking and left overnight.
Then 60 mL of both chloroform and MQ H2O were added, further mixed by shaking
and left for phase separation to occur for no less than 4 hours. In some instances, 58
mL of MQ H2O and 2 mL of a saturated NaCl solution were added instead of 60 mL
MQ H2O. This was done to aid in the separation of phases and prevent the formation
of an emulsion (Barthet et al. 2002). The lower phase containing chloroform and
lipid was decanted into pre-weighed round bottom flasks and the chloroform
subsequently removed by evaporation under reduced pressure. The total lipid values
calculated from the ‘small’ sample were then combined with cell count data to
calculate the amount of lipid in pg/cell. Subsequently, the oil was re-dissolved in
chloroform, transferred to a gas chromatography vial, sealed in an inert nitrogen
atmosphere and stored at -80ºC.
2.1.3 Separation and determination of lipid classes
The ‘large’ samples that underwent lipid extraction (as in Section 2.1.2) from the
three algal species were analysed and the lipid classes and fatty acid (FA) profiles
defined. The extracted lipid was separated into the lipid classes of neutral lipid (NL),
glycolipid (GL) and polar lipid (PL). The separation was performed using solid phase
extraction (SPE) silica columns (GracePureTM Silica 1000 mg/6 mL), with an elution
protocol according to Stern and Tietz (1993), with some modifications. The columns
were first conditioned with 17.5 mL of chloroform before they were loaded with
approximately 20 mg of lipid sample. Subsequently the first fraction containing NL
was eluted with 35 mL of chloroform, the second fraction containing GL with 30 mL
of chloroform/methanol (17:3, v/v) and the third fraction containing PL with 17.5
mL of methanol. The weight of this fraction was established in a pre-weighed glass
vessel after solvent evaporation. Lipid class fractions were subsequently methylated
for FA analyses by gas chromatography (GC).
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38
For determination of changes in lipid class, two aliquots of each oil sample diluted in
chloroform were spotted onto three pre-burned silica gel SIII Chromarods with a
Drummond microdispenser. After conditioning in a CaCl2 containing desiccator for 5
min, lipid classes were separated in hexane/diethyl ether/acetic acid (60:17:0.1,
v/v/v) for 25 min (Volkman et al. 1989). Lipid classes were then analysed with an
Iatroscan MK5 chromatography-flame ionization detector (FID) analyser (NTS
International, USA) and quantified using Chromstar 4.10 software (SES GmbH,
Bechenheim, Germany). Calculations were calibrated using an external standard
containing TAG (tripalmitin), FFA (palmitic acid), DAG (1,2-dipalmitoyl-rac-
glycerol) and phospholipid (L-α-phosphatidylcholine). The hydrogen pressure of the
Iatroscan was set at 160 mL/min-1 with an air-flow rate of 2 L/min-1.
2.1.4 Lipid methylation and determination by gas chromatography
FA composition was determined by GC analysis using two or three aliquots of each
lipid/lipid class sample, with a known concentration of an internal standard
(tridecanoic acid, C13:0). Samples were first methylated (Hartman & Lago 1997)
before being loaded onto the GC. Methylation was performed by adding 15 mg of
lipid sample to 0.5 mL of 0.5 M sodium methoxide solution and heated at 65°C for 5
min. A further 1.5 mL of methylating agent (ammonium chloride/methanol/sulphuric
acid (2:60:3, w/v/v)) was added and allowed to react for 3 min at 65°C. The FA
methyl esters (FAMEs) produced were extracted with hexane and injected into a
Hewlett Packard Series II GC equipped with a FID (Agilent Technologies, Santa
Clara, CA, USA) and a DB-225 capillary column (15 m x 0.25 mm, film thickness:
0.25 μm, J & W Scientific, Agilent Technologies, Santa Clara, CA, USA). The
carrier gas was helium with a pressure of 68.94 kPa and the make-up gas a
nitrogen/air mixture with pressures of 137.88 kPa and 206.82 kPa, respectively. The
injection volume was 1 μL of sample, while the injection and detector temperatures
were set at 230°C and 250°C respectively. The temperature profile of the GC oven
was initially set at 50°C, held for 1 min after injection and then ramped to 175°C.
This was done at a rate of 25°C/min without a holding time. A final temperature of
225°C was reached by increasing the temperature at increments of 4°C/min. This
final temperature was then held for 5 min. FAs were identified and quantified with an
external 38 FAME standard (0.5 mg/mL Supelco 37 Component FAME Mix, 0.025
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39
mg/mL methyl stearidonate). In addition, the FA composition of some samples were
confirmed by GC-MS (20-10 QP GC-MS, Shimadzu, Kyoto, Japan) run under
similar conditions as described for GC.
2.1.5 Algal cell lysis and protein extraction
Two methods were trialled for the lysis of microalgal cells for the purpose of
obtaining crude protein homogenates; bead-beating and homogenisation. Both
methods were carried out using the same starting material obtained as follows. Algal
culture (10 L of P. lutheri or Isochrysis sp.) from The Cawthron Institute was
centrifuged repeatedly at 2,500 x g for 10 min at 4°C until all cells from the total
volume had been pelleted.
2.1.5.1 Bead-beating
Bead-beating was carried out using a modification of Weis et al. (2002). All samples
were kept on ice between treatments. Algal cells were pelleted by centrifugation at
2,500 x g for 10 min at 4°C and the culture medium discarded. Cells were then
resuspended in 10 mL of filtered seawater (FSW) before being pelleted again by
centrifugation (as above) and the FSW discarded. The pellet of ~1.5 mL was
subsequently resuspended in 3.75 mL of extraction buffer (50 mM Tris-HCl pH 7.4,
1 mM EDTA and 50 mM NaCl). The protease inhibitor benzamidine was also added
fresh to the buffer to a final concentration of 2 mM. This was then transferred to a
glass test tube and 1.5 mL of acid washed glass beads (400-600 μm) added. The
mixture was then agitated 20 times using a vortex in 30 second bursts, with samples
kept on ice in between bursts. The cell homogenate was decanted, excluding the
glass beads, and centrifuged at 25,000 x g for 10 min at 4°C using a Beckman
Avanti® J-25 I centrifuge fitted with a JA-25.50 fixed angle rotor (Beckman Coulter,
Brea, CA, USA). The clarified algal supernatant was then retained and subsequently
analysed using the pNP activity and Lowry concentration assays. Due to the low
activity obtained, this method was later scaled up to allow for more biomass to be
processed. Volumes used were increased but remained at the same ratios. Everything
proceeded in the same manner with the exception of the vessel used for the bead-
beating, where the test tube was replaced with a ‘pear-shaped’ round bottomed flask.
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40
2.1.5.2 Homogenisation by Ultra-Turrax
Due to the nature of this method larger volumes were required than for bead-beating,
although all reagent ratios remained the same. This meant that 10 L of culture was
initially pelleted. Algal cells were treated the same as above with pelleting by
centrifugation at 2,500 x g for 10 min at 4°C and the media discarded. Cells were
then subsequently resuspended in FSW before being pelleted again by centrifugation
and the FSW discarded. Cells were then resuspended in extraction buffer. The
suspended cells were placed in a Schott bottle and homogenised with the Ultra-
Turrax (IKA , Staufen, Germany) for 1 min at 15,000 rpm, followed by 30 seconds
on ice. This was repeated 5 times. The homogenised suspension was subsequently
centrifuged at 25,000 x g for 10 min at 4°C. The clarified algal supernatant was then
retained and subsequently analysed using the pNP activity and Lowry concentration
assays.
2.1.6 Freeze-drying of algal protein homogenates
Freeze-drying was used as a method to both store and transport algal protein
homogenates. Aliquots (~10 mL, weighed to give 10 g) of P. lutheri were prepared
in duplicate with and without 5% glycerol. One of each duplicate was first weighed
and then frozen at -80°C. Following this they were processed using an AdVantage
Plus freeze-drier (VirTis, Gardiner, NY, USA) with a cycle of -25°C for 60 min,
40°C for 120 min and 20°C until samples were assessed as being dry (visual and
texture). The samples were then weighed again to determine the weight of water
removed. Prior to assay, samples were resuspended with the corresponding weight of
the water removed by freeze-drying. The resuspended samples and the appropriate
non-freeze-dried samples (stored at 4 C) were then assayed for activity by pNP
assay.
2.1.7 Lowry protein concentration assay
Measurement of protein concentration for protein homogenates and various fractions
from algal samples were obtained using a method adapted from Lowry et al. (1951).
A standard curve covering the range of 0-500 μg/mL was prepared with bovine
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41
serum albumin (BSA) for every set of assays and used to define concentration in
unknown samples. Reagents used in the assay are the ‘combined protein reagent’
(CPR) and ‘Folin-phenol solution’ (FPS), which were made fresh for each batch of
assays. CPR contained reagent A (0.94 M sodium carbonate in 0.5 M NaOH),
reagent B (40 mM calcium sulphate) and reagent C (88 mM potassium tartrate) at a
ratio of 20:1:1 (v/v/v), while FPS was Folin & Ciocalteu’s phenol reagent in MQ
H2O at a ratio of 1:9 (v/v). Standards and samples of unknown concentration (in
duplicate) were appropriately diluted to a final volume of 1 mL. An aliquot of CPR
(1 mL) was first added to each standard and sample, mixed by vortex and incubated
at room temperature for 15 min. Subsequently, 3 mL of FPS was added to each and
mixed. Samples were then incubated for a further 45 min at room temperature and
the absorbance of each measured at 540 nm using a Cary 300 Bio UV-visible
Spectrophotometer (Agilent Technologies, Santa Clara, CA, USA).
2.1.8 p-Nitrophenyl activity assay
A range of p-nitrophenyl (pNP)-acyl esters were used to determine lipase activity
spectrophotometrically in a modified assay based on that of Winkler et al. (1979). A
stock solution was made by dissolving each pNP-acyl ester in 2-propanol at 30 C to
a concentration of 15 mM. A final concentration of 0.25 mM was reached when the
stock solution was added to reaction buffer (20 mM Tris-HCl pH 8.0, 20 mM CaCl2,
0.01% gum arabic), also pre-warmed to 30 C. An aliquot (20-100 μL) of lipase was
then added to the buffered substrate to a final volume of 1.5 mL and mixed by
inversion before being placed in the spectrophotometer. Lipases were appropriately
diluted with reaction buffer to allow all reactions to take place without reactions
running to completion, ie; A410 was linear. A negative control (reaction buffer only)
was undertaken with each pNP-acyl ester to establish background levels of
spontaneous hydrolysis, which were later subtracted from the actual enzyme
hydrolysis rates. The reaction took place at 30 C and the release of pNP was
measured at 410 nm in a Cary 300 Bio UV-visible Spectrophotometer fitted with a
Cary temperature controller (Agilent Technologies, Santa Clara, CA, USA).
Depending on the data interpretation required absorbance data was dealt with in one
of two ways. When specific reaction rate was being investigated activity was
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42
calculated in units/mL (U/mL). This takes into account the A410/min, extinction
coefficient of pNP (relative to pH) ( M), sample volume added to assay in mL (Vs),
total assay volume in mL (Vt) and the dilution factor of the enzyme used (DF). The
calculation used is presented below.
Alternatively, if investigating relative reaction rates data is expressed as a
percentage. This is possible if the same enzyme (providing enzyme concentration is
uniform in all reactions) was being tested against different substrates or different
temperatures, providing all other variables remain the same (ie: pH). In these
instances the highest data point was converted to 100% and other data points
normalised to this.
2.1.9 Titrimetric activity assay
Titrimetric activity assays were used as a method for measuring lipase activity
against glyceride substrates. Triacylglycerol emulsions were formed by
homogenisation using an Ultra-Turrax (IKA , Staufen, Germany) at 19,000 rpm for
2 min; these emulsions were then mixed using a magnetic stirrer bar until required. A
number of different substrates and emulsion systems were used including tributyrin
(Committee 1981), tuna oil (Desnuelle et al. 1955) and lecithin. The substrate recipes
can be found in Section 2.0.4. The tuna oil emulsion was filtered after
homogenisation to remove soluble material originating from the gum arabic. A 907
Titrando pH-stat titration unit (Metrohm, Herisau, Switzerland) linked to a computer
running Tiamo 2.3 light software (Metrohm, Herisau, Switzerland) was used to carry
out and record each titration. Substrate emulsions were prepared fresh on the day of
use. Substrate was added to the stirred reaction vessel and allowed to warm to 30°C
before the pH was manually adjusted to slightly above pH 8.0. Lipase was then
added to the reaction vessel and the auto-titration begun. Lipases were diluted
appropriately in order to obtain titration rates of approximately 0.2-0.6 mL/min. The
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43
titrant used was 10 mM NaOH. To allow comparison, all activity data is presented as
the volume (mL) of 0.01 M NaOH titrated/min/mL of lipase in the reaction vessel.
This is necessary because titration methods do not allow calculation of mols of FA
released in a mixed FA substrate. All titration data used to calculate activity was
taken from the most linear regions of the titration curves. Blank titrations (no
enzyme) were also performed periodically to enable spontaneous substrate hydrolysis
to be calculated and subtracted from the relevant enzyme assays.
2.1.10 4-Methylumbelliferyl butyrate activity assay
This assay was used to measure lipase activity and is a modification of Roberts
(1985). Liposomal dispersions of 4-methylumbelliferyl butyrate (MUB) were
prepared fresh daily. The MUB (2 mM) was mixed with 0.2% soya lecithin and
solubilised in chloroform/methanol (2:1, v/v). Solvent was evaporated under a
nitrogen stream and then 10 mL of 150 mM NaCl was added. The solution was then
mixed by vortex, followed by sonication for 20 x 1 sec (repeated three times) bursts
at 3 Watts using a 500 Watt Ultrasonic processor fitted with a tapered microtip
(Sonics & Materials Inc., Newtown, CT, USA). The substrate solution was then
filtered through a 0.45 μm filter to remove larger aggregates and stored in darkness
before assays commenced. Each assay contained (in order of addition) 100 μL of
assay buffer, 300-400 μL of substrate solution and 0-100 μL of enzyme to make a
final volume of 500 μL for all assays. The buffer component usually consisted of 100
mM Tris-HCl pH 7.0 and 20 mM CaCl2. In assays investigating pH optima, Tris-HCl
was adjusted to the appropriate pH (pH 7.0, 8.0, 9.0) or replaced by a citrate buffer
(pH 3.0, 4.0, 5.0, 6.0). Once all components were added the assay solution was
mixed and placed in a water bath at 30ºC for 10 minutes. Assays were removed and
the reaction quenched by addition of 2.5 mL of ice cold 1.5 M Tris-HCl pH 7.5, and
samples placed on ice. The fluorescence of each assay was then quantified using a
Cary Eclipse Fluorescence spectrophotometer (Agilent Technologies, Santa Clara,
CA, USA), operating with excitation and emission settings of 365 nm and 450 nm,
respectively. The change in fluorescence was measured by subtracting the blank (0
μL enzyme, 100 μL buffer and 400 μL substrate in assay) from those assays
containing enzyme at each different pH. All buffers were checked to establish
background fluorescence, which was found to be negligible.
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44
2.1.11 1D SDS-PAGE
Denaturing, reducing, discontinuous 1D SDS-PAGE was conducted according to a
modified literature method (Davis 1964; Ornstein 1964). All SDS-PAGE analyses
were performed using a Mini-PROTEAN® Tetra cell (Bio-Rad, Hercules, CA, USA)
gel electrophoresis unit. Gels were made using the casting module, plates and well
combs provided with the electrophoresis unit. Before casting, gel plates were rinsed
in MQ H2O before being dried with Kimwipes, rinsed again with acetone and left to
air dry. The resolving gel (12% cross-linking) (10.2 mL MQ H2O, 7.5 mL 1.5 M
Tris-HCl pH 8.8, 0.15 mL 20% (w/v) SDS, 12 mL acrylamide/bis acrylamide
(30%/0.8%, w/v), 0.15 mL 10% (w/v) ammonium persulfate and 20 μL TEMED)
was poured, then saturated n-butanol layered on top to protect it from the air and to
give a flat gel surface. The gel was then left to polymerise for 30 min. Butanol was
removed and the surface of the gel rinsed with MQ H2O before being dried with
Kimwipes. The stacking gel (4% cross-linking) (3.075 mL MQ H2O, 1.25 mL 0.5 M
Tris-HCl pH 6.8, 0.025 mL 20% (w/v) SDS, 0.67 mL acrylamide/bis acrylamide
(30%/0.8%, w/v), 25 μL 10% (w/v) ammonium persulfate and 5 μL TEMED) was
poured and the comb put in place (taking care not to trap any air bubbles) and left to
polymerise. Gels were placed in the electrophoresis unit and running buffer (refer to
Section 2.0.4) added until the sample wells were submerged. Wells were flushed
with buffer to ensure no air bubbles or gel debris was present. Samples were
prepared by adding the protein solution to 5X sample buffer (refer to Section 2.0.4)
at a ratio of 4:1 (v/v), mixed by vortex and subsequently heated to 75°C for 10 min.
Denatured samples were then pipetted into their respective wells (sample volume
was dependent on the experiment) and 4 μL of a broad range molecular weight
standard (contains markers; 200, 116, 97, 66, 45, 31, 21, 14 and 6.5 kDa) (Sigma-
Aldrich, St. Louis, MO, USA) was loaded on each gel as a mass reference.
Electrophoresis was performed at 120 V for ~1.5 hours or until the dye front was
close to the bottom of the gel. After electrophoresis each gel was rinsed in MQ H2O
for 5 min before being stained. Staining with Coomassie used Bio-safe Coomassie
stain (Bio-Rad, Hercules, CA, USA) according to the manufacturer’s instructions.
Silver staining was also carried out according to the manufacturer’s instructions
using a Silver Stain Plus kit (Bio-Rad, Hercules, CA, USA). Images of gels were
acquired using a Molecular Imager Gel Doc™ XRS+ system (Bio-Rad, Hercules,
CA, USA).
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45
2.1.12 Non-denaturing PAGE (for proteins pI ≤ 7.0)
Non-denaturing (native), discontinuous 1D PAGE was conducted according to
Bollag et al. (1996). General gel pouring and electrophoresis procedures were carried
out as described in Section 2.1.11 with the following alterations. The resolving gel
(8% cross-linking) consisted of 4.8 mL MQ H2O, 2.5 mL of 1.5 M Tris-HCl pH 8.8,
2.7 mL acrylamide/bis acrylamide (30%/0.8%, w/v), 50 μL 10% ammonium
persulfate and 5 μL TEMED. The stacking gel (5% cross-linking) consisted of 2.3
mL MQ H2O, 1 mL 0.5 M Tris-HCl pH 6.8, 0.67 mL acrylamide/bis acrylamide
(30%/0.8%, w/v), 30 μL of 10% ammonium persulfate and 5 μL TEMED. Gels were
placed in the electrophoresis unit and pre-chilled electrophoresis buffer (25 mM Tris,
192 mM glycine, Section 2.0.4) added until the sample wells were submerged.
Samples were prepared by adding the protein solution to 5X sample buffer (321.5
mM Tris-HCl pH 6.8, 50% glycerol, 0.05% bromophenol blue, Section 2.0.4) at a
ratio of 4:1 (v/v) respectively and mixed by vortex. Electrophoresis was performed in
a cold room (4-8 C) at 150 V for ~1.5 hours or until the dye front was close to the
bottom of the gel. After electrophoresis each gel was further processed by either
MUB gel-overlay activity assay (see Section 2.1.14), or staining with Bio-safe
Coomassie stain (Bio-Rad, Hercules, CA, USA) according to the manufacturer’s
instructions.
2.1.13 Acidic non-denaturing PAGE (for proteins pI 7.0)
Acidic non-denaturing (native), discontinuous 1D PAGE was conducted essentially
as done by (Reisfeld et al. 1962). All gel electrophoreses were performed as in
Section 2.1.11 with the following differences. The resolving gel (7% cross-linking)
was made up of 3.3 mL MQ H2O, 3.35 mL of 1.5 M acetate-KOH pH 4.3, 3.2 mL
acrylamide/bis acrylamide (30%/0.8%, w/v), 160 μL of 10% ammonium persulfate
and 20 μL TEMED. The stacking gel was prepared with 2.13 mL MQ H2O, 0.83 mL
0.25 M acetate-KOH pH 6.8, 0.33 mL acrylamide/bis acrylamide (30%/0.8%, w/v),
33 μL of 10% ammonium persulfate and 5 μL TEMED. The electrophoresis buffer
recipe was 0.35 M β-alanine, 0.14 M acetic acid with pH adjusted to 4.3 (Section
2.0.4). Samples were prepared by adding the protein solution to 5X sample buffer
(1.45 mL 50% glycerol, 0.5 mL 0.25 M acetate-KOH pH 6.8 and 0.02% methyl
CHAPTER II
46
green, Section 2.0.4) at a ratio of 4:1 (v/v), respectively. Electrophoresis was
performed with the polarity of the anode and cathode reversed (black to red and vice
versa) at 200 V/30 mAH for ~1.5 hours or until the dye front was close to the bottom
of the gel.
2.1.14 Gel-overlay activity assay
Once electrophoresis had been performed, polyacrylamide gels run under non-
denaturing conditions were subsequently analysed by a gel-overlay activity assay
method modified from that of Diaz et al. (1999). Gels were rinsed in MQ H2O and
then equilibrated in pNP assay buffer (refer to Section 2.0.4) for 10 min, before being
placed inside an imaging system. A 20 mM stock solution of 4-methumbelliferyl
butyrate (MUB) was made in 2-methoxyethanol. This was then diluted in buffer to a
final concentration of 150 μM before being poured on the gel and left for 10 min.
Fluorescent activity bands were observed under UV illumination and images taken
using a Molecular Imager Gel Doc™ XRS+ system (Bio-Rad, Hercules, CA,
USA). Gels were subsequently rinsed with MQ H2O before being stained with Bio-
Safe Coomassie (Bio-Rad, Hercules, CA, USA) according to the manufacturer’s
instructions.
2.1.15 Ammonium sulfate precipitation
Ammonium sulfate precipitation was tested as an initial step for the concentration of
lipases in algal protein homogenates. The freeze-dried protein homogenate (600 mg)
was resuspended in 6 mL of MQ H2O and left to equilibrate for 2 hours at 4 C. The
sample was then centrifuged at 10,000 x g to remove insoluble material. Ammonium
sulfate fractions were prepared at 10, 20, 40 and 60% saturation in a beaker on a
magnetic stirrer at 4 C. Each fractionation step involved slowly adding solid
ammonium sulfate until it was dissolved and then leaving for 1 hour. This was
followed by centrifugation at 10,000 x g at 4 C, with the supernatant retained for
subsequent fractionation. The pellet was resuspended in 2 mL of 20 mM Tris-HCl
pH 8.0. Before further analysis (pNP and Lowry assays) all resuspended pellets were
dialysed using 10 kDa molecular weight cut-off (MWCO) dialysis tubing in 20 mM
CHAPTER II
47
Tris-HCl pH 8.0. The amount of ammonium sulfate required to achieve the
appropriate saturation for each fraction was calculated from Wingfield (2001).
2.1.16 Anion-exchange chromatography
Anion-exchange chromatography was trialled as a method for enriching lipases with
acidic isoelectric points from the crude protein homogenate. A BioLogic DuoFlow
(Bio-Rad, Hercules, CA, USA) FPLC fitted with a 1 mL HiTrapTM DEAE FF (weak
anion) or 1 mL HiTrapTM Q FF column (strong anion) (GE Healthcare, Little
Chalfont, UK) was used for this step. The column was equilibrated in buffer ‘A’ (20
mM Tris-HCl pH 8.0) before chromatography commenced. Aliquots of dialysed
crude protein homogenate were loaded onto the column with buffer ‘A’. Protein that
bound to the column was eluted with a concentration gradient of buffer ‘B’ (20 mM
Tris, 0.5 M NaCl pH 8.0). As the concentration of NaCl increased during the
chromatography the bound protein was displaced off of the column ligand. The
fractions collected that were of interest were analysed by the pNP activity assay.
2.1.17 Cation-exchange chromatography
A cation-exchange chromatography step was used as a method for enriching lipases
with basic isoelectric points from the crude protein homogenate. A BioLogic
DuoFlow (Bio-Rad, Hercules, CA, USA) FPLC fitted with a 1 mL HiTrapTM SP FF
(weak cation) (GE Healthcare, Little Chalfont, UK) was used for this step. The
column was equilibrated in buffer ‘A’ (20 mM MES pH 6.0) before chromatography
commenced. Aliquots of dialysed crude protein homogenate were loaded onto the
column with buffer ‘A’. Protein that bound to the column was eluted with a
concentration gradient of buffer ‘B’ (20 mM MES, 0.5 M NaCl pH 6.0). As the
concentration of NaCl increased during the chromatography the bound protein was
displaced off of the column. The fractions of interest were analysed by the pNP
activity assay.
CHAPTER II
48
2.1.18 Hydrophobic interaction chromatography
Hydrophobic interaction chromatography was used as a method for purifying lipase
from microalgae. The hydrophobic supports used to bind lipase were butyl-
Sepharose , octyl-Speharose (Sigma-Aldrich, St. Louis, MO, USA) and
Toyopearl butyl-650C (Supelco , Sigma-Aldrich, St. Louis, MO, USA). The
Sepharose resins were used to separate lipases sequentially with increasing
hydrophobicity. Freeze-dried protein homogenate was resuspended in MQ H2O to a
concentration of 100 mg/mL. The protein homogenate (10 mL) was dialysed
overnight at 4 C against 5 mM Tris-HCl pH 7.4, in dialysis tubing with a 10 kDa
MWCO. Wet resins (approximately 3 g) were washed three times with MQ H2O and
then equilibrated in 5 mM Tris-HCl pH 7.4. The homogenate was applied to the
butyl-Sepharose (less hydrophobic) and mixed by inversion at 4 C overnight. The
butyl-Sepharose was separated from the protein supernatant by a sintered glass filter
(under vacuum) and retained for analysis. A 1 mL aliquot of supernatant was also
retained, before the remainder was added to the octyl-Sepharose and left to incubate
overnight at 4 C. The supernatant and resin were again separated and retained for
further analysis. The butyl- and octyl-Sepharose resins were both washed with 5 mM
Tris-HCl pH 7.4 to remove any residual supernatant material and then left to air-dry
at room temperature. The dialysed homogenate (starting material) and the butyl-
/octyl-Sepharose supernatants were analysed by pNP and Lowry assays. The pNP
assay was performed with both pNP-butyrate (pNPB) and pNP-myristate (pNPM) to
allow monitoring of both esterase and lipase activity. Activity on the resins was
measured by pNP and titrimetric assays. The same procedure was followed with the
butyl-Toyopearl resin.
2.1.19 p-Nitrophenyl-acyl ester synthesis
p-Nitrophenyl (pNP)-acyl esters were synthesised using a modified Steglich
esterification (Neises & Steglich 1978). p-Nitrophenol (pNP) (334 mg, 2.4 mM, 1.2
equiv.), FA (1.0 equiv.), 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide (EDCI)
(575 mg, 3.0 mM, 1.5 equiv.) and 4-dimethylaminopyridine (DMAP) (12.2 mg, 0.01
mM, 0.05 equiv.) were dissolved in 10 mL of chloroform. The solution was stirred
continuously for 20 hours at room temperature, with the reaction carried out in
CHAPTER II
49
darkness and under nitrogen. Solvent was removed from the reaction mixture by
evaporation under reduced pressure to give a crude product. Subsequently, this was
dissolved in dichloromethane (DCM) and purified using silica gel flash
chromatography (Section 2.1.21). All reactions and chromatography fractions were
monitored in real time by thin layer chromatography (TLC) (Section 2.1.20)
2.1.20 Thin layer chromatography of pNP-acyl esters
Thin layer chromatography (TLC) was performed to monitor pNP-acyl ester
formation in real-time. A piece of filter paper was placed in the TLC tank, a mobile
phase of ethyl acetate and petroleum spirits 60-80 C (3:17, v/v) was then added. The
tank was sealed and left for 5 min to saturate the chamber atmosphere. Samples were
spotted onto silica gel 60 F254 aluminium TLC sheets (Merck, Darmstadt, Germany)
and the sheets then transferred into the TLC tank. The solvent was allowed to
migrate to ~1 cm from the top of the sheet before being removed from the tank and
the solvent front marked. Separation of compounds was then observed under UV
illumination and images taken in a Molecular Imager XRS+ system (Bio-Rad,
Hercules, CA, USA).
2.1.21 Silica gel chromatography of pNP-acyl esters
Silica gel column chromatography was carried out to isolate synthesised pNP-acyl
esters from the reaction mixture. A glass column was packed with silica gel 60 (70-
230 mesh ASTM) (stationary phase) and equilibrated with DCM and petroleum
spirits 60-80 C (1:1, v/v). The reaction mixture (dissolved in DCM) was loaded on to
the column and the mobile phase run continuously with 10 mL fractions collected.
Fractions were analysed by TLC and those containing the compound of interest were
subsequently pooled and the solvent removed by evaporation under reduced pressure.
Samples were further dried under nitrogen before being stored at -20 C. Unsaturated
pNP-acyl esters were obtained as clear oils ranging from colourless to light yellow in
colour.
CHAPTER II
50
2.1.22 Yield calculations of pNP-acyl esters
The amount of FA included in each respective reaction differed due to the varying
molecular weights. Yields were calculated once each of the synthesised pNP-acyl
esters were separated from the crude reaction mixture via column chromatography
and the solvent had been removed. The yield of each reaction product was calculated
as below.
2.1.23 Mass spectrometry of pNP-acyl esters
Mass spectra were gathered using a 6210 MSD TOF (time of flight) mass
spectrometer (Agilent Technologies, Santa Clara, CA, USA) used in positive
electrospray ionisation mode. The analysis conditions were: drying gas (N2) flow rate
and temperature (7 L min-1, 350 C), nebuliser gas (N2) pressure (30 psi), capillary
voltage (3.0 kV), vaporiser temperature (350 C), and cone voltage (60 V). Mass
spectrometry (MS) data acquisition was carried out using MassHunter Workstation
(version B.02.00 (B1128)) (Agilent Technologies, Santa Clara, CA, USA) and data
analysis carried out using MassHunter Qualitative Analysis (version B.03.01)
(Agilent Technologies, Santa Clara, CA, USA).
2.1.24 UV spectroscopy of pNP-acyl esters
A UV absorbance spectrum was obtained for each of the synthesised pNP-acyl esters
to determine their optimal absorbance. This was done by analysis of HPLC peaks
consistent with each of the compounds and utilising the diode array detector (see
Section 2.1.27).
CHAPTER II
51
2.1.25 FTIR of pNP-acyl esters
Fourier transform infrared spectroscopy (FTIR) was conducted using an Alpha FTIR
spectrometer (Bruker Optik GmbH, Ettlingen Germany) equipped with a deuterated
triglycine sulphate detector and a single reflection diamond attenuated total
reflectance (ATR) sampling module (Platinum ATR Quick-Snap™). ATR-FTIR
measurements were performed with 1 μL of the pNP-acyl esters being placed onto
the 2 x 2mm2 sensing surface of the diamond ATR crystal. In the case of p-
nitrophenol (powder) the sample was placed on the surface and pressure applied on
top of the sample using an ergonomic clamp. High quality spectra (16 scans) were
collected from each sample. The ATR-FTIR spectra were acquired at 4 cm-1 spectral
resolution with 64 co-added scans. Blackman-Harris 3-Term apodisation, Power-
Spectrum phase correction, and zero-filling factor of 2 were set as default acquisition
parameters using OPUS 7.0 software suite (Bruker Optik GmbH, Ettlingen
Germany). Background spectra of a clean ATR surface were acquired prior to each
sample measurement using the same acquisition parameters. All spectra were
normalised against the peak at ~1765 cm-1 associated with ester linkages (C=O) (Van
de Voort & Sedman 2000), giving a direct correlation of the C-H stretching band to
FA concentration. Normalised spectra were subsequently converted to the second
derivative using OPUS 7.0 software.
2.1.26 1H and 13C NMR of pNP-acyl esters
NMR was used to aid in compound characterisation, particularly to identify cis –
trans isomerisation. 13C NMR spectra were recorded with a AVANCE III 500MHz
NMR spectrometer (Bruker BioSpin GmbH, Rheinstetten, Germany), at 125.77MHz,
in CDCl3. 1024 scans were accumulated using a 90o pulse of 9.1us, with an
acquisition time of 1.1 seconds and a delay of 2 seconds. Broadband decoupling was
applied during acquisition. 1H spectra were also recorded with a AVANCE III
500MHz NMR spectrometer, at 500.13MHz, in CDCl3. 16 scans were accumulated
using a 90o pulse of 14.0 us, with a recycle time of 4.2 seconds. All chemical shifts
were measured using residual chloroform (7.26 ppm) as a secondary reference to
TMS as zero ppm.
CHAPTER II
52
2.1.27 Reverse-phase high performance liquid chromatography of
pNP-acyl esters
The stability of synthesised pNP-acyl esters was analysed using reverse-phase high
performance liquid chromatography (HPLC). An Agilent 1260 Infinity series HPLC
(Agilent Technologies, Santa Clara, CA, USA), equipped with a solvent degasser,
quaternary pump, auto-sampler, thermostatted column compartment, diode array
detector and fitted with a LiChrospher 100 RP-18 column (124 mm x 4 mm, 5 μm
particle size, Merck, Darmstadt, Germany) was used for the analyses. Storage
stability was investigated by storing samples over six months with and without 0.1%
butylated hydroxytolulene (BHT) (antioxidant) at -20 C under nitrogen. Samples
were removed from cold storage and subjected to HPLC periodically. Analytical
separations were carried out with a gradient flow of 35% acetonitrile + 0.01%
trifluoroacetic acid (TFA) and MQ H2O + 0.01% TFA to 100% acetonitrile + 0.01%
TFA over 15 min, followed by a 10 min hold at 100% acetonitrile + 0.01% TFA. The
column temperature was maintained at 25°C throughout. Samples were prepared in
acetonitrile and 20 μL injected. The signal was monitored at multiple wavelengths;
210 nm, 270 nm and 315 nm to detect FAs, pNP-acyl esters and pNP, respectively.
Each HPLC trace was normalised against subsequent traces (all trace data fitted to
the trace with the highest peak area) to allow for peak area ratios to be analysed
across different HPLC runs.
Assay stability was investigated by performing the pNP assay (Section 2.1.8) without
lipase present. FAs and derivatives were then extracted from the buffered substrate
solution by addition of diethyl ether (1:1, v/v), mixed by inversion and phases
allowed to separate before decanting the solvent layer. Diethyl ether was removed
under nitrogen and the dried sample resuspended in acetonitrile for analysis by
reverse-phase HPLC, as above.
2.1.28 Iatroscan analysis of lecithin and lecithin hydrolysis products
Two lecithin emulsions were produced as described in Section 2.1.9 using the recipe
from Section 2.0.4, with the addition of 100 mM Tris and the pH adjusted to pH 8.0
with HCl. One of the emulsions contained 5 mM CaCl2 the other did not. A 30 mL
aliquot of each emulsion was then reacted individually with each different lipase
CHAPTER II
53
while the pH was monitored. None of the hydrolysates changed by more than 0.1 pH
units for the duration of hydrolysis. The lipid was then extracted using the Bligh et
al. (1959) method. The solvent was removed by evaporation under reduced pressure
and the lipid transferred to a vial for storage under nitrogen before analysis. The lipid
content of each sample was the analysed by Iatroscan as in Section 2.1.3.
2.1.29 Analysis of micelle surface charge
The surface charge of the lecithin micelles (with and without CaCl2) was measured
using a Malvern Nano Zetasizer (Malvern Instruments Ltd., Malvern, UK).
Emulsions were produced as described in Section 2.1.9, using the recipe from
Section 2.0.4, and then diluted to allow for accurate unobscured measurement. The
emulsion without CaCl2 was diluted 1:4 (v/v) with MQ H2O, while the emulsion
containing CaCl2 was diluted in 5 mM CaCl2 to maintain the concentration of
calcium. Emulsions were analysed in duplicate and the zeta potential (surface
charge) obtained.
2.1.30 Analysis of micelle size distribution
Micelle size distribution in emulsions of lecithin with and without CaCl2 was
measured by dynamic light scattering. To perform the analyses a Malvern
Mastersizer (Malvern Instruments Ltd., Malvern, UK) was used. Emulsions were
produced as described in Sections 2.0.4 and 2.1.3. The volume mean diameter
(D[4,3] value) which indicates the presence of larger particles, was calculated using
the equation below.
The d(0.1), d(0.5) and d(0.9) values were also obtained. These values represent the
average micelle size corresponding to the cumulative distribution at 10%, 50% and
90%, respectively (i.e. d(0.1) is the micelle size at which 10% of the total distribution
is smaller than that size).
ni = number of droplets with the same diameter Di = the droplet size
CHAPTER III
54
CHAPTER III
Growth and lipid composition of Chaetoceros calcitrans,
Isochrysis sp. and Pavlova lutheri
3.0 Introduction
Microalgae are a highly diverse group of microorganisms providing a resource that is
largely untapped in terms of novel biologically active compounds. Relatively few
compounds have been studied from microalgae compared with other microorganisms
such as bacteria and fungi (Olaizola 2003). However, as they have the ability to
produce a multitude of unique compounds there is increasing interest in their
research and exploitation (Cardozo et al. 2007). As discussed in Section 1.2.2, some
species produce large quantities of LC n-3 PUFAs, such as EPA and DHA. Better
understanding of lipid production and modification in microalgae will enable
increased use of these organisms for the commercial production of specialised lipids.
Two main applications have driven research in algal LC n-3 PUFAs. The first of
these is the use of omega-3 in aquaculture feeds for juvenile molluscs, crustaceans
and fish (Borowitzka 1997). Multicellular organisms such as molluscs and fish have
a limited capacity for LC n-3 PUFA production, obtaining the majority of omega-3
through their diet. These FAs are important in animal health (Rico-Villa et al. 2006).
To ensure that aquaculture-reared organisms are healthy and contain the same lipid
biochemistry as their wild counterparts, they must be provided with these compounds
in their diet. This is achieved primarily by feeding them cultures of LC n-3 PUFA-
producing microalgae. The second main application is the use of microalgal oils as a
sustainable alternative to fish oil, for human consumption and health. Microalgal oil
is suitable for consumption by vegetarian and vegan consumers who want to
supplement their diet with EPA and DHA. A range of algae-derived commercial lipid
products are available, making up approximately 3-5% of the over-all nutraceutical
omega-3 market (Turchini et al. 2012).
Currently, the cost of microalgal culture is the main limitation to the expansion of
microalgal oil production for the above purposes (Borowitzka 1997; Naylor et al.
2009). Accordingly, research that furthers the understanding of microalgal lipid
CHAPTER III
55
metabolism is of interest for improving the efficiency of culture practices. The
versatility of microalgal lipid metabolism and production has been studied
increasingly in recent years. It is well established that their lipid composition changes
throughout the growth cycle (Alonso et al. 2000). Lipid production can also be
altered by environmental factors such as temperature, light and gas composition (eg:
nitrogen, carbon dioxide) (Fidalgo et al. 1998; Guedes et al. 2010; Yongmanitchai &
Ward 1991). The presence of certain lipid-modifying enzymes is required for
changes in lipid production and utilisation to take place. Elongases and desaturases
are instrumental in the organisms’ ability to synthesise LC n-3 PUFAs. Enzymes
involved in the modification/catabolism of TAG, such as lipases, are also of potential
utility. Throughout nature, organisms that utilise TAG as an energy source possess
lipases that allow for the hydrolytic release of FAs. Different lipases possess
different specificities depending on how they have evolved for interaction with
various FAs.
In this work we hypothesised that algal species that contain large quantities of LC n-
3 PUFAs may possess lipases with selective omega-3-related activities. Lipases with
specific activity for or against the hydrolysis of LC n-3 PUFAs are industrially
important, as they can enable the more specific concentration of omega-3 FA for
nutritional and pharmaceutical applications. The research described in this chapter
aimed to first investigate the growth of Chaetoceros calcitrans, Isochrysis sp.
(T.ISO) and Pavlova lutheri and second, to compare FA profiles of lipid classes
during the growth of these species. These microalgae were chosen as they were
readily cultured by our collaborators (Cawthron Institute). The organisms are also
known to produce EPA and DHA and as such were appropriate for the investigation
of LC n-3 PUFA-specific lipases. Each of the three species was periodically sampled
throughout their growth. The total lipid was extracted, lipid content per cell
established, lipid classes fractionated and FA composition for each respective class
determined. From these data, changes and trends in the lipid composition and
distribution in the algal cultures was assessed throughout their growth. It was
hypothesised that by monitoring the lipid distribution and composition during
growth, it could be possible to determine when lipases are active in the organisms.
As defined from the lipid classes, FA data was then analysed to determine if omega-
3-related lipase activity was present at different time points. Some of the lipid
analysis was performed by a colleague (Kathrin Stähler).
CHAPTER III
56
The results showed that this approach was not appropriate to determine when lipases
are active in these microalgae. Despite this, the data generated does provide
information that may be relevant to the use of these organisms in aquaculture. C.
calcitrans, Isochrysis sp. and P. lutheri are all currently utilised as feeds in the
aquaculture industry. As such, advancements in the understanding of lipid
metabolism in these organisms are of interest for the improvement of feed quality.
3.1 Growth of Chaetoceros calcitrans, Isochrysis sp. and
Pavlova lutheri
When grown under the same conditions (Section 2.1.1), the number of cells for each
of the microalgae species, Chaetoceros calcitrans, Pavlova lutheri and Isochrysis
sp., exhibited a growth curve typical of batch grown microorganisms (Figure 3.1).
That is, phases proceeding through lag, logarithmic, stationary and regression. All
three species underwent a short lag phase of 1 to 3 days, in which the cells adjusted
to their new environment. Subsequently the algae began dividing rapidly and
entering into the logarithmic growth phase. The cell density of C. calcitrans on day 1
was 1.4 x106 cells mL-1 and increased to 18.7 x106 cells mL-1 on day 5 of the trial,
before declining on day 12. This growth profile of C. calcitrans was similar to that
reported elsewhere (Ragg et al. 2010). Isochrysis sp. grew from an initial cell
concentration of 4.2 x 105 cells.mL-1 on day 1, to approximately 19 x 106 cells.mL-1
during the stationary phase before declining on day 17. The length of lag and
logarithmic growth periods is similar to that previously reported (Fidalgo et al. 1998;
Kaplan et al. 1986). P. lutheri displayed the longest observed time to reach the
regression phase. It grew from an initial cell concentration of 4.2 x 105 cells.mL-1 on
day 1, increasing to approximately 20 x 106 cells.mL-1 during the stationary phase,
before declining on day 70. This stationary growth period (~40 days) was unusually
long under batch culture conditions. Despite all three replicate cultures showing a
consistent growth pattern (suggesting the data is accurate), the slow growth times
observed for P. lutheri in this study are unlike any found in the literature. Most
previous research presents much shorter logarithmic and stationary growth periods
(Emdadi & Berland 1989; Tatsuzawa et al. 1995).
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57
Figure 3.1 Growth curves of C. calcitrans, Isochrysis sp. and P. lutheri with labels indicating
sampling points. Lag (LA), early logarithmic (EL), logarithmic (L), late logarithmic (LL), early
stationary (ES), stationary (S) and late stationary (LS). (A) C. calcitrans, (B) Isochrysis sp. and (C) P.
lutheri.
0
5
10
15
20
25
30
0 2 4 6 8 10 12 14
cell
x 10
6 /m
L
growth period (days)
0
5
10
15
20
25
30
0 2 4 6 8 10 12 14 16 18 20
cells
x 1
06 /m
L
growth period (days)
0
5
10
15
20
25
30
0 10 20 30 40 50 60 70 80
cells
x 1
06 /m
L
growth period (days)
A
B
C
EL
LA
LA
EL
L
LL
ES
S
LS
EL
L
LL
L
LL
ES
S LS
LS ES
S
CHAPTER III
58
A study by Dunstan et al. (1993) did grow P. lutheri to 40 days. However, this
culture was maintained under semi-continuous growth until day 30 before being
allowed to continue under batch conditions. Another batch culture study of P. lutheri
by Brown et al. (1993) also gives a similar logarithmic growth profile to that seen
here. However, the difference in stationary growth is unknown as they harvested the
culture before it went into regression. Likely contributors to the variation between
growth profiles are the differences in growth conditions and media used in the
experiments. Although no fully comparable results can be located in the literature, P.
lutheri is capable of growing for longer periods of time in continuous systems
relative to the other species (personal communication, Nicky Roughton, Cawthron
Institute). This indicates that these findings are consistent with the nature of this
organism’s growth. However, further studies are needed to confirm this.
3.2 Lipid content and class in Chaetoceros calcitrans, Isochrysis sp.
and Pavlova lutheri
To investigate the lipid distribution and composition of the microalgae, culture
samples were taken at times denoted on the growth curves (Figure 3.1); which were
approximately lag (LA), early logarithmic (EL), logarithmic (L), late logarithmic
(LL), early stationary (ES), stationary (S) and late stationary (LS). Total lipid (TL),
was isolated from each sample (Section 2.1.2) and was subsequently used to
calculate the amount of lipid (pg/cell) for each species over the growth period. TL
was then separated by SPE chromatography into the fractions of neutral lipid (NL),
glycolipid (GL) and polar lipid (PL). The amount of NL, GL and PL (pg/cell)
throughout the growth of each species was determined gravimetrically. The lipids
present in the three different lipid classes can be generalised as follows. NL are
uncharged lipids, of which the most common species in algae are TAG, FFA and
various sterols (Volkman et al. 1989). PL primarily consists of phospholipids which
form the various membrane structures throughout the cell. GL differ from other PL at
the sn-3 carbon of the glycerolipid backbone. While phospholipids (main type of PL)
contain various phosphate based moieties, GLs possess a carbohydrate moiety; most
commonly galactose either as a monomer (monogalactosyldiacylglycerol) or dimer
(digalactosyldiacylglycerol). A large amount of GL reside in the chloroplast
membranes where they play important roles in cell signalling and energy production
CHAPTER III
59
(Dormann & Benning 2002). The basic structures of the primary lipid species (TAG,
phospholipid and galactolipid) in the three lipid classes are provided in Figure 3.2.
Investigating changes in these lipid classes was used to determine possible lipase
activity. The quantity and composition of the respective lipid classes for each species
are discussed in the following sections.
Figure 3.2 Chemical structures of triacylglycerol (A), phosphatidylcholine (B) and
monogalactosyldiacylglycerol (C).
A
B
C
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60
3.2.1 Lipid content and classes of Chaetoceros calcitrans
The lipid class profiles over the growth of C. calcitrans are displayed in Figure 3.2.1.
The TL content of C. calcitrans cells was between 3.5 and 7.4 pg/cell over the 12
day growth period. TL fluctuated in the first 6 days of growth before significantly
increasing during the stationary phase. This increase can primarily be attributed to an
increase in NL, from 1.7 ±0.2 to 4.1 ±1.1 pg/cell on days 7 and 12, respectively.
Analysis of the NL fraction by Iatroscan demonstrated that it was mostly TAG (data
not shown). TAG is primarily used by cells as a FA store/reservoir. This occurrence
has been shown previously with various microalgal species (Dunstan et al. 1993;
Emdadi et al. 1989). It suggests that C. calcitrans is storing NL in the form of TAG
during stationary growth, possibly in preparation for the onset of the regression
phase. It is also possible that TAG may accumulate so that when conditions improve,
TAG is readily available for use in membrane synthesis and energy production.
These processes are required in the initial rapid cell division of the next growth cycle
for those individual cells that survive (Dunstan et al. 1993). The initial major lipid
class (66.2% of total lipid) was the GL fraction (3.9 ±0.9 pg/cell), which decreased to
1.8± 0.6 pg/cell by day 6. The accumulation of NL during the stationary phase to late
stationary (day 12, 4.1 ±1.1 pg/cell) meant that it became the major lipid class
(54.5% of TL). NL decreased between day 4 and 6, from 2.18 pg/cell to 1.31 pg/cell,
respectively. This decrease may signify the presence of lipase activity in the cells
and/or membrane synthesis. However, GL also decreased slightly, suggesting that the
drop in NL isn’t merely a result of lipid rearrangement in the cells. The PL fraction
was the minor class throughout the growth cycle and changed little, with 0.3-0.5
pg/cell equating to 6.7-11.1% of TL in the cells. This indicates that the amount of
plasma and organelle membranes remained relatively constant throughout growth.
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61
Figure 3.2.1 Lipid class composition over the growth of C. calcitrans. (A) The amount (pg/cell)
of total lipid (TL), neutral lipid (NL), glycolipid (GL) and polar lipid (PL) over the growth period
(days). (B) The percentage of NL, GL and PL at each defined growth phase of early logarithmic (EL),
logarithmic (L), late logarithmic (LL), early stationary (ES), stationary (S) and late stationary (LS).
0
1
2
3
4
5
6
7
8
9
0 2 4 6 8 10 12
pg o
il/ce
ll
Growth period (day)
TL
NL
GL
PL
0
20
40
60
80
100
EL L LL ES S LS
Lipi
d cl
ass (
%)
Growth phase
NL
GL
PL
A
B
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62
3.2.2 Lipid content and classes of Isochrysis sp.
Figure 3.2.2 shows the amount (pg/cell) and percentage of lipid class at each time
point over the growth of Isochrysis sp.. TL ranged from 27.3-6.7 pg/cell with an
average of 11.7 pg/cell over the growth period of 17 days. The amount of TL
(pg/cell) was highest in the lag phase before declining rapidly at the onset of
logarithmic growth. This may be a result of the starting cells being larger and
therefore containing more lipid per cell. Thus as the cells begin to divide in
logarithmic growth the lipid (pg/cell) decreases. The amount of TL then steadily
increased from day 7 to 17, shifting from 2.7 ±0.6 to 7.9 ±1.6 pg/cell, respectively.
As was observed in C. calcitrans, this increase in TL was predominantly due to a rise
in NL. As mentioned in Section 3.2.1, it would appear that many microalgae undergo
this accumulation of TAG during the stationary growth. The initial major lipid class
was NL (14.0 ±2.2 pg/cell) making up 51.1% of all lipids, this subsequently
decreased to 41% (2.7 ±0.6 pg/cell) in the logarithmic phase, while GL increased to
48.5% (3.2 ±0.3 pg/cell). NL was at its lowest between day 4 and 9, at ~3.2 pg/cell. It
is difficult to attribute this to lipase activity, as the major decrease in lipid (pg/cell)
from lag to logarithmic growth is more likely explained by the rapid cell division.
Furthermore, from day 7 onwards the NL content increased steadily, which is not
indicative of lipase activity. As growth advanced into late stationary phase, NL again
became the major class continuously increasing to reach 60.4% of TL. PL again
remained the minor class throughout growth, averaging <10% of TL.
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63
Figure 3.2.2 Lipid class composition over the growth of Isochrysis sp.. (A) The amount (pg/cell)
of total lipid (TL), neutral lipid (NL), glycolipid (GL) and polar lipid (PL) over the growth period
(days). (B) The percentage of NL, GL and PL at each defined growth phase of lag (LA), early
logarithmic (EL), logarithmic (L), late logarithmic (LL), early stationary (ES), stationary (S) and late
stationary (LS).
0
5
10
15
20
25
30
0 5 10 15
pg o
il/ce
ll
Growth period (day)
TL
NL
GL
PL
0
20
40
60
80
100
LA EL L LL ES S LS
Lipi
d cl
ass (
%)
Growth phase
NL
GL
PL
A
B
CHAPTER III
64
3.2.3 Lipid content and classes of Pavlova lutheri
Lipid class profiles over the growth of P. lutheri are displayed in Figure 3.2.3. A
substantial amount of time elapsed between the last two samples that were taken
(days 21 and 72). This occurred as the growth cycle duration for this species was
underestimated. The observed duration was significantly longer than previously
reported. Lipid samples taken from within this area of growth would give a more
informative analysis of P. lutheri lipid production and composition. Thus the
analyses provided for this species really only investigates lipid production and
composition over the lag and logarithmic growth phases. At an average of 17 pg/cell
P. lutheri contained the greatest lipid mass of the three species. This is consistent
with results from Volkman et al. (1989), although their cultures contained less total
lipid. TL ranged from 49.8 ±10 to 9.3 ±0.8 pg/cell and as was seen in Isochrysis sp.,
the sample taken during the lag phase contained the greatest amount of lipid
(pg/cell). Therefore, the same explanation is likely with regard to cell size and lipid
content. This is comparable to the findings of Emdadi et al. (1989), where lipid (μg
mm-3 of biovolume) was shown to be greatest in lag growth. However, since the
majority of studies investigating lipid class over time take their first sample in
logarithmic growth (Fernandezreiriz et al. 1989; Fidalgo et al. 1998), it is difficult to
determine in which phase maximal lipid production is achieved.
P. lutheri differed from Isochrysis sp. in that GL was present as the major lipid class
for the duration of growth, ranging from 47.7 to 51.2% of TL. NL and GL have
previously been found to be the major constituents of P. lutheri (Meireles et al.
2003). However, the study by Meireles et al. (2003) showed NL was more prevalent
than GL. Again, this may come down to discrepancies in growth conditions. As seen
in Isochrysis sp., the NL in P. lutheri decreased significantly from the lag phase.
Between days 8 and 18, NL was present at ~4.0 pg/cell. Variations in the amount of
lipid in all classes appeared to follow the same general trend as the TL, with no
individual lipid class fluctuating more than 15% for the entirety of growth. The PL
class was the least prevalent again, reaching a maximum of 13.4% of TL during
logarithmic growth. The lipid class profiles obtained for P. lutheri, Isochrysis sp. and
C. calcitrans did not give clear-cut decreases in NL. This was surprising as cells
entering into regression were expected to begin utilising stored TAG in order to
survive, requiring lipase to be expressed.
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65
Figure 3.2.3 Lipid class composition over the growth of P. lutheri. (A) The amount (pg/cell) of
total lipid (TL), neutral lipid (NL), glycolipid (GL) and polar lipid (PL) over the growth period (days).
(B) The percentage of NL, GL and PL at each defined growth phase of lag (LA), early logarithmic
(EL), logarithmic (L), late logarithmic (LL), early stationary (ES), stationary (S) and late stationary
(LS).
0
10
20
30
40
50
60
70
0 15 30 45 60 75
pg o
il/ce
ll
Growth period (day)
TL
NL
GL
PL
0
20
40
60
80
100
LA EL L LL ES S LS
Lipi
d cl
ass (
%)
Growth phase
NL
GL
PL
A
B
CHAPTER III
66
3.3 Fatty acid composition of Chaetoceros calcitrans, Isochrysis sp.
and Pavlova lutheri
Fatty acid (FA) composition was analysed in order to better understand the lipid
metabolism of these microalgae. Following the separation of NL, GL and PL classes,
the FA composition of each fraction was analysed. The NL fraction was of most
interest for the investigation of lipase activity, since lipases act on TAG. It was
initially hypothesised that this analysis may provide information on the expression of
lipases with omega-3-related activities. However, due to the difficulties in
determining lipase activity (from changes in NL) in the first place, this approach was
not possible. Regardless, trends in the FA compositions of each lipid class can
provide insight into microalgal lipid metabolism, some of which may be useful for
other applications, such as aquaculture. The data presented here gives a more
comprehensive (covering lag, logarithmic, stationary and regression phases) lipid
profile over batch culture than provided by previous studies. Most previous works
only report analyses for 1 to 3 time points (Dunstan et al. 1993; Fernandezreiriz et al.
1989; Fidalgo et al. 1998; Tatsuzawa et al. 1995; Volkman et al. 1989; Whyte 1987).
By analysing more time points and separating the lipid classes, we provided a more
in-depth analysis of the origins of changes to lipid composition in each microalgal
species. These data also provide information relevant to the growth of these
organisms for aquaculture feeds. That is, when LC n-3 PUFAs are most concentrated
in the cell. An overview of the FA composition of each species is provided and
discussed in the following sections.
3.3.1 Fatty acid composition of Chaetoceros calcitrans
The FA composition of the TL in C. calcitrans throughout its growth cycle is shown
in Appendix 8.1, Table 8.1.1. Initially saturates were the major FA class (52.4%),
predominantly consisting of myristic acid (C14:0, 32.2%) and palmitic acid (C16:0,
16.4%). Myristic and palmitic acids have previously been found to be the major
saturated FAs in C. calcitrans (Volkman et al. 1989). The amount of saturated FA
decreased significantly throughout the logarithmic and early stationary phases to
16.5% on day 6. The final sample taken saw saturated FA increase again to 33.5% of
lipid, driven primarily by an increase in palmitic acid, which accounted for 19.6% of
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67
TL in the late stationary sample. Monounsaturated fatty acids (MUFAs) increased
from 5.1% on day 1 to 28.9% on day 2, with a large increase (5.1 to 27.6%) in
palmitoleic acid (C16:1 n-7). This was also found to be the major MUFA species by
Volkman et al. (1989). From day 2, the major FA class was PUFAs. Initially the
major PUFAs were DHA (C22:6 n-3, 15.7%) and C22:2 n-6 (19.5%). DHA was only
observed in the total lipid during the early logarithmic growth, before decreasing to
2.3% in the logarithmic phase. DHA has previously been shown to be present in only
small quantities (<2.3% TL) in C. calcitrans (Pettersen et al. 2010; Ragg et al. 2010).
However, to our knowledge no previous studies have analysed the lipid of this
species this early in culture development. The results shown here demonstrate that
DHA is present in early culture growth, before being depleted rapidly. Subsequently,
DHA was a minor component for the remainder of the culture growth cycle. From
day 2 onwards the major PUFAs were C16:2 n-6, C16:2 n-4, C16:3 n-4 and EPA
(C20:5 n-3), consistent with that observed by Volkman et al. (1989). EPA increased
significantly from 1.4% on day 1 to 23.0% on day 2. The amount of EPA then
remained stable throughout growth until declining to 15.8% in the late stationary
phase.
The FA composition of the NL fraction in C. calcitrans over the growth period is
shown in Appendix 8.1, Table 8.1.2. In early logarithmic growth saturated FAs were
the major FA class (51.1%). The proportion of saturated FA decreased during
logarithmic growth to only 14.1% on day 6, before increasing to 36.7% in late
stationary phase. MUFAs in the NL were comprised almost exclusively of
palmitoleic acid, which increased two fold after day 1, from 10.4% to 22.2% on day
2, which was the major FA change in this period. PUFA increased throughout the
logarithmic growth phase (38.5 to 62.1% from day 1 to day 6 respectively) and
decreased during the late stationary phase (35.6%). The major PUFAs in the NL
fraction were EPA, C16:2n-6 and C16:3n-4. EPA was most abundant between day 2
and day 4 (31.4-33.4%), while concentrations were lower in early (7.5%) and late
growth (16.2%). DHA in the neutral lipid fraction decreased significantly after day 1
(9.9 to 1.1%), indicating a rapid use of this FA by the algae in early culture
development.
GL FA composition in C. calcitrans over the growth period is shown in Appendix
8.1, Table 8.1.3. The major FA class throughout the trial was PUFA (48.8-55.3%), of
which C16:2 n-6, C16:2 n-4, C16:3 n-4 and EPA were the major FA species. This
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68
was unsurprising since GL makes up a large amount of lipid in the chloroplast, where
FA synthesis and modification takes place (Marechal et al. 1997). Recent work by
Liang et al. (2013) has shown that EPA synthesis takes place in the chloroplast of the
marine diatom species Fistulifera. EPA in the GL fraction was constant over its
growth, averaging 14.9% over all samples taken. DHA decreased significantly
throughout the growth period from day 1 (from 11.4 to ≤ 2.0%). MUFA was again
composed primarily of palmitoleic acid, which remained relatively stable before
decreasing in the late stationary growth. Saturated FA decreased significantly from
day 1 to day 2 (24.6 to 14.4%), then steadily increased again to reach 28.9% on day
12.
FA composition in the PL fraction of C. calcitrans is shown in Appendix 8.1, Table
8.1.4. In early logarithmic growth the majority of FAs were saturates (52.1%). This
decreased rapidly on day 2 as the proportion of MUFAs and PUFAs in the PL
increased. Palmitoleic acid was the major MUFA, comprising approximately 30-50%
of all FA in PL from logarithmic to late stationary growth. The PUFA content of the
PL fraction decreased over the logarithmic growth phase (47.9% lipid on day 1 to
33.2% on day 4). This is mainly due to the decrease of the two most prevalent
PUFAs, DHA and C22:2 n-6. The amount of DHA relative to other FA was the
highest in the PL fraction, indicating that DHA plays an important role in the
membrane chemistry of C. calcitrans. It is well known that DHA plays vital roles in
membrane structure and function in a range of organisms (Stillwell et al. 2003). EPA
was also a major PUFA in PL and whilst not being detected on day 1, its levels
remained constant throughout the rest of the growth period (12.6-17.0%). Apart from
EPA, a significant proportion of the PUFAs found in the PL fraction was comprised
of C16 PUFAs.
3.3.2 Fatty acid composition of Isochrysis sp.
The FA profile of the TL present in Isochrysis sp. over the course of its growth is
shown in Appendix 8.1, Table 8.1.5. Saturated FAs initially formed the majority of
TL (44.0%), of which myristic acid (21.8%) and palmitic acid (17.9%) were the
primary species. This is consistent with previous studies (Dunstan et al. 1993;
Fidalgo et al. 1998; Volkman et al. 1989). The amount of saturated FA decreased
significantly during early logarithmic growth, to 32.3% on day 7. Total MUFA
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69
concentrations remained relatively unchanged throughout the growth of Isochrysis
sp.; ranging from 24.7-30.8%, with the main FA being oleic acid (C18:1 n-9). There
is some discrepancy in the literature over which MUFA is most common. Some
authors record palmitoleic acid as the most prevalent (Volkman et al. 1989), while
other trials in the same laboratory report oleic acid as the most abundant (Dunstan et
al. 1993). PUFA were initially the minor FA class at 26.1% of TL. However, they
increased significantly to 43.0% on day 7 before gradually decreasing to 30.4% for
the remainder of growth. The major FA species present were linoleic acid (C18:2 n-
6), -linolenic acid (ALA, C18:3 n-3), stearidonic acid (SDA, C18:4 n-3) and DHA.
EPA was only observed in minute amounts (<0.6%) in the early stages of logarithmic
growth which is consistent with previous studies of the Isochrysis sp. T.ISO strain
(Roncarati et al. 2004). The PUFA profiles found here are comparable with data
published previously (Dunstan et al. 1993; Volkman et al. 1989).
FAs present in the NL fraction of Isochrysis sp. over the duration of its growth are
shown in Appendix 8.1, Table 8.1.6. Saturates were the dominant FA class for most
of the growth period, exceptions being the logarithmic and late logarithmic samples,
where PUFA and MUFA were most prevalent, respectively. Myristic and palmitic
acids were again the major saturates species. MUFA in the NL ranged from 32.5-
37.5%, remaining relatively unchanged throughout the entire growth period. Oleic
acid was the major MUFA making up approximately one third of all NL. The PUFA
content increased significantly from 17.0% on day 1 to 35.4% on day 7, before
decreasing to 25.9% in late logarithmic growth (day 9). Linoleic acid, SDA and
DHA made up the major FA present in this fraction. DHA underwent an increase
from 3.2% in the lag phase to 9.6% in late logarithmic growth, then steadily
decreasing to 4.2% in the late stationary phase. C16:2 n-6 and ALA were also
present, with concentrations peaking during logarithmic growth.
The FA profile of the GL fraction in Isochrysis sp. over the course of its growth is
shown in Appendix 8.1, Table 8.1.7. On day 1 saturated FA were the major species
present (42.7%) and primarily consisted of myristic and palmitic acids. The saturate
content declined significantly throughout logarithmic growth with a concurrent
increase in PUFA levels. MUFA were the least prevalent FA class throughout the
duration of growth. The major MUFAs were again palmitoleic and oleic acids.
PUFA represented the major FA class in GL for the majority of the growth period
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70
accounting for approximately half the FA during stationary growth. The amount of
PUFA on day 1 (38.7%) steadily rose to 53.8% on day 14. The primary PUFAs were
linoleic acid, ALA, SDA and DHA. SDA showed the greatest change in
concentration, significantly increasing from 17.3% on day 1 to 27.9% on day 17.
This culture of Isochrysis sp. contained very little EPA in the GL when compared to
the other two species. However, it did contain an abundance of SDA. GL is known to
be involved in PUFA synthesis in the chloroplast (Marechal et al. 1997). It would
appear that the 5-elongase and 5-detaturase enzymes required for the conversion
of SDA to EPA may act slowly, in this strain of Isochrysis sp.. The presence of DHA
indicates that flux through the PUFA biosynthetic pathway is occurring, although the
specific biosynthetic pathway in this organism is still unknown.
The FA composition of the PL fraction in Isochrysis sp. during its growth cycle is
shown in Appendix 8.1, Table 8.1.8. At all stages of growth PUFA was the major FA
class present (41.5-48.8%). The PUFA species were composed of low amounts of
linoleic acid, ALA and SDA. DHA was present at higher levels ranging from 23.3 to
38.0% of PL, indicating the importance of DHA in membrane biochemistry in this
organism. Saturated FA was the second most prevalent class (33.7-42.7%)
throughout growth, with myristic and palmitic acids being the most abundant. MUFA
was the least abundant FA type and primarily consisted of oleic acid.
3.3.3 Fatty acid composition of Pavlova lutheri
The FA profile in the TL over the growth cycle of P. lutheri is shown in Appendix
8.1, Table 8.1.9. Saturated FA concentrations were the highest (40.1%) in the early
stages of growth (lag), before decreasing to 23.3% in early logarithmic growth. This
occurred concurrently with an increase in PUFA, primarily EPA. The major saturates
were myristic and palmitic acids. MUFA formed the minor FA class in the TL,
ranging from 11.5 to 18.3%, which was primarily palmitoleic acid. PUFA formed the
majority of FA present in P. lutheri over the growth cycle. In logarithmic growth
(day 8) PUFA accounted for 67.9% of TL. The most prevalent FA species were
SDA, EPA and DHA. The significant increase in PUFA concentration can be
attributed to increases in EPA (19.4% on day 1 to 36.3% on day 8). The amount of
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71
DHA remained relatively consistent throughout logarithmic growth until reaching
stationary growth where DHA then increased from 10% to 20.5%.
The NL FA profile of P. lutheri over its growth is shown in Appendix 8.1, Table
8.1.10. Both saturated FA and MUFA followed similar trends over the growth period
starting in the lag phase at 28.0% and 21.4%, respectively. The saturated FA and
MUFA content then decreased to 15.5% and 12.8% in early stationary growth,
respectively. Amounts of FA then remained relatively stable during the logarithmic
phase, before increasing again in early stationary phase growth. The increase in
saturate FA and MUFA from logarithmic to stationary phase is consistent with that
found by Dunstan et al. (1993). The major FAs found were myristic, palmitic and
palmitoleic acids. As was found in the TL analysis, PUFA was the main class of FA
present, averaging 57.6% over the growth cycle. PUFA increased significantly from
50.6% on day 1 to 75.9% on day 8 (logarithmic). From day 8 the general trend was a
decrease in PUFA with the final sample taken in late stationary growth showing a
low of 39.7%. The main FAs that contributed to these changes in PUFA levels were
C16:2 n-6 (4.9% on day 1, 15.6% on day 8, 4.5% on day 72), SDA (5.9% on day 1,
10.0% on day 8, 2.3% on day 72), EPA (15.9% on day 1, 26.2% on day 8, 15.1% on
day 72) and DHA (11.2% on day 1, 15.6% on day 8, 10.4% on day 72). This influx
of shorter less saturated FA is consistent with cells preparing for starvation
conditions, or the onset of more favourable conditions and the next stage of rapid cell
division (Dunstan et al. 1993).
The FA profile of the GL present in P. lutheri over its growth cycle is shown in
Appendix 8.1, Table 8.1.11. FA in the GL class also showed both saturate FA and
MUFA following similar trends. Both underwent initial decreases from lag to early
logarithmic growth and then remained relatively unchanged for the remainder of
growth. The resulting PUFA profile showed an increase from 42.8% in the lag phase
to 60.5% in early logarithmic growth. Following this initial increase PUFA
fluctuated little (62.1-66.2%). From day 8 onwards EPA was the major PUFA
comprising >40% of all FA present in GL. SDA was also prevalent, averaging 11.8%
of GL. As observed in C. calcitrans, it would appear that GL is involved in the
biosynthesis of EPA in this species also.
The PL FA composition present in P. lutheri is shown in Appendix 8.1, Table 8.1.12.
The saturated FA in the PL decreased in the lag phase from 56.6 to 27.2% during the
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72
late stationary growth. This was primarily due to a decrease in palmitic acid (39.5%
to 18.7%). PUFA levels increased, with 32.4% in the lag phase increasing to 68.3%
in the late stationary phase. The decrease in palmitic acid was met by a
corresponding increase in EPA and DHA which were the major PUFA species
present. Amounts of EPA ranged from 14.5 to 26.6% of PL for the duration of
growth, while DHA showed an even greater range of 7.0 to 42.4%. In late stationary
growth it was found that the LC n-3 PUFAs made up ~67% of all FA in the PL. This
was the highest concentration of LC n-3 PUFAs found in the PL of the three species
investigated. Recent studies suggest LC n-3 PUFA bioavailability may be increased
in phospholipid forms (Schuchardt et al. 2011), thus P. lutheri may provide a
promising source of omega-3 for inclusion into nutraceuticals.
3.4 LC n-3 PUFA content in the neutral lipid of Chaetoceros
calcitrans, Isochrysis sp. and Pavlova lutheri
As NL in these species is composed almost entirely of TAG, the lipid quantity and
FA composition of this lipid class was of primary importance when investigating
potential lipase activity. Fluctuations in the concentration of EPA and DHA may also
be indicative of omega-3-related lipase activity. However, the NL profiles observed
could not readily be interpreted in terms of lipase activity. That microalgal lipid
metabolism is relatively poorly understood also made interpretation more difficult
(Khozin-Goldberg et al. 2011). To view the trends and particular enzymatic
influences in the quantity of EPA and DHA in NL, the data were plotted both as a
percentage and an amount (pg/cell) over the growth cycle of each species (Figures
3.4.1, 3.4.2 and 3.4.3).
As discussed in Section 3.3.1, C. calcitrans contained very little DHA, except for on
day 1. EPA was the major LC n-3 PUFA in this species. When the amount of these
FAs is expressed as a percentage stationary growth shows increased EPA in the TL.
This is mirrored in the NL fraction (Figure 3.4.1A). EPA in the TL then steadily
decreases until the late stationary phase. The NL EPA also follows the same general
decrease. There is a sharp dip within this decrease, which may indicate lipase
activity. However, the error bars are relatively large for this data point and it may be
an anomaly. When EPA in the NL is plotted as an amount (pg/cell) the contour
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73
tracks EPA in the TL over the growth cycle (Section 3.4.1B). This indicates changes
in the concentration of other FA are the major influence and that changes in EPA are
simply representative of overall lipid change. Looking at the FA profile of the early
stationary sample (Appendix 8.1, Table 8.1.2), it is clear that a number of FAs
(C16:2 n-6, oleic acid and linoleic acid) increased significantly in this phase,
although the sample standard deviation is large. The concentration of LC n-3 PUFAs
in the NL fraction remained unchanged, indicating that limited omega-3 lipase
activity was taking place.
Figure 3.4.1 EPA and DHA content in the total lipid and neutral lipid of C. calcitrans over the
duration of growth. (A) Shows content as a percentage of FAs in each respective fraction, (B) shows
to amount (pg/cell) of each FA in the respective fractions. Total lipid EPA ( ), neutral lipid EPA
( ), total lipid DHA ( ), neutral lipid DHA ( ).
0
5
10
15
20
25
30
35
40
0 50 100 150 200 250 300
Fatt
y ac
id (%
)
Time (hrs)
0
0.2
0.4
0.6
0.8
1
1.2
1.4
0 50 100 150 200 250 300
Fatt
y ac
id (p
g/ce
ll)
Time (hrs)
A
B
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74
In contrast to C. calcitrans, Isochrysis sp. contains DHA and very little EPA (Figure
3.4.2). When expressed as a percentage, DHA in both TL and NL increased in the lag
and logarithmic growth phases. From day 5 onwards DHA in the TL remained
relatively stable, while there is a general decrease in DHA in the NL. A reduction in
the percentage of DHA in NL occurred (Figure 3.4.2A), which may be due to lipase
hydrolysis or FA restructuring. However, as seen for C. calcitrans, when DHA is
plotted as a concentration (pg/cell) the profile changes dramatically.
Figure 3.4.2 EPA and DHA content in the total lipid and neutral lipid of Isochrysis sp. over the
duration of growth. (A) Shows content as a percentage of FAs in each respective fraction, (B) shows
to amount (pg/cell) of each FA in the respective fractions. Total lipid EPA ( ), neutral lipid EPA
( ), total lipid DHA ( ), neutral lipid DHA ( ).
0.0
2.0
4.0
6.0
8.0
10.0
12.0
14.0
0 100 200 300 400 500
Fatt
y ac
id (%
)
Time (hrs)
0
0.5
1
1.5
2
2.5
0 100 200 300 400 500
Fatt
y ac
id (p
g/ce
ll)
Time (hrs)
A
B
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75
While initially decreasing markedly, the amount of DHA present in the TL remained
stable until day 8, after which DHA increased steadily. The profile of NL DHA is
similar, but remained more stable throughout the growth cycle. Therefore, changes in
the amount of other FA species, and not lipase activity, are the major influence
causing change in the relative amount of DHA in the NL. Looking at the FA profile
of the logarithmic and stationary samples (Appendix 8.1, Table 8.1.6) fluctuation in
C18 FAs was the major cause of the observed relative decrease in DHA.
P. lutheri differed from the other species in that it contained relatively high amounts
of both EPA and DHA in its NL (Figure 3.4.3). When the amount of EPA in TL is
given as a percentage, lag and logarithmic growth phases show increases in EPA. On
day 14 EPA in the TL decreased significantly. A similar profile is seen in EPA
within NL, but the decrease begins after day 8. However, as with the other species,
when EPA is plotted as a concentration (pg/cell), the amount of EPA present in NL
follows the same profile as that of TL. In P. lutheri (Appendix 8.1, Table 8.1.10) it
appears that changes in C16 FA species (C16:0, C16:1, n-7, C16:2 n-2 and C16:4 n-
1) are the primary cause of the observed reduction of EPA relative to other FA in the
NL fraction. The same profile is seen in the amount of DHA present in this species.
Overall these results indicated that for all microalgal species studied, changes in
omega-3 content could not be clearly linked with omega-3-specific lipase activity.
This approach only gives an indication of lipid metabolism in the organisms. The
comparison of FA profiles from different lipid classes for demonstrating lipase
activity was not as useful as initially thought. To our knowledge no comprehensive
studies have been performed to analyse what happens to the biochemical
composition of algae once they reach the regression phase of culture. The NL content
of cells may decrease as cells in starvation begin to utilise the FAs stored as TAG, in
order to survive. The release of FAs for energy production would require lipase
activity. Our findings suggest that these species of algae do not utilise all stored TAG
once in regression, as NL (TAG) continued to increase into this phase of the growth
cycle (see Figures 3.2.1. and 3.2.2). It may be a lack of other nutrients, vital to
microalgae survival, which causes cell death no matter the amount of TAG stored for
use in energy production, membrane maintenance and other physiological uses. At
the other end of the growth spectrum (lag phase) NL decreases significantly. We
speculate that this is mainly due to a decrease in cell size. However, to some degree
it may also be due to the use of FA from TAG for energy and the synthesis of new
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76
membranes as the cells divide rapidly. While it is possible that lipases are active at
this time point, the volume of culture required to obtain sufficient biomass for lipase
isolation would make carrying this out extremely challenging.
Figure 3.4.3 EPA and DHA content in the total lipid and neutral lipid of P. lutheri over the
duration of growth. (A) Shows content as a percentage of FAs in each respective fraction, (B) shows
to amount (pg/cell) of each FA in the respective fractions. Total lipid EPA ( ),
neutral lipid EPA ( ), total lipid DHA ( ), neutral lipid DHA ( ).
0
5
10
15
20
25
30
35
40
45
0 500 1000 1500 2000
Fatt
y ac
id (%
)
Time (hrs)
0
2
4
6
8
10
12
14
16
0 500 1000 1500 2000
Fatt
y ac
id (p
g/ce
ll)
Time (hrs)
A
B
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77
As photoautotrophs microalgae can produce energy (adenosine triphosphate (ATP))
via both photosynthesis (photophosphorylation) and the catabolism of stored
carbohydrates and lipids (substrate-level phosphorylation and oxidative
phosphorylation). Under normal (un-stressed) growth conditions, carbohydrates are
the major carbon store in many species of microalgae (Chauton et al. 2013; Li et al.
2011), suggesting that glycolysis is the predominant form of energy-yielding
catabolism. Microalgae have been shown to possess the enzymes required for FA
catabolism and β-oxidation and thus can generate energy from lipids (Winkler et al.
1988). However, in microalgae FAs appear to be used less in energy production and
more as structural and functional compounds for membrane synthesis and
remodelling in the cell. TAG accumulation occurs in cells when conditions for
growth become limiting (stressed), suggesting FA are not the preferred energy source
under these conditions.
A wealth of genetic information on algal lipid metabolism has become available over
the last decade and as of 2011, seven algal genomes have been sequenced (Khozin-
Goldberg et al. 2011). Research on algal lipids has largely focussed on lipid
biosynthesis, accumulation and composition modification for applications in
biofuels, aquaculture and nutraceuticals. Investigation of the overall lipid metabolism
and expression profiles of related genes/enzymes in some species has come in the
form of transciptomic analysis (Guarnieri et al. 2011; Rismani-Yazdi et al. 2012).
These relatively new transciptomic approaches provide extremely powerful tools for
monitoring the expression of enzymes involved in different metabolic pathways. The
use of these methods in place of, or in conjunction with the lipid-based approach
employed here, may have provided a more accurate profile of lipase expression in
microalgae. However, relatively little work has been carried out on characterising the
enzymes involved in regulating lipid catabolism (Trentacoste et al. 2013). Lipases
from microalgae have been particularly neglected, which is emphasised by the fact
that to date no lipase of eukaryotic microalgal origin has been isolated and
characterised in full. The very nature of microalgal lipid composition makes them a
potentially interesting source of lipases. Many species contain large quantities of LC
n-3 PUFAs, which are important for structural and functional purposes (Khozin-
Goldberg et al. 2011). Metabolically it makes little sense for microalgae to break
down such valuable lipids for energy production when simpler saturated FAs are
available. Accordingly, it is possible that lipases present in some species have
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78
developed omega-3 related specificities. For example, it is possible that as LC n-3
PUFAs are relatively metabolically expensive for cells to synthesise, and they are
used for certain cellular structures or functions, that lipases may have evolved to
specifically not target these FAs in TAGs.
After investigating the lipid distribution and composition in these three algal species,
it was not possible to show the presence of specific lipase activity. Therefore, for
lipase isolation work, algae were harvested at a point that gave maximal biomass in
the shortest time duration. Cells were harvested at late logarithmic phase growth in
each species, this being days 6-8 for C. calcitrans, days 8-10 for Isochrysis sp. and
days 18-20 for P. lutheri.
3.5 Summary
Microalgae are capable of producing large quantities of the LC n-3 PUFAs EPA and
DHA. It was hypothesised that by analysing the lipid production and composition of
C. calcitrans, Isochrysis sp. and P. lutheri over their growth, it may be possible to
determine when lipase was being expressed. Information on lipid composition in
these species is also useful for aquaculture purposes. The duration of growth cycle
(inoculation to the onset of regression) varied for each species. These were 12, 17
and 72 days for C. calcitrans, Isochrysis sp. and P. lutheri, respectively. The duration
of P. lutheri growth was longer than any previously published examples. The total
lipid content was highest in P. lutheri and lowest in C. calcitrans. Lipid class
analyses of each species over the growth period showed that the PL content in all
species remained largely unchanged. The NL content increased significantly in C.
calcitrans and Isochrysis sp. from late logarithmic growth phase onwards. Therefore,
it was not possible to determine when the lipase activity was present, and further
analysis of LC n-3 PUFA content could not be used for indicating potential omega-3-
specific lipase activities. The FA composition and LC n-3 PUFA content of each
species was still relevant for other applications such as aquaculture. The primary LC
n-3 PUFA present in C. calcitrans was EPA, in Isochrysis sp. it was DHA, and P.
lutheri contained both EPA and DHA in similar quantities. LC n-3 PUFA content
was generally highest in early culture between lag and logarithmic phases. Cultures
were subsequently grown to the late logarithmic phase for lipase isolation.
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CHAPTER IV
Investigation of lipases from Isochrysis sp. and
Pavlova lutheri
4.0 Introduction
Research demonstrating the importance of LC n-3 PUFAs in human health,
especially EPA and DHA, has led to these compounds becoming commercially
significant. Accordingly, their use in food and nutraceuticals has steadily increased in
recent years. In 2012, the ‘Global Organisation for EPA and DHA Omega-3’
(GOED) estimated annual global consumer spending on products containing omega-
3 to be US$25.4 billion (Ismail 2013). Conventional methods for concentrating LC
n-3 PUFAs include urea complexation, fractional and molecular distillation,
winterisation and super critical fluid extraction (Bimbo 1998; Shahidi et al. 1998).
However, all of these methods have drawbacks (see Section 1.1.3). Lipases offer an
alternative method of concentration with substantial advantages. These include
reduced chemical and solvent usage, milder pH and temperature processing
conditions (and therefore lower energy inputs) and a more selective and targeted
approach. However, for this method of processing to become feasible new lipases
with specific activities for LC n-3 PUFA hydrolysis, or that select for FA other than
PUFA, are required. For commercial purposes desirable lipase activity for omega-3
concentration would involve either, excluding EPA and DHA hydrolysis completely
(i.e. concentrating LC n-3 PUFA in the glyceride fraction), or a targeted hydrolysis
of these compounds (i.e. specific removal of LC n-3 PUFAs from the hydrolysate).
Microalgae have been utilised as a source of biological compounds, including
polysaccharides, lipids, peptides and carotenoids (Cardozo et al. 2007). However, so
far relatively few enzymes, and no lipases, have been isolated and characterised from
these organisms. Microalgae contain many lipids, including EPA and DHA
(Harwood et al. 2009) which they are able to synthesise, not just acquire from their
diet. It may therefore be possible that lipases originating from these organisms
possess specific activities selective for or against the hydrolysis of LC n-3 PUFAs.
Currently no specific lipases have been identified that specifically hydrolyse LC n-3
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80
PUFAs from TAG substrates. However, LC n-3 PUFA specificity has been
demonstrated with the digestive juices of fish (Halldorsson et al. 2004a; Lie et al.
1985). The majority of lipases are inefficient at hydrolysing compounds such as EPA
and DHA because of the long chain-length and numerous unsaturated bonds in these
FAs (Wanasundara et al. 1998). For this reason a number of lipases have been shown
to concentrate LC n-3 PUFA in the acylglycerol fraction (Akanbi et al. 2013;
Hoshino et al. 1990; Wanasundara et al. 1998) (i.e. they hydrolyse FAs other than
PUFAs from the glycerol backbone).
With the initial investigation of algal growth and lipid production completed
(Chapter III), the lipases present in the microalgae were investigated. To our
knowledge there is currently no literature describing the isolation and biochemical
characterisation of a lipase from microalgae. Previous isolation of lipases from a
range of sources has been undertaken using a variety of methods. The most
commonly employed approaches are non-specific, such as ammonium sulfate
precipitation, ion-exchange chromatography, gel filtration chromatography and
hydrophobic interaction chromatography (Ghosh et al. 1996; Saxena et al. 2003).
Most of the time a single isolation step is not enough to achieve sufficient purity and
a combination of steps are generally used. One or more of these methods is used in
almost all published lipase isolation protocols. As the /β-hydrolase fold of lipases is
highly conserved (Holmquist 2000), it was assumed that these methods could also be
applicable for the isolation of lipases from microalgae.
This research aimed to demonstrate the presence of lipases in various microalgae and
then to isolate and characterise the new enzymes. Methodology was first established
to access intracellular protein. Multiple techniques were then tested for the
purification of lipase from the crude material. Lastly, enzymatic analysis of activity
optima and FA selectivity of lipases from different species were investigated.
4.1 Detection of lipases in microalgae
The presence of lipases in the microalgae was established using two methods; the p-
nitrophenyl (pNP) assay, and the 4-methylumbelliferyl-butyrate (MUB) gel-overlay
assay in conjunction with native-PAGE (i.e. gel zymography). The microalgal
species Isochrysis sp., P. lutheri and C. calcitrans were all tested for activity. Prior to
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the optimisation and testing of alternate methods for cell breakage, bead-beating
(Section 2.1.5.1) was used as a previous study found it achieved adequate cell
breakage (Lee et al. 1998). Further work involving cell lysis methods is presented
and discussed in Section 4.2.
The crude protein homogenate was clarified by centrifugation (25,000 x g for 15 min
at 4 C) prior to use in the experiments. Activity against pNP-palmitate (pNPP) was
measured (see Section 2.1.8). The release of long chain FAs such as palmitate upon
hydrolysis is usually indicative of lipase activity, while shorter more soluble
substrates are associated with esterase activity (Gupta et al. 2003). Hydrolysis of
pNPP was observed for homogenates obtained from Isochrysis sp. and P. lutheri, but
not C. calcitrans (data not shown). A possible reason for the lack of activity in C.
calcitrans is its cell structure. As a diatom, the organism possesses a frustule (porous,
rigid cell wall composed almost entirely of polymerised silicic acid (Schmid et al.
1981)). Relative to the other non-diatom species, this structure may have reduced the
effectiveness of the bead-beating method. Subsequently, further trials were only
performed with Isochrysis sp. and P. lutheri.
Crude protein homogenate from the two species was analysed by native-PAGE (see
Section 2.1.12). After electrophoresis was performed the gel was exposed to a 4-
methylumbelliferyl butyrate (MUB) gel-overlay assay (Section 2.1.14). Fluorescent
bands appeared on the gel after approximately 10 minutes, showing the position of
proteins capable of hydrolysing the fluorophore (4-methylumbelliferone) from
butyrate. Multiple bands were present in P. lutheri and Isochrysis sp. samples (Figure
4.1), suggesting that there are multiple lipolytic enzymes (possibly esterases), or
isoforms of such enzymes present. This result further confirmed the presence of FA-
hydrolysing enzymes, possibly lipases, in P. lutheri and Isochrysis sp.. The presence
of multiple enzymes of interest also indicated a need for separation before accurate
biochemical characterisation could take place.
Subsequent to the assays using MUB, attempts to elucidate lipase activity from that
of other lipid hydrolases were made using 4-methylumbelliferyl palmitate (MUP). As
discussed earlier, lipases typically hydrolyse longer insoluble FA substrates. Initially
the substrate was prepared as for MUB (Section 2.1.14). However, it was found that
MUP was not soluble in 2-methoxyethanol. Subsequently a range of organic solvents
and surfactants were trialled to solubilise the MUP. The solvent also had to be
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82
miscible with water, so the MUP emulsion could form once the stock substrate was
added to the assay buffer. Miscibility tables (Phenomenex) were used to aid in
solvents selection (see Appendix 8.2, Table 8.2 for solvents tested). Dioxane with
1% Triton X-100 was the only solvent combination found that solubilised MUP and
formed an emulsion. However, fluorescent bands were not observed when this
substrate was applied in gel-overlay assays. There are a number of possible
explanations for this: (1) The structure of MUP micelles is different than MUB
micelles and does not allow interaction/movement of the substrate into the gel to
permit lipase interaction; (2) lipase activity against MUP is too low to visualise or (3)
the presence of Triton X-100 is detrimental to lipase activity. The second explanation
is unlikely due to the high sensitivity of UV methods; likewise the amount of Triton
X-100 present in the overlaid substrate is minimal. Therefore it is likely that the
emulsion itself is the cause and further testing of this substrate is required.
Figure 4.1 4-Methylumbelliferyl butyrate gel-overlay assay performed on a basic native-
PAGE. Visualised under UV light. Lane 1; 30 μL of protein homogenate from P. lutheri, lane 2; 30
μL of protein homogenate from Isochrysis sp.. White dashed line marks the beginning of the resolving
gel. Black arrows indicate fluorescent bands.
1 2
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83
4.2 Algal cell lysis methods
Rupturing of the cell wall/membrane was required to release the cellular contents
before lipase isolation could be attempted. There are numerous methods that have
been trialled for achieving cell lysis in microorganisms. Most previously published
work that discussed cell lysis in microalgae has focussed on the extraction of lipids
and to a lesser extent, nucleic acids. Physical and chemical extraction methods are
used for cell lysis. However, chemical/solvent methods are not compatible with
lipases (and other enzymes) as protein denaturation and/or inhibition can occur in the
presence of solvents. Lee et al. (1998) reported on physical methods including bead-
beating, sonication, homogenisation, freeze-thaw cycling and French pressure cells.
A later study from the same laboratory used microwave energy, however the high
temperatures generated by this method meant it was not suitable for protein isolation
(Lee et al. 2010). Bead-beating appeared to be a relatively effective method of
achieving high levels of cell breakage (Lee et al. 2010; Lee et al. 1998). It has also
been noted that this method generally causes less damage to biologically active
compounds, such as proteins, than other physical methods (Chisti & Mooyoung
1986). This method is also easily established in the laboratory with minimal outlay of
resources required. Bead-beating was trialled on a small scale (1-2 mL algal cell
pellet), as outlined in Section 2.1.5.1. The method gave satisfactory cell lysis when
cell preparations were observed by microscopy before and after bead-beating (Figure
4.2.1).
Figure 4.2.1 P. lutheri cells before and after bead-beating. Visualised under 100 x magnification.
50 m 50 m
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84
The amount of lipase activity present in the crude homogenate was relatively low. To
obtain sufficient material for research to progress, larger amounts of biomass needed
to be processed. Due to the physical dimensions of test tubes, when the volume of
both the algal biomass and glass beads was increased, less cell breakage and lower
protein yields were achieved. This appeared to result from poor movement of the
beads in the bottom of the tube, which ultimately meant less interaction between the
cells and beads. Subsequently, it was decided that modifications to the protocol, as
well as alternative methods, should be investigated that could enable more biomass
to be processed. The glass vessel used for bead-beating was changed to allow more
material to be processed. Not only was the volume changed but also the shape.
Instead of a 100 mL standard test-tube, a 100 mL ‘pear shaped’ round bottomed flask
was tested. It was observed that while vortex mixing in the flask, movement/mixing
of the glass beads in solution was improved, presumably increasing interaction with
the cells. This was confirmed when identical volumes (10 mL algal pellet in 30 mL
extraction buffer) of algal cells were bead-beaten in both a 100 mL test tube and 100
mL flask. There was more cell breakage accomplished using the latter (data not
shown).
Homogenisation using an Ultra-Turrax was investigated due to its suitability for
processing relatively large volumes of biomass (see Section 2.1.5.2). An Ultra-
Turrax generates shear forces that are strong enough to disrupt the cell
wall/membrane (Chisti et al. 1986). The protein homogenates obtained from
homogenisation and bead-beating (in a flask) were analysed for protein concentration
by the method of Lowry (Section 2.1.7) and for lipase activity using pNP assays
(Table 4.2.2).
Table 4.2.2 Lipase activity and protein concentration of homogenate obtained via
homogenisation and bead-beating of P. lutheri cells.
Method Protein conc. mg/mL
Activity on pNPP U/mL
Specific activity U/mg
Homogenisation 2.04 0.012 0.005
Bead-beating 7.55 0.157 0.020
Data are averages of duplicate measurements
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85
Bead-beating was found to deliver superior protein yield (mg/mL) at more than three
times that obtained via homogenisation. The specific enzyme activity obtained with
bead-beating was also higher. This was surprising as homogenisation has been
shown previously not to influence enzyme activity (Chisti et al. 1986). It is possible
that if the lipases were particularly sensitive to increases in temperature, the high
shear forces and/or localised heat generated from homogenisation may have caused
some protein denaturation, resulting in lower activity. As a result of these findings
bead-beating was established as the most appropriate method available for accessing
intracellular protein from the microalgae.
4.3 Calcium-dependence of lipase activity
Lipase activity from the crude extracts tested with the pNP assay was found to be
dependent on the presence of CaCl2. The initial buffer for the pNP assay contained
20 mM Tris-HCl pH 8.0 and 0.01% gum arabic. This gave no measureable activity.
The second pNP buffer contained 20 mM Tris-HCl pH 8.0, 20 mM CaCl2, 5 mM
sodium cholate and 0.01% gum arabic. This assay set-up had been optimised at Plant
and Food Research for use with lipases isolated from the caeca of salmon and hoki
(Kurtovic et al. 2010). When the crude algal protein homogenate was tested with this
buffer, pNPP hydrolysis was observed. Not all lipases require calcium and/or bile
salts for activity, particularly those from microbial sources (Knoche & Horner 1970;
Patkar et al. 1993; Yu et al. 2007). The lipases that require bile salts for optimal
activity are usually sourced from the digestive tract of multicellular organisms
(Kurtovic et al. 2009). To investigate these cofactor dependencies in the microalgal
lipases, pNP buffers were tested in the presence and absence of CaCl2 and sodium
cholate. The activity observed with different buffers for each microalgal species is
summarised in Table 4.3. When sodium cholate was excluded from the buffer no
significant change was observed in the activity. However, when CaCl2 was removed
from the buffer, the hydrolysis of pNPP was almost completely abolished. These
findings showed that lipase activity in both species was dependent on calcium.
Calcium-dependence or stimulation of lipase activity has been observed with a
number of lipases from other sources. Pancreatic lipases have been shown to require
calcium for optimal activity (Bourne et al. 1994; Vantilbeurgh et al. 1993). Notably,
many plant lipases are also most active in the presence of calcium (Abigor et al.
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86
2002; Enujiugha et al. 2004; Hills & Beevers 1987; Piazza et al. 1989). Similarities
in the enzymatic requirements of higher plants and microalgae possibly demonstrate
the evolutionary closeness of these organisms.
Table 4.3 Lipase activity against pNPP from microalgal protein homogenates with different
assay buffers.
Species Buffer components
Activity* CaCl2 Sodium cholate
P. lutheri + + +
P. lutheri + - +
P. lutheri - + -
P. lutheri - - -
Isochrysis sp. + + +
Isochrysis sp. + - +
Isochrysis sp. - + -
Isochrysis sp. - - -
*Assay value required for a + result ( A410/min >0.01)
4.4 Storage of algal protein homogenates
The nature of this research project required algal protein samples to be processed and
transported from Plant and Food Research in New Zealand to Deakin University,
Australia. This was required as all algal biomass for this work was obtained from
The Cawthron Institute, Nelson, New Zealand, where microalgal culture facilities are
well established. A permit was acquired from the Australian Quarantine and
Inspection Service (AQIS) for the import of algal protein homogenates.
Freeze-drying was considered the most suitable method for storage and transport of
the protein homogenates, due to the reduced volume and weight of samples. It also
greatly reduces the chance of any viable cells remaining (Cordero & Voltolina 1997).
It was important that consistent enzyme activity was maintained in the samples once
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rehydrated. P. lutheri crude homogenates were used to trial this process. Duplicate
protein samples were prepared with and without 5% glycerol (Section 2.1.6). After
freeze-drying, the sample without glycerol weighed 440 mg and was in a dry flaky
biscuit-like form. The sample with 5% glycerol weighed 1014 mg, resulting in a
highly viscous gum-like material. The difference in weight was largely due to the
weight of the glycerol (500 mg) that was not removed by freeze-drying. However,
glycerol also aided in the retention of some water, with an additional 74 mg
unaccounted for. Once both samples had been resuspended with MQ H2O to the
appropriate weight (10 g), 100 μL of each sample was assayed for activity against
pNPP. The same samples that had not been freeze-dried were also assayed to
compare activity before and after freeze-drying (Figure 4.4). The activity observed
for the freeze-dried sample without glycerol decreased slightly more (~10%) than
that of the sample with 5% glycerol (~5%). However, the highly viscous product that
resulted from the addition of glycerol was both difficult to handle and transfer
between storage vessels. It was decided that despite the loss of slightly more activity,
the dried form of the sample without glycerol was more suitable for the storage and
transport of material. It also reduced issues that may have arisen with AQIS and
concerns regarding the presence of viable microalgae cells in the ‘less dry’ glycerol
samples.
Figure 4.4 Relative activities against pNPP for P. lutheri protein homogenate before and after
freeze-drying. ( ) Non-freeze-dried sample, ( ) freeze-dried sample. Measurements are averages of
duplicates.
75
80
85
90
95
100
sample sample with %5glycerol
Rela
tive
activ
ity a
gain
st p
NPP
(%)
sample with 5% glycerol
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88
4.5 Lipase purification by precipitation and ion-exchange
chromatography
Isolation of the lipases was required to obtain accurate biochemical characterisation
data. A number of methods were trialled to obtain purified or partially purified lipase
from the crude protein homogenates. The methods trialled in the work presented here
are not exhaustive but are methods commonly used in the purification of enzymes
and were available in the laboratory at the time of experimentation. They include
ammonium sulfate precipitation and ion-exchange chromatography. Hydrophobic
interaction chromatography was also tested and is discussed in Section 4.6.
4.5.1 Ammonium sulfate precipitation
Ammonium sulfate precipitation is a technique that works by altering protein
solubility and is the most common precipitation method undertaken in the isolation
of microbial lipases (Saxena et al. 2003). Ammonium sulfate precipitation was the
first method investigated in the current study for the concentration of lipases from
protein homogenates (see Section 2.1.15). Fractions obtained at differing saturation
levels (10, 20, 40 and 60%) were analysed by pNP activity and Lowry protein
concentration assays. As an example, the activity and protein concentration data
obtained for Isochrysis sp. is shown in Table 4.5.1. Specific activity (U/mg) was
calculated from these data and indicates if the protein had been enriched.
The greatest protein concentration and activity was obtained in the 40% fraction.
However, the 40% fraction had the lowest specific activity (activity/mg protein),
indicating that the lipase was not concentrated. The 10, 20 and 60%+ fractions gave
very small increases in specific activity. However, the spreading of lipase activity
throughout the fractions meant that considerable losses occurred and showed that this
method did not effectively concentrate lipase. With the low activity present in the
starting material further dilution of activity by the ammonium sulfate precipitation
was unhelpful. Similar results were found for the protein homogenate in P. lutheri,
with already low starting activity diluted further (data not shown). Therefore,
ammonium sulfate precipitation was rejected as a first step in concentrating lipase
activity in the microalgal samples.
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Table 4.5.1 Specific activity of ammonium sulfate precipitated fractions from Isochrysis sp.
protein homogenate.
Fraction Protein conc. mg/mL
Activity on pNPP U/mL
Specific activity U/mg
Start 10.79 0.025 0.0023
10% 1.156 0.0035 0.0030
20% 5.50 0.0138 0.0025
40% 12.61 0.0198 0.0016
60% 4.36 0.0082 0.0019
60%+ 1.09 0.0032 0.0029
Data are averages of duplicate measurements
4.5.2 Anion-exchange chromatography
Anion-exchange chromatography was used as a method for enrichment of lipases
from Isochrysis sp. and P. lutheri. Chromatography was performed with a BioLogic
DuoFlow (Bio-Rad, Hercules, CA, USA) FPLC fitted with a 1 mL HiTrapTM DEAE
FF column (GE Healthcare, Little Chalfont, UK) (Section 2.1.16). Algal protein
homogenate was dialysed against anion-exchange buffer A (20 mM Tris-HCl pH
8.0) prior to chromatography. Sample load was 1 mL and the chromatography was
run at a flow rate of 1 mL/min. An example of the chromatogram of protein
homogenate from P. lutheri is displayed in Figure 4.5.2. Protein was eluted as the
concentration of buffer B (20 mM Tris-HCl pH 8.0, 0.5 M NaCl) increased (Figure
4.5.2A). However, most of the starting material appears to have travelled through the
column in the flow through fraction. The activity of the starting material, flow-
through and fractions of interest were tested against pNP-myristate (pNPM) (Figure
4.5.2B). The starting material and flow-through had activity, with the flow-through
activity approximately 3-4 times lower. This is due to the dilution of protein in the
flow-through fraction (4 min collection time). All fractions of interest (matched to
chromatogram peaks) were found to have no significant activity.
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90
Figure 4.5.2 Anion-exchange chromatography of the protein homogenate from P. lutheri. (A)
The chromatogram of P. lutheri protein homogenate, green line represents the absorbance at 280 nm,
black line represents concentration of buffer B, red line indicates conductivity and light green bars
represent fractions of which the activity was investigated. (B) Activity ( A410/min) of starting material
(start), flow through (ft) and fractions against pNP-myristate. (C) Activity ( A410/min) of fractions as
in B, mixed with starting material (1:1) against pNP-myristate.
0
0.1
0.2
0.3
0.4
0.5
start ft 6 7 8 9 11 12 13 17 18 19
Activ
ity (
A 410
)
Fraction
00.05
0.10.15
0.20.25
0.30.35
start ft 6 7 8 9 11 12 13 17 18 19
Activ
ity (
A 410
)
Fraction
A
B
C
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91
We hypothesised that the lack of activity in the fractions may have been due to the
separation of a lipase co-factor present in the starting material. A second set of
activity assays were performed where the fraction material was mixed 1:1 with the
starting material. It was expected that if a co-factor was reintroduced, the activity in
fractions containing lipase would be restored. The addition of starting material to the
relevant fractions did not result in the activity of any particular fraction being
significantly higher (Figure 4.5.2C). This indicated that the activity seen was
associated with the added starting material. These assays, and the presence of activity
in the flow through, show that the lipase(s) were not bound by the anion-exchange
column.
The same experiments were performed for the protein homogenate from Isochrysis
sp. and can be found in Appendix 8.3, Figure 8.3.1. As was observed with P. lutheri,
protein was loaded onto the column and eluted as the concentration of buffer B
increased. Fractions relating to elution peaks on the chromatogram were then tested
for activity against pNPM. As shown in Figure 8.3.1B, very low levels of activity
were present across a range of fractions. Activity in the flow-through was again
approximately 3-4 times less than that of the starting material. No noticeable
increases in activity were observed in any of the fractions when the starting material
was added to them in an attempt to restore activity (Figure 8.3.1C). This indicated
that very little lipase from Isochrysis sp. was being eluted and that no co-factor had
been lost. SDS-PAGE was not performed on any of the fractions from either species,
as the lack of activity and known molecular weight meant it would not provide
further indication of the presence lipases in the fractions. These results demonstrated
that anion-exchange chromatography was not an appropriate method for the
purification of lipases from these microalgae.
A stronger anion-exchange column (1 mL HiTrapTM Q FF column (GE Healthcare,
Little Chalfont, UK)) was also tested. However, this was also unsuccessful in
enriching the lipases from these microalgae.
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4.5.3 Acidic native-PAGE analysis of P. lutheri extract
Based on the behaviour of anion-exchange chromatography it was hypothesised that
the lipases present in the crude protein homogenate may have basic (>7.0) isoelectric
points (pI). Therefore they would not be bound by the anion-exchange column (at pH
8.0 a protein with a pI >7.0 will likely carry either no charge or a positive charge and
will not be able to interact with the positively charged anion column support). The
presence of lipases with basic pI’s was examined in P. lutheri by acidic native-PAGE
(Section 2.1.13) with a MUB gel-overlay assays. Acidic native-PAGE enables the
separation of proteins with pI’s >7.0 (Reisfeld et al. 1962) that cannot be separated
by standard (basic) native-PAGE conditions. A single band is clearly visible in the
acidic native-PAGE (Figure 4.5.3), indicating the presence of a potential lipase with
a pI >7.0. The band did not migrate far into the gel, indicating that the proteins pI is
not strongly basic. After detecting activity in the acidic gel cation-exchange
chromatography was selected as a potentially suitable method for the purification of
lipase(s) from the microalgae.
Figure 4.5.3 4-Methylumbelliferyl butyrate gel-overlay assay on an acidic native-PAGE of P.
lutheri protein homogenate. (1) The acidic native-PAGE with 30 μl of protein sample loaded, (2) inset
of the basic native-PAGE from Figure 4.1 for comparison. White dashed line marks the beginning of
the resolving gel. Black arrows indicate observed fluorescence bands.
1 2
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4.5.4 Cation-exchange chromatography
Cation-exchange chromatography was performed using a BioLogic DuoFlow (Bio-
Rad, Hercules, CA, USA) FPLC fitted with a 1 mL HiTrapTM SP FF column (GE
Healthcare, Little Chalfont, UK) (Section 2.1.17). Algal protein homogenate was
dialysed against cation-exchange buffer A (20 mM MES pH 6.0) prior to
chromatography. An example chromatogram of the protein homogenate from P.
lutheri is shown in Figure 4.5.4. A very low level of protein was eluted as the
concentration of buffer B (20 mM MES pH 6.0, 0.5 M NaCl) increased (Figure
4.5.4A). The majority of the starting material travelled through the column in the
flow-through fraction. The activity of the starting material, flow-through and
fractions of interest were measured using the pNP assay (Figure 4.5.4B). The starting
material and flow-through displayed activity, with flow-through activity again being
3-4 times lower than that of the starting material. The range of fractions analysed
were found to have no significant activity. As with the anion-exchange experiments,
a second set of activity assays were performed with starting material mixed 1:1 with
the fraction material. However, the addition of starting material did not result in
activity being restored (Figure 4.5.4C), indicating no co-factor had been lost.
Protein homogenate from Isochrysis sp. was used in the same experiments, with
results shown in Appendix 8.3, Figure 8.3.2. Little protein was bound, or eluted by
the column as the concentration of buffer B increased. The general lack of
discernible peaks makes isolation difficult. Fractions relating to elusion peaks on the
chromatogram were then tested for activity against pNPM (Figure 8.3.2B). A range
of fractions were analysed and found to have little or no activity. Again no noticeable
increase in activity was observed in any fractions when starting material was added
(Figure 8.3.2C). These results demonstrated that cation-exchange was also not an
appropriate method for the purification of lipases from the protein homogenates of
these microalgal species.
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94
Figure 4.5.4 Cation-exchange chromatography of the protein homogenate from P. lutheri. (A)
The chromatogram of P. lutheri protein homogenate, green line represents the absorbance at 280 nm,
black line represents concentration of buffer B, red line indicates conductivity and light green bars
represent fractions of which the activity was investigated. (B) Activity ( A410/min) of starting material
(start), flow through (ft) and fractions of interest against pNP-myristate. (C) Activity ( A410/min) of
fractions as in B, mixed with starting material (1:1) against pNP-myristate.
0
0.1
0.2
0.3
0.4
0.5
start ft 6 7 8 9 10 11 14 16 18
Activ
ity (
A 410
)
Fraction
00.05
0.10.15
0.20.25
0.30.35
start ft 6 7 8 9 10 11 14 16 18
Activ
ity (
A 410
)
Fraction
A
B
C
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Ion-exchange chromatography has been used routinely for isolating lipase from a
variety of sources. However, non-specific methods, such as ion-exchange
chromatography, are known to result in significant losses of target protein yields
(Saxena et al. 2003; Taipa et al. 1992). The lipase activity present in the starting
material was low making purification challenging. Furthermore, given that the
purification of lipase to homogeneity in most protocols requires 4 to 5 steps, this
approach was simply not achievable with the low starting activity. A higher level of
activity in the starting material would be required for future enrichment studies using
ion-exchange chromatography. This could be achieved if the lipases-encoding genes
were cloned and expressed recombinantly.
4.6 Lipase purification by hydrophobic interaction
Lipases interact with insoluble lipid substrates at the lipid-water interface. The nature
of this interaction involves the lipase undergoing a conformational change whereby
hydrophobic surfaces surrounding the lipase active site interact with the lipid
interface (Fernandez-Lafuente et al. 1998). This is often accompanied by an
increased activity (hyper-activation) and is referred to as interfacial activation
(Fernandez-Lorente et al. 2007; Palomo et al. 2006). It is thought that this process
also occurs when lipases are adsorbed onto hydrophobic supports (Fernandez-
Lorente et al. 2008; Hanefeld et al. 2009). Binding of the lipase to a hydrophobic
support is usually carried out at low ionic strength, minimising non-specific binding
of unwanted proteins (Fernandez-Lafuente et al. 1998; Sabuquillo et al. 1998). This
property has been exploited extensively for the isolation and/or immobilisation of
lipases from a range of sources (Bastida et al. 1998; Kurtovic et al. 2011; Palomo et
al. 2002). In some instances hydrophobic interaction chromatography has been used
to purify lipases to homogeneity in a single step (Gupta et al. 2005).
Owing to the potential selectivity of this method, it was tested for purifying and/or
immobilising lipases from the microalgal samples. The potential hyper-activation of
lipase activity could also address the difficulties of analysing low amounts of lipase
activity. Finally, the use of hydrophobic supports has been shown to be useful in
separating multiple lipolytic enzymes in crude mixtures (Sabuquillo et al. 1998). As
multiple bands were observed in Figure 4.1, this method could potentially provide a
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96
means for separating different lipolytic enzymes in the crude algal homogenates and
warranted further investigation.
Hydrophobic interaction chromatography was tested as a method for the isolation of
lipase from P. lutheri. The hydrophobic supports used were butyl- and octyl-
Sepharose and butyl-Toyopearl resins (refer to Section 2.1.18). Two different
supports were trialled as lipases may interact with various supports differently and
this can alter both lipase binding and activity (Fernandez-Lorente et al. 2008;
Kurtovic et al. 2011). Sepharose is a carbohydrate-based resin that is hydrophilic in
nature, but when coated in butyl/octyl chains the resin becomes hydrophobic
(Fernandez-Lafuente et al. 1998). Toyopearl is an acrylic support that is hydrophobic
in nature and its hydrophobicity is enhanced when hydrophobic groups are added
(Fernandez-Lorente et al. 2008). Thus the two supports may interact differently with
the microalgal lipases. Hydrophobic adsorption was performed on algal protein
homogenates in low ionic strength conditions (Section 2.1.18). The supernatant was
monitored by pNP activity and Lowry protein concentration assays, to define the
specific activity (activity/mg protein) and thus determine the specificity of lipase
binding.
4.6.1 Hydrophobic interaction with Sepharose® resins
The less hydrophobic butyl-Sepharose resin was trialled first followed by octyl-
Sepharose (more hydrophobic), to test if the separation of different lipolytic enzymes
was possible using resins of different hydrophobicities. The starting material and
butyl-/octyl-Sepharose supernatants were analysed by pNP and Lowry assays. The
pNP assay was performed with both pNP-butyrate (pNPB) and pNP-myristate
(pNPM) so that both lipase and esterase activities could be monitored (Figure 4.6.1).
The highest activity was found in the starting material. Activity in the supernatant
dropped by approximately 50% after exposure to the butyl-Sepharose and then
further again after incubation with octyl-Sepharose (Figure 4.6.1A). This was
promising as it suggested fractionation of different lipolytic enzymes by the varying
degrees of support hydrophobicity. Furthermore, the uptake of activity onto the
supports was lipase specific (Figure 4.6.1B), as the specific activity (U/mg protein)
decreased after each step, indicating more lipase was adsorbed than non-target
proteins. Activity against both pNPB and pNPM decreased in the same fashion.
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These findings demonstrated that the lipase had been adsorbed onto the hydrophobic
supports.
To confirm enzyme adsorption the activity of the lipase bound to the Sepharose
resins had to be measured. Accurate measurement of lipase activity on hydrophobic
supports cannot be performed using the pNP assay. Although hydrolysis may occur,
the released pNP appears to bind to the supports, thereby removing it from the
spectrophotometrically measureable aqueous phase. The development of a yellow
colour on the resin indicated that activity was likely to be present, but it was not
quantifiable. Attempts to obtain quantitative data on the activity of hydrophobically
bound lipase were made using the titrimetric assay (see Section 2.1.9) with a
tributyrin substrate. No measurable activity was observed against tributyrin with
either butyl- or octyl-Sepharose preparations. A large aliquot (2 mL) of starting
material (protein homogenate) was also assayed and found to have low activity (data
not shown). The large volume ( 25 mL) of the titrimetric assay means that a
significant amount of lipase is required to achieve measurable hydrolysis rates.
Although lipase was bound by the Sepharose resins, there was not enough to enable
determination of its presence by this assay method.
Figure 4.6.1 Relative and specific lipase activities of the Sepharose adsorption fractions against
pNP-acyl esters. (A) Activity of all fractions relative to that of the starting material against pNPB. (B)
Specific activity (activity/mg protein) of all samples. pNPB ( ), pNPM ( ). Data are averages of
duplicates.
0
20
40
60
80
100
120
start post butyl post octyl
Rela
tive
activ
ity (%
)
Sample/fraction
0
0.005
0.01
0.015
0.02
0.025
0.03
0.035
0.04
start post butyl post octyl
spec
ific a
ctiv
ity (U
/mg
prot
ein)
Sample/fraction
120
0 035
0.04
n)
A B
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98
Elution of the bound material from the Sepharose resins was also investigated as a
means to enrich the lipase. This has been achieved with low concentrations (<1%,
v/v) of Triton X-100 in other studies (Fernandez-Lorente et al. 2008). The washed
butyl- and octyl-Sepharose resins were incubated in 5 mM Tris-HCl pH 7.4
containing Triton X-100 at concentrations of 0.5, 1.0 and 2.0% (v/v). The eluted
samples were then analysed for activity by pNP assay. No measurable activity was
observed (data not shown). This indicated that either: (1) The lipase was bound
tightly to the support and not eluted; or (2) that Triton X-100 had destroyed the
activity of the lipase. Triton X-100 can be detrimental to the activity of lipases from
some sources (Diaz et al. 2007; Gargouri et al. 1983; Weselake et al. 1989).
4.6.2 Hydrophobic interaction with Toyopearl resin
The second trial used butyl-Toyopearl resin as a hydrophobic support. The same
protocol was followed as described for the Sepharose resins (Section 4.6.1). The
supernatant and resin were separated and retained for analysis. Samples were
analysed by pNP activity (pNPB and pNPM substrates) and Lowry concentration
assays (Figure 4.6.2). Activity in the supernatant fraction after incubation with butyl-
Toyopearl was less than 20% of the starting material (Figure 4.6.2A). This showed
that the butyl-Toyopearl bound the activity more readily than the butyl-Sepharose
resin. When specific activity (Figure 4.6.2B) was analysed, it showed that the uptake
of activity onto the support was lipase specific. The activity of the supernatant
against the two substrates differed, with the specific hydrolysis against pNPM
decreasing more than for pNPB. This may indicate that lipase was bound more
readily than other lipolytic enzyme species in the homogenate (such as esterases
which cleave shorter chain substrates). As was found with the Sepharose resins,
measurement of ‘bound lipase’ activity was not achieved by titrimetric methods.
Elution of lipase from butyl-Toyopearl by Triton X-100 treatment also could not be
detected (data not shown).
Hydrophobic supports appear to be able to adsorb lipases from a crude homogenate
in a lipase specific manner. However, to achieve binding of measurable amounts of
activity in the future, a different approach would be required to obtain more activity
in the starting material.
CHAPTER IV
99
Figure 4.6.2 Relative and specific lipase activities of the fractions from Toyopearl adsorption
against pNP-acyl esters. (A) Activity of all fractions relative to that of the starting material against
pNPB. (B) Specific activity (activity/mg protein) of all samples. pNPB ( ), pNPM ( ). Data are
averages of duplicate readings.
The use of molecular biological methods is one such approach. This project aimed to
isolate these enzymes via classical biochemical methods (i.e. native isolation). This
is often difficult, as has been demonstrated by the current study. The use of
recombinant systems for the expression of these lipases offers an alternative
approach. The ability to use not only bacteria and yeast as microbial expression
hosts, but microalgae themselves, has improved as understanding of model
organisms has progressed in recent years (Specht et al. 2010). Examples of
recombinant expression approaches, and the benefits they provide, can be seen in
recent work by French researchers Godet et al. (2010) and Kerviel et al. (2014),
where they have isolated and analysed sequences, and recombinantly expressed lipid
hydrolases from Isochrysis galbana. Comprehensive biochemical characterisation of
these enzymes has not yet been described.
0
20
40
60
80
100
120
start post butyl
Rela
tive
activ
ity (%
)
Sample/fraction
0
0.005
0.01
0.015
0.02
0.025
0.03
start post butyl
spec
ific a
ctiv
ity (U
/mg
prot
ein)
Sample/fraction
120 0.03
in)
A B
CHAPTER IV
100
4.7 Enzymatic analysis of lipase-containing crude material
Despite being unable to purify lipases from the microalgal protein homogenate we
decided to undertake biochemical characterisation of activity in the crude material to
see if anything could be learned about the lipase specificity/behaviour without
purification. This could give an indication of whether future efforts to isolate
these lipases via molecular biological means would be worthwhile (i.e. if the
enzymes had useful properties). The molecular weight of the lipase from P. lutheri
was investigated, followed by the effects of temperature and pH on hydrolysis rate to
determine optima. The FA selectivity of lipases in the crude material was also
analysed.
4.7.1 Molecular weight analysis by multi-PAGE and mass spectrometry
An approach using multiple PAGEs and matrix-assisted laser desorption/ionisation
time-of-flight/time-of-flight (MALDI TOF/TOF) mass spectrometry was used to
elucidate the molecular weight of lipase in P. lutheri. Images of the fluorescent bands
observed by MUB gel-overlay assays on acidic and basic native-PAGEs were
recorded (refer to Appendix 8.4, Figure 8.4). Subsequently, both gels were stained
with Coomassie blue and imaged again. The gels were then aligned and Coomassie
bands cut out (with a scalpel) from the area corresponding to the fluorescent bands
(band 1 from acidic native-PAGE and bands 1 and 2 from basic native-PAGE, see
Figure 8.4). The excised gel bands were then suspended in SDS-PAGE loading
buffer and prepared for electrophoresis (Section 2.1.11). Starting material and the gel
bands were loaded into the appropriate wells. After electrophoresis, the gel was
silver stained as there was not enough material present to visualise bands by
Coomassie staining. The silver stained gel showed that a number of protein bands
were present in each of the gel bands cut from the native-PAGE gels (Figure 4.7.1).
Bands from lane 3 (3A and 3B) and lane 4 (4A, 4B and 4C), correspond to bands 1
and 2 from the basic native-PAGE, respectively. As numerous tightly-stacked bands
were present in lane 7 (band 1 from the acidic native-PAGE), analysis was not
feasible. Bands were selected on the basis of approximate molecular weight (i.e.
range of 20 to 90 kDa) band intensity and adequate separation (i.e. a clear band could
be seen).
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101
Figure 4.7.1 SDS-PAGE of fluorescent bands cut from native-PAGE of P. lutheri protein
homogenate. Lane 1; Molecular weight marker (4 μl), lane 2; starting material (20 μl load), lane 3; gel
band 1 from basic native-PAGE, lane 4; gel band 2 from basic native-PAGE, lane 5; no sample, lane
6; starting material (20 μl load), lane 7; gel band 1 from acidic native-PAGE. Bands denoted 3A, 3B,
4A, 4B and 4C were cut out and sent for analysis by mass spectrometry.
Protein bands of interest were then cut out again and sent to the Centre for Protein
Research at the University of Otago, New Zealand, for analysis by MALDI
TOF/TOF mass spectrometry. Proteins were identified by comparing the measured
collision induced dissociation pattern of tryptic peptides (trypsin digested) with the
calculated fragmentation pattern of tryptic peptide digestion, as predicted in silico by
the Mascot database (Matrix Science, Boston, MA,USA).
The bands 3A and 3B in lane 3 (Figure 4.7.1) gave no matches for lipase sequences.
Band 3A gave the highest score for an S-adenosyl-L-homocysteine hydrolase from
Pavlova lutheri. However, this band also provided numerous hits for ribulose-1,5-
bisphosphate carboxylase/oxygenase (RuBisCO) (see Appendix 8.5). The presence
of RuBisCO in the band demonstrates that the band was not clean. This was not
3A
3B
4A
4B
4C
200 - 116 -
97 -
66 -
45 -
31 -
21 -
14 -
6.5 -
CHAPTER IV
102
completely unexpected, as RuBisCO is the most common protein on earth (Raven
2013) and is often a contaminant in plant based samples. Lane 3B gave a single hit
for a cytosolic aldolase from Fragaria x ananassa (garden strawberry) (see
Appendix 8.5). Bands from lane 4 were found to be too dilute for analysis by
MALDI TOF/TOF mass spectrometry. They were subsequently analysed by LTQ-
Orbitrap hybrid mass spectrometry, which offers increased sensitivity. However,
while numerous hits were obtained none were for lipase. The main sequence matches
were for trypsin and keratin (types I and II cytoskeletal) from mammalian sources
(data not shown). This suggested the main peptide species present arose from sample
contamination, most likely from gel handling post electrophoresis. The findings from
this work indicate that the protein homogenate must be at least partially purified to
remove potential sources of contamination, before molecular weight analyses are
performed.
4.7.2 Effect of temperature on lipase hydrolysis
Temperature optima characterisation of the crude/mixed lipases from Isochrysis sp.
and P. lutheri was carried out using the pNP assay. The substrate pNPM was used in
assays ranging from 20 to 60 C. Protein extracts from P. lutheri and Isochrysis sp.
gave optimal hydrolysis of pNPM at 30 C and 55 C, respectively (Figure 4.7.2).
There was a clear difference observed in temperature optima between the two
microalgal species. The P. lutheri lipase was most active under lower temperatures,
as would be expected for an organism living in an aquatic environment. Surprisingly
Isochrysis sp. displayed a notably higher optimal temperature. It is unclear why a
marine microalga would possess an enzyme that is optimally active at temperatures
far exceeding those in which the organism lives. However, this property could be
useful in commercial settings as most industrial processes using lipases are carried
out above 45 C (Sharma et al. 2002). Comparison of these findings with other
microalgae is not yet possible as no lipases have been characterised. The temperature
optima of lipases sourced from other organisms varies greatly, as summarised by
Ghosh et al. (1996). There are examples of lipases isolated from various
psychrotrophiles (Rashid et al. 2001) and thermophiles (Li & Zhang 2005) that
exhibit optimal activities representative of the organisms origins. However, in many
instances there is no clear relationship between optimal activity and the environment.
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103
Thermostability could not be explored using the crude material, as significant protein
precipitation meant the samples became unmeasurable.
These findings help to explain the lower activity observed from Isochrysis sp.
compared to P. lutheri in earlier experiments, where standard assays were carried out
at 30 C. Measurement of lipase activity at high temperatures has numerous
challenges. While it can be achieved using spectrophotometric methods as long as
the non-enzymatic activity is carefully monitored, titration methods can become
particularly difficult. For example, titration of tributyrin hydrolysis at high
temperatures causes condensation (containing butyric acid) inside the reaction vessel,
which subsequently drips back into solution causing instability in measured pH.
Figure 4.7.2 Effect of temperature on the activity of protein homogenates from Isochrysis sp. and
P. lutheri. The relative activity of each crude homogenates against pNPM was normalised to the
optimal activity obtained for each sample; 30 C for P. lutheri ( ) and 55 C for Isochrysis sp. ( ).
0
20
40
60
80
100
120
0 20 40 60 80
Rela
tive
activ
ity (%
)
Temperature ( C)
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104
4.7.3 Effect of pH on lipase hydrolysis
Investigating the pH optima of lipases from Isochrysis sp. and P. lutheri was less
straightforward than for temperature. The pNP assay was used initially, with pNPM
as the substrate. This assay has limitations when investigating acidic conditions, as
the absorbance of pNP cannot be measured effectively below pH 6.0 (see Section
1.5). The extinction coefficients of pNP were determined for each pH value assayed,
before the measurement of lipase activity (see Appendix 8.6, Table 8.6). These
values are very similar to those previously reported by Kurtovic et al. (2010). Protein
extracts from P. lutheri and Isochrysis sp. were assayed at pH 6.0-9.0 (Figure 4.7.3).
Optimal hydrolysis of pNPM was carried out by P. lutheri and Isochrysis sp. at pH
6.0 and 7.0, respectively. A modified pNP assay was tested in an attempt to gather
measurements below pH 6.0. The assay was performed at the desired pH before
being adjusted to pH 8.0 for measurement. However, results were inconsistent (data
not shown).
Figure 4.7.3 Effect of pH on the activity of protein homogenates from Isochrysis sp. and P.
lutheri. Plot contains data for the hydrolysis of both pNPM and MUB. The relative activity of each
crude homogenate was normalised to the optimal activity obtained for each set of assays. Assays were
performed at 30 C for P. lutheri and 45 C for Isochrysis sp.. P. lutheri against MUB ( ) and
pNPM ( ), Isochrysis sp. against MUB ( ) and pNPM ( ).
0
20
40
60
80
100
120
3 4 5 6 7 8 9
Rela
tive
activ
ity (%
)
pH
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105
Fluorescence assay methods were investigated to measure lipase activity under acidic
conditions. Work by Roberts (1985) indicated that this was possible using 4-
methylumbelliferone derivatives as substrates. Activity profiles were obtained for the
hydrolysis of MUB at pH 3.0-8.0. Overlaying these data points on the same graph as
those from the pNP assays gave an estimation of pH optima (Figure 4.7.3). The
optima obtained for the lipase present in P. lutheri was pH 5.0-6.0, while for
Isochrysis sp. it was approximately pH 7.0. The lipase from P. lutheri had a broad
range of activity with >70% of the optimum activity performed between pH 4.0 and
7.0. Isochrysis sp. activity appeared more restricted and gave a sharper pH optimum
profile. Again, there are no previous studies defining the pH optima of lipase from
microalgae for comparison. The pH optima obtained for the lipases from these
microalgae was relatively neutral, with P. lutheri tending towards more acidic
optima. This is unsurprising as the majority of lipases characterised from eukaryotic
microbial sources appear to possess relatively neutral pH optima (Ghosh et al. 1996;
Vakhlu & Kour 2006). In contrast, numerous bacterial lipases show more extreme
pH optima, many of which are optimally active at alkaline pH (Gupta et al. 2004).
4.7.4 Fatty acid selectivity of algal homogenates
Defining general fatty acid (FA) selectivity for the crude lipase mixtures was
challenging due to the low level of activity present in the microalgal protein samples.
Conventional methods for investigating lipase FA selectivity involve the direct
hydrolysis of oil or oil emulsions (outlined in Sections 1.4-1.4.3). These methods
could not be used with the low level of activity present in the microalgal samples.
Colorimetric methods offer increased sensitivity and the pNP assay method was
already established in our laboratory. However, pNP substrates are limited to those
that are commercially available (i.e. saturated acyl groups only). We hypothesised
that if a range of unsaturated acyl group derivatives were synthesised to complement
the commercial substrates, this assay platform could be further utilised to screen FA
selectivity (see Chapter V). The extended pNP-acyl ester library was applied to the
protein homogenates from Isochrysis sp. and P. lutheri to produce lipase FA profiles
(Figure 4.7.4).
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106
Figure 4.7.4 Activity profiles of protein homogenates from Isochrysis sp. and P. lutheri against a
range of pNP-acyl esters. The relative activity of each substrate is normalised to the highest hydrolysis
value, as 100%. (A) Isochrysis sp. and (B) P. lutheri. Substrates used in screen were pNP-acetate
(C2:0), pNP-butyrate (C4:0), pNP-octanoate (C8:0), pNP-dodecanoate (C12:0), pNP-myristate
(C14:0), pNP-palmitate (C16:0), pNP-palmitoleate (C16:1), pNP-stearate (C18:0), pNP-oleate
(C18:1), pNP-linoleoate (C18:2), pNP-linolenoate (C18:3), pNP-eicosapentaenoate (C20:5) and pNP-
docosahexaenoate (C22:6).
Optimal hydrolysis was achieved with substrates that were C4 or shorter. These
substrates are soluble under the conditions of the assay. It is possible that the activity
is a result of hydrolysis by other lipid-hydrolases, such as esterases as lipases usually
hydrolyse longer insoluble substrates more readily (Gupta et al. 2003). With
substrates ≤ C4 omitted from the profiles both homogenates showed optimal
0
20
40
60
80
100
120
Rela
tive
activ
ity (%
)
pNP-acyl ester
0
20
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Rela
tive
activ
ity (%
)
pNP-acyl ester
A
B
CHAPTER IV
107
hydrolysis against pNPM (C14:0). Saturated acyl groups shorter and longer than this
were hydrolysed less readily. The hydrolysis of pNP-acyl esters with olefinic acyl
groups occurred at levels approximately 70-80% of C14:0, indicating that the FA
specificity was relatively broad. This was particularly surprising with regard to the
hydrolysis of EPA and DHA, as most lipases hydrolyse these FA poorly (Mukherjee
et al. 1993). These profiles challenge the suggestion that lipases from microalgae
(that contain large quantities of LC n-3 PUFAs) may have evolved not to hydrolyse
these FAs, due to their metabolically expensive synthesis, as outlined in Section 3.4.
However, it is important to note that the pNP assay uses mono-FA substrates and
does not allow for analysis of positional selectivity (regio-selectivity). Lipase
positional selectivity could be particularly important in microalgae if TAG is used
primarily as a precursor for the synthesis of various phospholipids and glycolipids, as
hypothesised by Khozin-Goldberg et al. (2011). Still, these results suggest that the
lipases from these organisms may carry out hydrolysis on a relatively broad range of
FAs.
4.8 Summary
Microalgae have the ability to synthesise a vast array of lipids, including EPA and
DHA, suggesting they have lipases/lipid synthases with useful specificities. Lipase
was detected in both Isochrysis sp. and P. lutheri by the pNP activity and MUB gel-
overlay assays, with the latter demonstrating there was likely more than one lipase
present. The highest degree of algal cell lysis was achieved by the bead-beating
method. Lipase activity from both species was found to be calcium dependent.
Attempts at purification and enrichment of lipase from crude protein homogenate by
ammonium sulfate precipitation and ion-exchange chromatography were
unsuccessful. This was largely due to the low amount of activity in the crude
homogenate. The lipase from P. lutheri was adsorbed onto both Sepharose and
Toyopearl resins via hydrophobic interaction. However, efforts to measure activity
on the resins, or removal of lipase from the resin were not successful due to the low
amount of bound enzyme. Elucidation of the molecular weight of lipase present in P.
lutheri via native-PAGE and MALDI TOF/TOF mass spectrometry was not
achieved, with most sequence matches arising from non-target species or
contaminants.
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Activity optima were determined for the lipases in crude protein homogenates of
both Isochrysis sp. and P. lutheri. Temperature optima for lipase(s) in Isochrysis sp.
and P. lutheri extracts were 55 C and 30 C, respectively. The optimal pH for activity
was approximately 7.0 and 5.0-6.0 for lipases from Isochrysis sp. and P. lutheri,
respectively. The FA selectivity of the lipases from both species was similar. Short
chain substrates (≤ C4) were hydrolysed most readily, although this may have been
due to the presence of esterases in the crude homogenate. Optimal hydrolysis of
longer-chain insoluble substrates was seen against pNPM (C14:0). Activity against
longer unsaturated substrates was quite uniform, suggesting these enzymes are active
against a relatively broad array of FAs.
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CHAPTER V
Novel omega-3 pNP-acyl esters for the characterisation of
lipase fatty acid selectivity
5.0 Introduction
Lipases are used extensively as biocatalysts in many commercial and industrial
applications, such as pharmaceuticals, cosmetics, detergents, oil and biodiesel
processing (Gotor-Fernandez et al. 2006). A current application for lipases is the
processing and concentration of LC n-3 PUFAs for use in nutraceuticals,
pharmaceuticals and functional foods. Conventional processing methods for omega-3
oils often entail harsh processing conditions with respect to both temperature and pH.
Given the unstable nature of EPA and DHA, this can lead to degradative oxidation,
polymerisation and geometric isomerisation (Shahidi et al. 1998). As previously
mentioned, the use of lipases and the associated milder processing conditions
provides a number of advantages over chemical methods (Section 1.3.4). The
characterisation of lipase activity is important for deducing the appropriate
application of lipases. The assignment of enzyme activity via structural or sequence
based approaches is limited by predictive chemical theories. Therefore, practical
methods are still the best approach for accurately establishing enzyme activity
(Reymond et al. 2002).
Established methods for the screening and characterisation of novel lipases for LC n-
3 PUFA selectivity, as well as the modification of already known lipases for this
purpose are greatly limited by the time required to perform the methods. The use of
spectrophotometric assay systems and synthetic substrates, using a suitable
chromophore or fluorophore provide an alternative. These methods can be used for
determining the catalytic activity and relative FA selectivity of hydrolytic enzymes.
These methods are well established and widely used (Roberts 1985; Schmidinger et
al. 2005; Wahler et al. 2002a; Wahler & Jean-Louis 2002b). Such methodologies are
relatively simple and can be carried out in a high-throughput manner.
The coupling of an acyl group to the chromophore p-nitrophenol (pNP) is an
example of a substrate type that is utilised in the detection and screening of activity
CHAPTER V
110
in enzymes such as lipases. The hydrolysis of the ester bond is homologous to the
reaction these enzymes catalyse in vivo. The assay is based on the release of pNP and
its colorimetric detection at 400-410 nm as a measurement of enzymatic activity
(Winkler et al. 1979). When utilising a range of substrates with various acyl groups
the assay can be used both as a quantitative and qualitative measure of activity. This
is apparent when substrates are assembled into libraries and provides a powerful
analytical tool for characterising enzyme activity (Goddard & Reymond 2004;
Reymond et al. 2002). The pNP assay is particularly useful in defining acyl-chain-
length specificity in lipases, of which examples can be found in the literature
(Abramic et al. 1999; Brundiek et al. 2012; Couto et al. 2010; Kurtovic et al. 2010;
Qian et al. 2011; Yang et al. 2002).
While pNP has been used with numerous substrates in a variety of enzyme assays
(Liggieri et al. 2009; Nanjo et al. 1988; Nourooz-Zadeh et al. 1992; Qian et al. 2011),
its use has never included LC n-3 PUFA substrates. Current lipase applicable
substrates are limited primarily to saturate or mono-/di-unsaturate acyl groups
(Acharya et al. 2002; Nourooz-Zadeh et al. 1992). The pNP-acyl ester platform could
provide a simple and rapid method for identifying lipases suitable for omega-3
concentration. This work aimed to develop/establish a method for synthesising
olefinic pNP-acyl esters, followed by chemical characterisation of the new
compounds. The stability of these compounds was then monitored before the pNP-
acyl esters were incorporated into the assay for examining the FA selectivity of
lipases. The findings demonstrate the utility of the method when used for defining
lipase FA selectivity.
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5.1 pNP-acyl ester synthesis protocol development and
optimisation
The synthesis of pNP-acyl esters (described in Section 2.1.19) was first trialled with
the saturated FAs octanoic and stearic acid. The protocol was subsequently modified
to gain the best yields possible. The protocol was designed with the FA component
being the limiting reagent and all other components present in excess. The initial
method included an H2O washing step, whereby the reaction mixture (post reaction)
was added to 100 mL of chloroform and 100 mL of MQ H2O in a separating funnel.
The solution was mixed and the phases left to separate; in the instances where
separation was poor, 2 mL of saturated NaCl solution was added to aid in phase
separation (Barthet et al. 2002). The solvent phase was decanted and magnesium
sulphate added as a drying agent to remove any residual water. The solvent was then
removed by evaporation under reduced pressure before the sample underwent
column chromatography. It was thought that this step would aid in the removal of the
water soluble EDCI and its urea derivative ((N-ethyl)-N’-(3-dimethylaminopropyl)
urea) that is produced in the reaction. While this synthesis worked, the additional
processing steps resulted in a loss of product, giving low yields. Subsequently, it was
found that silica gel column chromatography (Section 2.1.21) alone was sufficient
for the removal of EDCI and its respective reaction products from the desired pNP-
acyl ester compounds; rendering the washing step unnecessary. Furthermore, when
this process was used for reactions involving LC n-3 PUFAs such as EPA and DHA,
problems such as oxidation must be taken into consideration. The exposure of such
compounds to an aqueous oxygen-rich environment was thus undesirable.
Altering the reaction temperature was also briefly investigated as a means of
increasing yield. However, no significant increase in yield was observed when the
reaction was performed at temperatures up to 55 C instead of room temperature (data
not shown). Despite the relatively poor yields (<50%) it was possible to obtain
enough material for the characterisation of the saturated pNP-acyl esters via 13C 1H
NMR. These NMR spectra together with TLC analysis confirmed that the protocol
was successfully linking the pNP with the FA (data not shown). The developed
protocol was used for the synthesis of the new olefinic compounds.
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5.2 Synthesis of novel olefinic pNP-acyl esters
Acyl groups were chosen for esterification based on those olefinic FAs most
prevalent in fish oils such as tuna and anchovy, which are currently the main source
of commercial omega-3 oils. These were palmitoleic acid (C16:1 n-7), oleic acid
(OA, C18:1 n-9), linoleic acid (LA, C18:2 n-6), -linolenic acid (ALA, C18:3 n-3),
eicosapentaenoic acid (EPA, C20:5 n-3) and docosahexaenoic acid (DHA, C22:6 n-
3). The synthesis protocol used for each olefinic pNP-acyl ester was initially carried
out in accordance with that discussed in Section 5.1, with yields ranging from 40 to
70% (data not shown). It was necessary to perform the synthesis of these compounds
in an inert nitrogen atmosphere and with minimal light exposure. Failure to do so,
particularly with EPA and DHA, resulted in a range of reaction products observed as
a smear by TLC (data not shown). The addition of 4-dimethylaminopyridine
(DMAP), as used previously by Qian et al. (2011) in pNP ester synthesis, was also
important and led to significantly increased yields.
DMAP was first used as a catalyst for the esterification of carboxylic acids by Neises
et al. (1978) and subsequently was called the ‘Steglich esterification’. Neises et al.
(1978) found that addition of 3-10% molar equivalent of DMAP to these
esterification reactions significantly increased yields. The Steglich esterification
involves the use of a carbodiimide to form a good leaving group on the carboxylic
acid. The catalytic role of DMAP appears to involve it reacting nucleophilically with
the carbodiimide activated ester (O-acylisourea in the case of EDCI) further
activating the esters as a reactive amide. DMAP therefore acts as an acyl transfer
reagent for the reaction with alcohol (or phenolic alcohol) to form an ester. For
reaction with pNP the phenolic reacts with the activated FA to form the pNP-acyl
ester product (see Figure 5.2.1 for reaction mechanism).
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113
Figu
re 5
.2.1
R
eact
ion
mec
hani
sm o
f the
Ste
glic
h es
terif
icat
ion
with
ED
CI.
R
1 =
acyl
cha
in g
roup
, R2
= p-
nitro
phen
ol.
CHAPTER V
114
In addition to the inclusion of DMAP, Qian et al. (2011) also implemented a washing
step in their pNP ester synthesis. However, we found it unnecessary and a single
silica gel column chromatography step with a mobile phase of DCM and petroleum
spirits (60-80 C) (1:1, v/v) was sufficient for purification without washing.
Furthermore, EDCI was used in this synthesis while N,N'-dicyclohexylcarbodiimide
(DCC) was used in the method of Qian et al. (2011). EDCI is known to give a more
soluble urea by-product than DCC and is more readily removed during workup
(Joullie & Lassen 2010). The modified protocol led to the synthesis of the olefinic
pNP-acyl esters with high yields using a simpler workup than previously reported.
Figure 5.2.2 shows an overview of the synthesis, including chemical structures and
reaction yields.
Figure 5.2.2 Synthesis, complete structures and yields of pNP-acyl esters.
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5.3 Characterisation of synthesised pNP-acyl esters
Unsaturated FAs, particularly LC n-3 PUFAs are generally regarded as unstable
given their sensitivity to oxygen. The presence of oxygen, elevated temperatures and
light can lead to oxidation, polymerisation and geometric isomerisation of double
bonds (cis - trans) (Shahidi et al. 1998). Therefore, it was important to fully
characterise the reaction products to ensure that no undesirable reactions had
occurred during synthesis. The presence of oxidative degradation or double bond
rearrangement would mean that lipase activity assays were not an accurate
representation of the enzymes affinity for that FA. A range of techniques were
utilised in the chemical characterisation, including mass spectrometry (MS), UV
spectroscopy, Fourier transform infrared (FTIR) spectroscopy and 1H and 13C NMR.
Compound characterisation data:
pNP-palmitoleate (3a); colourless oil. max 272nm. max 3004 cm-1. Found; [M+Na]+
398.2394 C22H35NO4 requires 398.2302. 1H NMR (500 MHz, CDCl3): δ/ppm 8.30
(d, 2H, J= 9.3 Hz), 7.30 (d, 2H, J=9.2 Hz), 5.41-5.34 (m, 2H,), 2.62 (t, 2H, J=7.6
Hz), 2.06-2.01 (m, 4H), 1.78 (qu, 2H, J=7.6 Hz), 1.45-1.24 (m, 16H), 0.90 (t, 3H,
J=7.1 Hz). 13C NMR (125 MHz, CDCl3): δ/ppm (pNP) 155.65, 145.36, 125.29,
122.54, (acyl) 171.40, 130.21, 129.78, 34.44, 31.91, 29.85, 29.78, 29.24, 29.17,
29.14, 29.11, 27.35, 27.25, 24.84, 22.78, 14.22.
pNP-oleate (3b); colourless oil. max 272nm. max 3003 cm-1. Found; [M+Na]+
426.2716 C24H39NO4 requires 426.2615. 1H NMR (500 MHz, CDCl3): δ/ppm 8.29
(d, 2H, J=9.1 Hz), 7.30 (d, 2H, J=9.2 Hz), 5.42-5.34 (m, 2H), 2.62 (t, 2H, J=7.5 Hz),
2.06-2.02 (m, 4H), 1.79 (qu, 2H, J=7.6 Hz), 1.45-1.29 (m, 20H, J= Hz), 0.90 (t, 3H,
J=7.1 Hz). 13C NMR (125 MHz, CDCl3): δ/ppm (pNP) 155.66, 145.37, 125.31,
122.55, (acyl) 171.40, 130.22, 129.78, 34.45, 32.03, 29.89, 29.79, 29.65, 29.46,
29.46, 29.25, 29.18, 29.15, 27.36, 27.27, 24.85, 22.81, 14.24.
pNP-linoleate (3c); pale yellow oil. max 272nm. max 3008 cm-1. Found; [M+Na]+
424.2554 C24H37NO4 requires 424.2458. 1H NMR (500 MHz, CDCl3): δ/ppm 8.30
(d, 2H, J=9.2 Hz), 7.30 (d, 2H, J=9.2 Hz), 5.40-5.35 (m, 4H), 2.84 (t, 2H, J=6.1 Hz),
2.63 (t, 2H, J=7.5 Hz), 2.09-2.05 (m, 4H), 1.78 (qu, 2H, J=7.5 Hz), 1.45-1.37 (m,
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116
14H), 0.91(t, 3H, J=7.1 Hz). 13C NMR (125 MHz, CDCl3): δ/ppm (pNP) 155.66,
145.39, 125.32, 122.56, (acyl) 171.41, 130.39, 130.09, 128.28, 128.00, 34.46, 31.67,
29.71, 29.48, 29.27, 29.20, 29.16, 27.35, 27.30, 25.78, 24.86, 22.72, 14.22.
pNP-linolenate (3d); pale yellow oil. max 272nm. max 3010 cm-1. Found; [M+Na]+
422.2427 C24H35NO4 requires 422.2302. 1H NMR (500 MHz, CDCl3): δ/ppm 8.30
(d, 2H, J=9.0 Hz), 7.30 (d, 2H, J=9.3 Hz), 5.45-5.33 (m, 6H), 2.80 (t, 4H, J=6.6 Hz),
2.63 (t, 2H, J=7.6 Hz), 2.13-2.07 (m, 4H), 1.78 (qu, 2H, J=7.5 Hz), 1.45-1.33 (m,
8H), 1.00 (t, 3H, J=7.5 Hz). 13C NMR (125 MHz, CDCl3): δ/ppm (pNP) 155.65,
145.38, 125.32, 122.55, (acyl) 171.4, 132.11, 130.30, 128.45, 128.34, 127.96,
127.22, 34.45, 29.68, 29.25, 29.19, 29.14, 27.31, 25.75, 25.66, 24.85, 20.68, 14.41.
pNP-eicosapentaenoate (3e); pale yellow oil. max 272nm. max 3011 cm-1. Found;
[M+Na]+ 446.2397 C26H35NO4 requires 446.2302. 1H NMR (500 MHz, CDCl3):
δ/ppm 8.3 (d, 2H, J=9.2 Hz), 7.30 (d, 2H, J=9.2 Hz), 5.51-5.34 (m, 10H), 2.88-2.83
(m, 8H), 2.64 (t, 2H, J=7.5 Hz), 2.24 (q, 2H, J=7.2 Hz), 2.10 (qu, 2H, J=7.6 Hz),
1.88 (qu, 2H, J=7.3 Hz), 1.0 (t, 3H, J=7.5 Hz). 13C NMR (125 MHz, CDCl3): δ/ppm
(pNP) 155.59, 145.40, 125.33, 122.53, (acyl) 171.21, 132.20, 129.52, 128.74,
128.61, 128.46, 128.45, 128.16, 128.13, 127.95, 127.11, 33.76, 26.55, 25.78, 25.78,
25.76, 25.67, 24.63, 20.69, 14.41.
pNP-docosahexaenoate (3f); light yellow oil. max 272nm. max 3013 cm-1. Found;
[M+Na]+ 472.2572 C28H37NO4 requires 472.2458. 1H NMR (500 MHz, CDCl3):
δ/ppm 8.29 (d, 2H, J=9.2 Hz), 7.30(d, 2H, J=9.2 Hz), 5.55-5.34 (m, 12H), 2.92-2.82
(m, 10H), 2.70 (t, 2H, J=7.1 Hz), 2.56 (q, 2H, J=7.0 Hz), 2.10 (qu, 2H, J=7.5 Hz),
1.0 (t, 3H, J=7.6 Hz). 13C NMR (125 MHz, CDCl3): δ/ppm (pNP) 155.55, 145.43,
125.33, 122.55, (acyl) 170.73, 132.19, 130.24, 128.73, 128.61, 128.44, 128.30,
128.15, 128.14, 127.96, 127.93, 127.20, 126.12, 34.37, 25.78, 25.78, 25.78, 25.75,
25.66, 22.74, 20.69, 14.41.
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5.3.1 Mass spectrometry and UV spectroscopy
The molecular formula of each pNP derivative was confirmed using mass
spectrometry, with all recorded masses being in agreement with the theoretically
calculated values (see Section 5.3). UV-spectra were defined from data obtained by
the HPLC’s diode array detector. An absorbance maximum of approximately 270 nm
corresponds to the pNP-acyl ester system and is clearly distinguished from the free
pNP absorbance maximum of 315 nm. The absorbencies of ~270 nm and ~315 nm is
consistent with that found by Iacazio et al. (2000) in regard to pNP-ester and pNP
UV absorbance, respectively.
5.3.2 Fourier transform infrared spectroscopy
Attenuated total reflectance Fourier transform infrared spectroscopy (ATR-FTIR)
was also used for molecular characterisation of FAs present in the synthesised
compounds. The 2nd derivative ATR-FTIR spectra of the derived pNP-acyl esters
(obtained using a 9-point Savitzky-Golay algorithm (Savitzky & Golay 1964)),
reveal the C-H stretching band of olefinic C=CH- chains in the FAs were observed
within the spectral range of 3035-2980 cm-1 (Figure 5.3.2). The 2nd derivative is used
as it increases peak resolution when multiple spectra are over-lapping or in close
proximity to one another. The olefinic (C=CH-) band occurs as a result of
unsaturation in the FA moiety present in the pNP-acyl esters and is therefore
commonly used for examining the degree of unsaturation in lipids and oils (Sills et
al. 1994; Yoshida & Yoshida 2004). The higher the number of olefinic bonds the
higher the wavenumber of the peak maximum (Sills et al. 1994; Yoshida et al. 2004).
In accordance with this, the peak maximum of 3003 cm-1, observed for pNP-OA
(C18:1 n-9), shifted to 3013 cm-1 for pNP-DHA (C22:6 n-3). The higher intensity of
the band presented in the highly unsaturated pNP-DHA further suggests a significant
increase in the degree of unsaturation in the oils. These results are in agreement with
those found via the other characterisation methods.
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118
16:1 n-7 (3004)
22:6 n-3 (3013)
20:5 n-3 (3011)
18:3 n-3 (3010)
18:2 n-6 (3008)
18:1 n-9 (3003)
Figure 5.3.2 ATR-FTIR analyses of the synthesised pNP-acyl esters. Presented as the normalised
2nd derivative C-H stretching band of olefinic C=CH- chains observed within the 3035-2980 cm-1
spectral range of the ATR-FTIR spectra. Synthesised pNP-acyl esters including pNP-palmitoleate
(C16:1 n-7), pNP-OA (C18:1 n-9), pNP-LA (C18:2 n-6), pNP-ALA (C18:3 n-3), pNP-EPA (C20:5 n-
3) and pNP-DHA (C22:6 n-3).
5.3.3 1H and 13C NMR
NMR data was important for confirming the structures, and determining if
isomerisation or oxidative degradation had occurred during synthesis. The 1H and 13C NMR spectra (unassigned) for the six synthesised pNP-acyl esters are presented
in Section 5.3. 13C NMR spectra are presented in a table format and relevant carbons
labelled for easier interpretation (Table 5.3.3). Spectra were also generated for the
FAs prior to the ester synthesis to ensure correct conformation (refer to Appendix
8.7, Table 8.7). Ester bond formation resulted in a chemical shift from ~180.5 ppm to
~171.5 ppm for the carboxylic carbon. The correct number of carbon signals was
found by 13C NMR for each synthesised compound. The right number of olefinic
carbon shifts were also observed for each compound, with two shifts observed for
each alkene in the acyl chain; ranging from 2-12 shifts in the pNP-palmitoleate
(C16:1 n-7) and pNP-DHA (C22:6 n-3) derivatives, respectively. All shifts involving
olefinic groups are in the approximate spectral region of 127-132 ppm, while those
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associated with allylic carbons are found between 14-35 ppm (Aursand & Grasdalen
1992). Chemical shifts from allylic carbons adjacent to two olefinic carbons
(methylene interrupted double bonds) found in polyunsaturated acyl groups were
also present in the expected numbers. The desired shift associated with these allylic
carbons is δ ≈ 25.6, indicating that both neighbouring olefinic groups are in the cis
configuration. This chemical shift migrates upfield toward δ ≈ 30.5 or δ ≈ 35.5 if one
or two trans isomers are present, respectively (Aursand et al. 1992; Rakoff & Emken
1983; Vatele et al. 1995).
Table 5.3.3 The observed 13C NMR chemical shifts (δ) of synthesised pNP-acyl esters.
Carbon Fatty acid C16:1 n-7 C18:1 n-9 C18:2 n-6 C18:3 n-3 C20:5 n-3 C22:6 n-3 pNP
155.65 155.66 155.66 155.65 155.59 155.55 145.36 145.37 145.39 145.38 145.40 145.43 125.29 125.31 125.32 125.32 125.33 125.33 122.54 122.55 122.56 122.55 122.53 122.55
Acyl group Carbonyl (C1) 171.40 171.40 171.41 171.4 171.21 170.73
130.21θ 130.22θ 130.39θ 132.11θ 132.20θ 132.19θ 129.78θ 129.78θ 130.09θ 130.30θ 129.52θ 130.24θ 34.44 34.45 128.28θ 128.45θ 128.74θ 128.73θ 31.91 32.03 128.00θ 128.34θ 128.61θ 128.61θ 29.85 29.89 34.46 127.96θ 128.46θ 128.44θ 29.78 29.79 31.67 127.22θ 128.45θ 128.30θ 29.24 29.65 29.71 34.45 128.16θ 128.15θ 29.17 29.46 29.48 29.68 128.13θ 128.14θ 29.14 29.46 29.27 29.25 127.95θ 127.96θ 29.11 29.25 29.20 29.19 127.11θ 127.93θ 27.35ψ 29.18 29.16 29.14 33.76 127.20θ 27.25ψ 29.15 27.35ψ 27.31ψ 26.55ψ 126.12θ 24.84 27.36ψ 27.30ψ 25.75ǂ 25.78ǂ 34.37 22.78 27.27ψ 25.78ǂ 25.66ǂ 25.78ǂ 25.78ǂ 14.22 24.85 24.86 24.85 25.76ǂ 25.78ǂ
22.81 22.72 20.68ψ 25.67ǂ 25.78ǂ 14.24 14.22 14.41 24.63 25.75ǂ
20.69ψ 25.66ǂ 14.41 22.74ψ
20.69ψ Methyl (ω) 14.41 Olefinic carbon θ, allylic carbon adjacent to a single olefinic carbon ψ, allylic carbon adjacent to two olefinic carbons ǂ.
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From this information it was determined that all double bonds in the synthesised
pNP-acyl esters were in the native cis configuration. In Table 5.3.3 with pNP-EPA as
an example, this means that 20 carbons can be found in relation to the acyl chain, 10
of which are involved in the 5 double bonds (~127-132 ppm). Between the 5 double
bonds there are 4 allylic carbons (methylene interrupted double bonds, ~25.6 ppm)
and another one allylic carbon at each end (20.69 and 26.55 ppm in EPA). This
indicates the correct chain length and number of double bonds are present and that
the double bonds are in the cis configuration.
The 1H NMR spectra and integration values for the derivatives were also consistent
with the expected structures. Chemical shifts associated with pNP were
approximately 8.3 and 7.3 ppm which is consistent with previous findings (Qian et
al. 2011). The acyl group shifts were found to be consistent to their grouping in
respect to allylic and olefinic groups. For example, the DHA derivative had 12
olefinic protons with the associated shifts (δ = 5.55-5.34 ppm) and 10 bis-allylic
protons adjacent to two olefins (δ = 2.92-2.82 ppm), which is in agreement with
previous assignments (Gunstone 1990).
5.4 Stability analyses of synthesised pNP-acyl esters
Due to the high sensitivity of EPA and DHA to oxidation, it was important to
determine the stability of these synthesised pNP-acyl esters, both in terms of their
‘assay’ and ‘storage’ stabilities. Gas chromatography (GC) was initially trialled as a
method for analysing compound stability. However, the high temperatures required
to enable compound migration through the GC column resulted in EPA and DHA
esters showing significant peak tailing; presumably as a result of compound
breakdown. Thus peak areas obtained were inaccurate and this method was
unsatisfactory (data not shown). Reverse phase HPLC was then trialled. This method
gave good peak shape and thus was subsequently used for assessing the breakdown
(if any) of the pNP-acyl esters, including the appearance of resultant breakdown
products. Before analyses began, compounds (pNP, butylated hydroxytolulene
(BHT), palmitoleic acid, OA, LA, ALA, EPA, DHA and the respective pNP
derivatives) were run to determine the elution times. This aided in differentiation
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between spontaneous hydrolysis products and those that arose through possible
degradation and isomerisation.
5.4.1 Storage stability analyses
Stability of the olefinic pNP-acyl esters stored at -20 C, both in the presence and
absence of BHT (+/-BHT) was monitored for a period of 6 months. Samples were
removed from cold storage and subjected to HPLC periodically, to determine long-
term storage stability. HPLC traces were analysed at 270 nm with the absorbance
spectrum of individual peaks analysed. Peaks optimally absorbing at ~270nm
(indicative of a pNP-acyl ester) were included in the analyses. Over a 6 month period
little degradation or isomerisation was observed for any of the pNP esters. Figure
5.4.1.1 shows the HPLC traces for pNP-palmitoleate, -OA and -LA. Traces for the
omega-3 compounds synthesised are shown in Figure 5.4.1.2. On day 1 all
compounds presented clean traces with a single peak optimally absorbing at 272 nm,
this was denoted as 100% of the pNP-acyl ester compound. After 6 months of
storage +/-BHT, HPLC analysis of compounds indicated that a number of minor
peaks with absorbance optima ~270 nm had developed in some samples. The total
peak areas were integrated and used to calculate the percentage of the peak area
relative to that identified on day 1. pNP-palmitoleate and pNP-LA showed no
breakdown products +/-BHT, with each respective peak integrated at 100% of the
peak area after 6 months. After 6 months without BHT the major peak in the pNP-
OA trace represented 96.82% peak area, while in the presence of BHT this was
increased to 98.27%. For the omega-3 compounds all samples after 6 months had
small amounts of breakdown products. As expected, pNP-ALA was the most stable
with peak area percentages of 99.23 and 98.37%, with and without BHT,
respectively. Without BHT the longer more unsaturated compounds of pNP-EPA and
DHA made up 96.67 and 94.94% of the relevant peak area, respectively. In the
presence of BHT, pNP-EPA and DHA accounted for 96.87 and 99.19% of the
relevant peak areas, respectively. BHT appeared to have little protective effect on the
EPA compound while DHA showed less breakdown in its presence.
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122
Figure 5.4.1.1 Reverse phase-HPLC of mono-/di-unsaturated pNP-acyl esters. Compounds
analysed (A) pNP-palmitoleate, (B) pNP-OA and (C) pNP-LA. Bottom trace on each plot is day 1,
middle trace 6 months without BHT and top trace 6 months with BHT.
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Figure 5.4.1.2 Reverse phase-HPLC of omega-3 pNP-acyl esters. Compounds analysed (A) pNP-
ALA, (B) pNP-EPA and (C) pNP-DHA. Bottom trace on each plot is day 1, middle trace 6 months
without BHT and top trace 6 months with BHT.
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The levels of breakdown over 6 months were found to be low with all compounds
showing <6% appearance of breakdown products. Addition of the antioxidant BHT
did reduce the amount of breakdown; however in its absence breakdown was still
low. The low level of degradation for all compounds was confirmed by 1H and 13C
NMR analysis of all stored samples after 6 months. The spectra produced were found
to be identical to those obtained immediately after synthesis with little sign of
contaminants (data not shown). When examining the signals at 210 nm (FFA
maxima) and 315 nm (pNP maxima) low levels of spontaneous hydrolysis were
observed amounting to <5% over 6 months. The addition of BHT did not appear to
reduce levels of spontaneous hydrolysis in any of the samples.
5.4.2 Assay stability analyses
Analysis of the stability of pNP-acyl esters under the conditions of the assay was also
required to ensure that the olefinic acyl groups did not undergo any structural
changes as a result of being introduced into an aqueous oxygen-rich environment. To
determine stability during the course of the assay, pNP-acyl esters were assayed in
the absence of any lipase, extracted, and analysed by HPLC for the presence of any
degradation products (see Section 2.1.27). As in Section 5.4.1 the peak areas of each
HPLC trace were integrated. No degradation products were observed that would
suggest modification of the acyl chain had occurred. This is probably a result of the
short assay duration, where the compounds are not exposed long enough to oxidative
conditions for oxidation or isomerisation to occur. Extra precautions such as using a
nitrogen blanket or carrying out the assay in darkness are possible, but appear to be
unnecessary.
All pNP-acyl esters underwent a low level of spontaneous hydrolysis, as is known to
occur for other pNP esters when performing the assay (Qian et al. 2011). There were
small differences in the level of spontaneous hydrolysis between omega-3 derivatives
and the other three substrates. However, the level of spontaneous hydrolysis is small
and can be negated with the correct use of assay blanks. With the above investigation
completed the pNP-acyl esters were deemed suitable for use in the pNP activity
assay protocol (Section 2.1.8).
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5.5 Screening of lipase fatty acid selectivity using pNP-acyl esters
Assays based on synthetic substrates, such as p-nitrophenyl esters for the
measurement of lipase activity have long been established in the field. They provide
a relatively simple, rapid and dependable method for analysing lipase and esterase
activity that can be applied in a high-throughput fashion. However, in regard to
lipase activity, they cannot be substituted for natural substrates (TAG), and caution
must be observed when attempting to make direct comparisons (discussed in Section
1.5). Thus this extended pNP-acyl ester library was designed not as an assay for total
characterisation, but as an additional tool for use in screening, characterisation and
analysis of lipase activity, particularly regarding LC n-3 PUFAs.
Selectivity profiles were generated for five commercial lipases using a range of
commercially available saturated pNP-acyl esters (C2:0, C4:0, C8:0, C12:0, C14:0,
C16:0, C18:0) and the newly synthesised unsaturated pNP-acyl esters (C16:1 n-7,
C18:1 n-9, C18:2 n-6, C18:3 n-3, C20:5 n-3, C22:6 n-3). All assays were performed
in triplicate and repeated once. The five lipases used were lipase B from Candida
antarctica (CalB), Aspergillus niger lipase (AnL), Candida rugosa lipase (CrL),
Rhizomucor miehei lipase (RmL) and Thermomyces lanuginosus (formerly
Humicola) lipase (TlL). The fungal lipases were chosen as they represent the group
of lipases most widely utilised in industry (Singh et al. 2012). They also provide an
appropriate starting point to validate these pNP-ester substrates, as a great deal of
work has previously been carried out with them. All of these lipases, with the
exception of AnL, have also had their three-dimensional structures resolved
(Brzozowski et al. 1991; Derewenda et al. 1994; Grochulski et al. 1994a; Uppenberg
et al. 1994) allowing for structure function relationships to be considered. These
analyses were performed to investigate relative rates of hydrolysis by these lipases
towards the various FAs in the pNP assay format. This was done as a ‘proof of
concept’ to demonstrate that this assay format could be used to investigate not only
acyl chain-length selectivity, but also acyl chain-structure selectivity. Furthermore,
these screens could be used to elucidate potential applications for particular enzymes,
such as omega-3 concentration.
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5.5.1 Fatty acid selectivity profiles of Candida antarctica lipase B and
Aspergillus niger lipase
CalB is one of the most studied and applied microbial lipases, particularly in regard
to LC n-3 PUFA processing, due to its activity and compatibility with immobilisation
systems (Kralovec et al. 2010; Tecelao et al. 2010; Watanabe et al. 2009). CalB
displayed a relatively broad range of activity across the substrate array (Figure
5.5.1A). The poorest activity was observed against C2:0, while other saturated acyl
groups were all hydrolysed more readily with the highest against C14:0. Saturated
substrates of 8, 12, 16 and 18 carbons in length were hydrolysed at an almost
uniform rate. This is in agreement with published data (Qian et al. 2011). Mono- and
di-unsaturated substrates reacted relatively poorly by comparison to saturated
substrates of the same chain length. However, the omega-3 substrates of pNP-C18:3
n-3 (ALA) and pNP-C20:5 n-3 (EPA) were hydrolysed at rates similar to those
saturated esters of medium chain length. The substrate pNP-22:6 n-3 (DHA) was
more slowly hydrolysed than the shorter omega-3 acyl groups, with the rate of
reaction comparable to that of the poorly hydrolysed C2:0. The activity profile
generated for AnL resembled that of CalB (Figure 5.5.1B). The relative FA
selectivity of AnL, as observed with CalB, again favoured medium chain saturate
substrates with an optimal length of C14. Furthermore it discriminated against the
C2:0 and DHA pNP esters. These data indicate that these enzymes share a similar FA
selectivity that is most likely due to their similar tertiary structure.
The structure of AnL has not yet been resolved via x-ray crystallographic means and
thus structure function comparisons are speculative. However, a structure assembled
by homology and molecular modelling has been put forward by Shu et al. (2009).
Both CalB and AnL possess small lids relative to those of other lipases, with only
five (residues 142-146) and seven (residues 86-92) amino acids respectively (Shu et
al. 2009; Uppenberg et al. 1994). Many reports in the literature state the absence of a
lid in CalB. However, most recent data supports the presence of a small lid, although
the significance of its contribution in interfacial activation appears to be negligible
(Ferrario et al. 2011; Skjot et al. 2009). The active site of CalB is situated at the
bottom of a funnel-type binding site (Gutierrez-Ayesta et al. 2007; Pleiss et al. 1998).
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Figure 5.5.1 Activity profiles depicting the relative activity of CalB and AnL against a range of
pNP-acyl esters. The relative activity of each substrate is normalised to C14:0 as 100%. (A) CalB and
(B) AnL. Substrates used in screen are pNP-acetate (C2:0), pNP-butyrate (C4:0), pNP-octanoate
(C8:0), pNP-dodecanoate (C12:0), pNP-myristate (C14:0), pNP-palmitate (C16:0), pNP-palmitoleate
(C16:1), pNP-stearate (C18:0), pNP-oleate (C18:1), pNP-linoleoate (C18:2), pNP-linolenoate (C18:3),
pNP-eicosapentaenoate (C20:5) and pNP-docosahexaenoate (C22:6).
0
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The structure proposed by Shu et al. (2009) in conjunction with the similar activity
profiles observed here, indicates that the active site of AnL may also be based around
a funnel-type topology. Lipases possessing this active site structure accept a broad
range of acyl groups and also exhibit activity usually associated with esterases; by
acting on soluble short chain substrates. For this reason these lipases may not be
particularly suitable for the concentration of LC n-3 PUFAs in glyceride substrates.
5.5.2 Fatty acid selectivity profile of Candida rugosa lipase
CrL was more selective than CalB and AnL, particularly in regard to omega-3
substrates (Figure 5.5.2). CrL was most active against C12:0. Short chain saturate
acyl groups were poorly hydrolysed with activity against C2:0 negligible. Activity
for C18:0 was also relatively poor, approximately 5-fold lower than that of C12:0.
The synthesised olefinic pNP substrates provide insight into the lipases activity.
Esters of palmitoleic acid (C16:1 n-7) and ALA (C18:3 n-3) groups were the most
readily hydrolysed, with the latter hydrolysed at rates close to that for C12:0.
However, on pNP-acyl esters longer than C18, CrL performed the hydrolysis at
vastly lower rates. Activity against EPA (C20:5 n-3) was 10-fold lower than the most
favoured C12:0 substrate, while DHA (C22:6 n-3) hydrolysis was minimal. These
results indicate that CrL may be useful for the concentration of EPA and DHA in
glyceride substrates. The structure of CrL is unique compared to the other lipases
tested here, as its active site resides within a deep tunnel that is buried inside the
protein (Pleiss et al. 1998). Observations from a crystal structure obtained for CrL
with a hexadecyl substrate, equivalent to a C17 acyl chain, show that the catalytic
triad lies near the entrance to the tunnel, with the tunnel narrowing near where the
omega terminus of an C3 acyl group would be positioned (Pleiss et al. 1998). This is
consistent with the poor activity observed with substrates ≤ C4, as without the
required length for sufficient interaction between the tunnel surface and acyl group,
the substrate may not be suitably positioned for hydrolysis. The cavity also appears
to be too small to effectively hold acyl chains greater than C18, possibly explaining
the low level of hydrolysis for EPA and DHA substrates. The hydrolysis profile
generated here is consistent with the hydrolysis of TAG substrates by CrL, where
DHA is selectively retained in the glyceride fraction (personal communication, Dr.
Matthew Miller, Plant and Food Research New Zealand).
CHAPTER V
129
Figure 5.5.2 Activity profile depicting the relative activity of CrL against a range of pNP-acyl
esters. The relative activity of each substrate is normalised to C12:0 as 100%. Substrates used in
screen are pNP-acetate (C2:0), pNP-butyrate (C4:0), pNP-octanoate (C8:0), pNP-dodecanoate
(C12:0), pNP-myristate (C14:0), pNP-palmitate (C16:0), pNP-palmitoleate (C16:1), pNP-stearate
(C18:0), pNP-oleate (C18:1), pNP-linoleoate (C18:2), pNP-linolenoate (C18:3), pNP-
eicosapentaenoate (C20:5) and pNP-docosahexaenoate (C22:6).
5.5.3 Fatty acid selectivity profiles of Thermomyces lanuginosus and
Rhizomucor miehei lipases
The FA selectivity of TlL was different from the three lipases discussed previously.
This lipase was selective for saturated FA’s of C8:0 and longer, with the highest
relative hydrolysis rates against pNP-acyl esters of C8:0 and C12:0 (Figure 5.5.3A).
Substrates C14:0 and C16:0 were also accepted relatively well by the lipase, while
C18:0 was to a lesser extent. TlL showed very poor hydrolysis of the shortest
saturates (≤ C4:0). Interestingly, all pNP-acyl esters containing any unsaturation
were relatively poorly hydrolysed, irrespective of chain length. Of the olefinic
substrates palmitoleic acid (C16:1 n-7), followed by ALA (C18:3n-3) were the most
readily accepted substrates while DHA was the least preferred. This selectivity
partially explains why TlL has been shown to be useful for the partial concentration
of EPA and DHA from fish oil (Akanbi et al. 2013). The profile obtained for RmL
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CHAPTER V
130
(Figure 5.5.3.B) resembled that of TlL. RmL displayed an obvious preference for
saturated medium to long chain (C8-C18) acyl groups, with an optimal activity for
C12:0. Short saturate acyl esters (≤ C4) were hydrolysed extremely poorly. This is
consistent with the observation that RmL displayed very low activity towards water-
soluble substrates (Derewenda et al. 1994). Palmitoleic acid and ALA substrates
were the most readily hydrolysed olefinic acyl esters, while DHA was again the most
poorly hydrolysed. It is likely that these underlying similarities between TlL and
RmL can be linked to the similar active site structure of these two lipases.
The identification of lipases based on their coordination-substrate site has led to the
defining of three main lipase types (see Figure 1.3.3) (Gutierrez-Ayesta et al. 2007;
Pleiss et al. 1998). Both TlL and RmL belong to a group of lipases which contain a
cleft-type active site on the surface of the enzyme. This is unlike other lipases which
may have funnel- or tunnel-type active sites that recede back into the protein
structure (Gutierrez-Ayesta et al. 2007). The topology of the active sites possessed
by TlL and RmL have been found to be virtually identical (Derewenda et al. 1994),
again explaining the similarities in observed activity. It would appear that this type of
active site interacts with unsaturated acyl groups less favourably, perhaps being
unable to orientate the ester bond, in regard to the catalytic triad, as freely as with
saturated substrates.
CHAPTER V
131
Figure 5.5.3 Activity profiles depicting the relative activity of TlL and RmL against a range of
pNP-acyl esters. The relative activity of each substrate is normalised to C12:0 as 100%. (A) TlL and
(B) RmL. Substrates used in screen are pNP-acetate (C2:0), pNP-butyrate (C4:0), pNP-octanoate
(C8:0), pNP-dodecanoate (C12:0), pNP-myristate (C14:0), pNP-palmitate (C16:0), pNP-palmitoleate
(C16:1), pNP-stearate (C18:0), pNP-oleate (C18:1), pNP-linoleoate (C18:2), pNP-linolenoate (C18:3),
pNP-eicosapentaenoate (C20:5) and pNP-docosahexaenoate (C22:6).
0
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80
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CHAPTER V
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5.6 Observations from lipase activity profiles
From the pNP-acyl ester activity profiles generated for these five lipases, three
distinct areas of activity can be described: (1) The activity against short chain
saturated substrates; (2) the influence unsaturation has on activity with substrates of
the same chain length; (3) the activity and selectivity towards LC n-3 PUFAs.
The low activity of all assessed lipases for pNP-acetate (C2:0) and pNP-butyrate
(C4:0) is partially due to the aqueous solubility of these substrates under the assay
conditions, resulting in a lack of micelle structure formation. It is well established
that lipases react most favourably with water-insoluble substrates and factors such as
interfacial activation play an important role. Interfacial activation describes the
conformational change which lipases undergo when they make contact with the
water-lipid interface of an insoluble substrate. It is generally accepted that interfacial
activation causes a substantial increase in the rate of lipase hydrolysis (Nardini et al.
1999; Verger 1997). The activity of CalB and AnL for these substrates was
noticeably higher than that of the other three lipases, indicating that the active site
structure has some impact on the activity with short chain FAs. CalB has previously
been shown to be more accepting of water soluble substrates when compared with
other lipases (Kirk et al. 1992; Pleiss et al. 1998). It has been suggested that CalB is
an intermediate between esterases and true lipases (Uppenberg et al. 1994), which is
consistent with its ability to hydrolyse soluble substrates in our assays. These
findings indicate that AnL, with its similar structural features and activity profile, fits
into the same category.
The C18 series of substrates with none, one, two or three double bonds present, is a
useful series for comparing activity versus unsaturation for a single chain length.
Degree of saturation is not linearly correlated with ease of hydrolysis, as C18:0 was
hydrolysed more readily than ALA (C18:3 n-3) by most lipases, the exception being
CrL where the opposite was true. However, a general trend in those substrates with
double bonds present is that most of the lipases hydrolyse the more highly
unsaturated ALA (C18:3 n-3) more readily than OA (C18:1 n-9) and LA (C18:2 n-
6). This indicates that as unsaturation increases in substrates of the same length, so
does the rate of hydrolysis. The impact of unsaturation on the rate of hydrolysis can
be partially explained through describing the different substrate-protein interactions.
Two possible reasons for the observed differences are as follows. Firstly, the impact
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133
unsaturation has on surface tension and viscosity. It has been suggested previously
that lipase activity towards insoluble substrates changes with varying surface
tensions (Gargouri et al. 1989). Variance in chain-length and unsaturation can
directly influence surface tension and viscosity (Kordel et al. 1991). Thus the
increase in activity may not be due to lipase FA specificity itself, but more to the
qualities/properties of the lipid interface as a function of FA composition. Others
have proposed similar suggesting that the quality of the emulsion of certain
substrates influences lipase activity (Huang et al. 2004). Secondly, lipase FA
selectivity dictates which FA are more readily hydrolysed. The difference in activity
could be attributed to the increase in flexibility and disorder seen in increasingly
unsaturated acyl groups, often associate with membrane dynamics and fluidity
(Feller et al. 2005; Feller et al. 2002; Hyvonen et al. 2005). These properties could
allow the polyunsaturated acyl chains to better conform to the active site structure of
various lipases. This is of particular interest in regard to CrL, with such a structurally
involved and constrained active site it seems intuitive that as unsaturation increases
in the C18 substrates, so too does the hydrolysis rate. We propose that the impact of
emulsion quality is minimal and that the primary reason for differences in ease of
hydrolysis is lipase specificity. Defining these properties is largely outside of the
scope of this research. Understanding whether either explanation, or possibly a
combination of the two is responsible, requires further research.
The final area of discussion in this chapter involves the activity of these lipases
towards the LC n-3 PUFA pNP esters of EPA and DHA. Lipase activity observed
with these substrates was generally poor when compared with other acyl groups.
DHA was poorly hydrolysed by all five lipases, while EPA was hydrolysed more
readily than DHA by all lipases tested. The most obvious factor leading to
differences in EPA and DHA hydrolysis is likely related to chain-length, to
investigate this further would require the synthesis of saturate C20:0 and C22:0
substrates and comparing hydrolysis rates. Interestingly CalB and AnL acted on
ALA more poorly than EPA despite its shorter chain length, while the remaining
three lipases hydrolysed ALA better than EPA; suggesting that chain-length alone
isn’t the defining factor. Another factor leading to poor hydrolysis of DHA may be
related to steric hindrance. DHA can exist in a number of conformers, some of which
position the methyl end of the acyl chain in close proximity to the ester bond. The
association between FA conformation and steric hindrance of lipase access to the
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134
ester bond was first suggested by Bottino et al. (1967). The final possibility involves
the positioning of the double bond closest to the FA carboxyl terminus. This plays a
significant role in the rate of hydrolysis. Bottino et al. (1967) discussed this also and
the area has subsequently been studied in more detail by (Hills et al. 1990;
Mukherjee et al. 1993) using esterification reactions. It was concluded that acyl
chains with an initial double bond in the cis -4, cis -6 or cis -8 positions were
discriminated against by various lipases, while those with the first double bond
present in an odd numbered position were hydrolysed more readily. This supports
why EPA (cis -5) is cleaved more readily relative to DHA (cis -4), despite both
having an extensively unsaturated structure.
5.7 Summary
The pNP assay has been used extensively for defining the chain-length specificity of
lipases. We hypothesised that this assay platform could also be used for defining the
FA selectivity of lipases. The synthesis of six olefinic pNP-acyl esters, including
EPA and DHA derivatives, was optimised to achieve yields in excess of 85%.
Chemical analysis showed that all six compounds were present in the correct
conformation. All six compounds were stable under the conditions of the assay and
under storage conditions for at least 6 months. In conjunction with seven commercial
substrates, the six new pNP-acyl esters were then used for the screening of FA
selectivity in five commercial lipases. It was found that the extended pNP-ester
library provides a useful tool for determining FA selectivity. The findings
demonstrated that the profiles produced were consistent with regards to active site
structure-function relationship in the different lipases. CalB and AnL had similar
activity profiles which were relatively broad compared to the other lipases tested.
TlL and RmL, which have near identical active sites, had similar activities and
preferentially hydrolysed medium chain saturated substrates. It was also found that
CrL may be the most applicable for the concentration of EPA and DHA in glyceride
substrates. All lipases tested performed optimally on C14:0 and C12:0 pNP esters.
CHAPTER VI
135
CHAPTER VI
Calcium-stimulated activity in fungal lipases with cleft-type
active sites
6.0 Introduction
Understanding the physicochemical and structure-function relationships of
lipase/lipid-substrate catalytic mechanisms is of great importance, particularly as
optimal lipase activity is required for enzyme-mediated commercial processes.
Understanding these relationships is required not only for the particular enzymes and
substrates, but also at the level of protein-lipid interaction at an interface. There is
great interest in understanding how different lipases act at the lipid interface and the
factors that influence their action, particularly as this relates to methods for
manipulating the rate or specificity of reactions. Examples of this can be seen in the
modification of lipase activity for use in biofuel production (Juan et al. 2011),
detergents (Hasan et al. 2010), chemical synthesis (Baldessari 2012) and omega-3
concentration (Wanasundara et al. 1998). Despite the extensive research in this area,
the huge variety of lipase active site structures and conformational changes they
undergo, means that new findings and applications continue to emerge.
The profiling of the five commercial lipases (lipase B from Candida antarctica
(CalB), Aspergillus niger lipase (AnL), Candida rugosa lipase (CrL), Rhizomucor
miehei lipase (RmL) and Thermomyces lanuginosus (TlL)) with pNP-acyl esters
(Chapter V) was initially carried out using a generalised activity assay (Section
2.1.8) suitable for both calcium-dependent and independent lipases, where CaCl2 was
included in the buffer. The five fungal lipases used in these studies had been found
previously to act in a calcium-independent manner, with none of the resolved crystal
structures containing an associated calcium ion (Brzozowski et al. 1991; Derewenda
et al. 1994; Grochulski et al. 1994a; Uppenberg et al. 1994). Because of the apparent
independence from a calcium requirement for the tested enzymes, the assays were
subsequently run in the absence of calcium, expecting this to have no effect on the
rate of hydrolysis. Surprisingly, this was found not to be the case for some of the
lipases.
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Research on the influence of calcium on lipase activity has primarily focussed on
pancreatic lipases, as they possess a calcium ion binding site (Bourne et al. 1994;
Vantilbeurgh et al. 1993). Calcium is required for optimal pancreatic lipase activity
(Kimura et al. 1982) and its role in human health, the adsorption of poorly
bioavailable drugs and lipid hydrolysis in the human gut, have been extensively
researched (Chakraborty et al. 2009; Devraj et al. 2013; Golding & Wooster 2010).
In contrast, the function of calcium in stimulating the activity of fungal lipases has
not been reported on in-depth. This is perhaps due to the commercial importance and
sensitivity of such information. Generally speaking calcium and other divalent
cations such as magnesium are thought to stimulate lipase activity by removing
hydrolysed FAs as calcium-soaps (Gupta et al. 2004). However, many fungal lipases
appear not to be, or are only marginally stimulated by calcium (Knoche et al. 1970;
Patkar et al. 1993; Yu et al. 2007). In some cases calcium inhibits the activity of
fungal lipases (Khor et al. 1986; Linfield et al. 1984). Deciphering the influence of
calcium on lipases is difficult as findings can be greatly influenced by the substrates
used (Wang et al. 1988). As fungal lipases are extensively used in industry,
improvements in lipase catalysed hydrolysis are of value (Singh et al. 2012). Thus,
further investigation of the fundamental structure-function relationships of these
lipases is required.
Although not an initial research objective, this work arose from the study in Chapter
V and required further investigation. As discussed in Chapter V, the profiled lipases
fall into three classes; cleft-, funnel- and tunnel-type active sites. The most notable
effect of calcium exclusion was seen with RmL and TlL, which both possess cleft-
type active sites. This phenomenon was investigated further using both pNP-acyl
ester substrates and a number of emulsified glyceride substrates; including tributyrin,
tuna oil and lecithin. Hydrolysis of these substrates was analysed both in the
presence and absence of CaCl2 to observe differences in hydrolysis rates. The aim of
this study was to determine how different substrates or substrate mixtures react with
lipases in the presence and absence of calcium, establishing a starting point for future
research in this area.
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6.1 Increased pNP-acyl ester hydrolysis in the presence of calcium
It was observed that the removal of CaCl2 from the assay buffer resulted in an altered
rate of pNP ester hydrolysis by some lipases. An initial experiment was carried out
using the substrates pNP-C12:0 and pNP-C14:0 with a number of different buffer
compositions. Four different solutions were made up (all of which contained 20mM
Tris-HCl pH 8.0 and 0.01% gum arabic) that varied in the inclusion of 20 mM CaCl2,
5 mM CaCl2, 20 mM NaCl or nothing. The pH of the buffers was manually adjusted
by addition of HCl once salts had been added, this ensured changes in pH could not
influence activity. The respective buffers were then used under the same conditions
with each of the commercial lipases (CalB, AnL, CrL, RmL and TlL). Two
concentrations of CaCl2 were used to check if the change in activity was
concentration dependent. The NaCl was included to test if the ionic strength of the
buffer was causing the change in activity. The activity of each lipase in the different
buffer systems are displayed in Figure 6.1.
As was found in Chapter V, trends can be seen that appear to reflect the active site
topology of the lipases. CalB and AnL (funnel-type) again showing a similar
response, TlL and RmL (cleft-type) are also similar to each other, while CrL (tunnel-
type) is dissimilar to the rest. As expected the activity of CalB and AnL for
pNPC12:0 was essentially unchanged regardless of the salts used. However, the
hydrolysis of pNP-C14:0 gave an unexpected result. Reactions carried out in buffers
containing CaCl2 gave significantly higher hydrolysis rates than those with NaCl or
nothing. The NaCl and no-salt buffers gave essentially the same rate of hydrolysis,
suggesting that the ionic strength of the buffer does not influence the activity of these
lipases. It is currently unclear why increased hydrolysis of the myristic acid (C14:0)
substrate in particular, is observed in the presence of calcium. In subsequent work
(discussed in Section 6.2) the majority of FA’s tested resulted in little or no change
in activity.
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Figure 6.1 Activity of commercial lipases against pNP-acyl esters of medium chain-length in
the presence and absence of CaCl2 and NaCl. Lipases used were (A) CalB, (B) AnL, (C) TlL, (D)
RmL and (E) CrL. Blue and red bars represent the hydrolysis ( A410/min) of pNP-C12:0 and pNP-
C14:0, respectively.
0
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0.6
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00.10.20.30.40.50.60.70.80.9
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A B
C D
E
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139
The influence of calcium in the buffers was more pronounced with TlL and RmL.
Both enzymes showed a large difference in the hydrolysis of pNP-C12:0 in buffers
containing calcium. The 5 mM CaCl2 buffer gave greater activity with both enzymes
than the 20 mM, the reason for this is unclear. Activity was lowest in the buffer with
nothing added, while the addition of 20 mM NaCl appeared to raise levels of
hydrolysis; but more modestly than the CaCl2 buffers. The hydrolysis of pNP-C14:0
by RmL followed the same profile as that for C12:0, with calcium-containing buffers
giving the most activity. However, the hydrolysis by TlL on pNP-C14:0 did not
follow the same trend as C12:0. Activity was still higher in buffers containing CaCl2,
but the reaction carried out in 20 mM gave more activity than in 5 mM.
The data obtained for CrL showed the inclusion of CaCl2 in buffers generally had
less of an effect than on the other enzymes. There was a small increase in activity
against pNP-C14:0 when calcium was present, however this was to a lesser extent
than that observed for TlL and RmL. Activity against pNP-C12:0 showed slightly
higher hydrolysis with the 5 mM CaCl2 and 20 mM NaCl buffers relative to the
others. However there was no apparent difference between the buffer containing 20
mM CaCl2 and the one with nothing added.
These results demonstrated that the addition of calcium (in the form of CaCl2) was
affecting the hydrolysis rates for certain lipases against pNP-acyl ester substrates and
that this effect was not due purely to ionic strength. The reason for increased CalB
and AnL activity against C14:0 in the presence of calcium is currently uncertain.
However, one thing that could be determined is that when compared to the other
three lipases, both TlL and RmL displayed notable changes to their activity against
the substrates in the presence of calcium.
6.2 Calcium alters hydrolysis rate and not specificity
After the initial experiment showing calcium had altered the rate of substrate
hydrolysis by lipases, particularly lipases with cleft-type active sites, further profiling
was carried out with the pNP-acyl ester library. This was done to check if the
increase in hydrolysis was uniform across all FAs, or whether the removal of calcium
changed the FA selectivity of these lipases. The lipases TlL and RmL were chosen as
they were most influenced by CaCl2. CalB was used as a ‘reference’ lipase that was
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less affected by the addition/removal of calcium. Assays using the range of pNP-acyl
esters were performed as previously in Chapter V, with the omission of CaCl2 from
the assay buffer. The assay values ( A410/min) were then compared with those values
obtained for assays with calcium and all values converted to relative activity (%).
This manipulation allowed the activity recorded for each assay to be visualised
against all other assays for the respective lipase in a comparative manner. Figure 6.2
shows the pNP-acyl ester FA profiles for each of the three lipases when assayed both
with and without calcium. It can be clearly seen that TlL and RmL show a significant
decrease in activity against all substrates when CaCl2 is absent from the assay buffer,
with the exception of short-chain substrates where activity was already minimal.
Most substrates show approximately a 5-10 fold decrease in their hydrolysis when
calcium is excluded from the assay. Alternatively CalB, while showing some
variance in the hydrolysis of C12:0 and C14:0 (mentioned in Section 6.1), is
relatively indifferent in its activity across all FAs. As well as demonstrating the
change in hydrolysis rate for TlL and RmL, changes in the FA selectivity were
investigated. For ease of viewing (due to the low values when compared with
calcium included assays) the activity profile generated for each lipase without
calcium is presented in Appendix 8.8, Figure 8.8.
Despite a change in hydrolysis rate the profiles obtained without calcium display
similar FA selectivity to those obtained with calcium. Notably, CalB shows the
decrease in hydrolysis of C14:0, but the remaining profile remains very similar. TlL
also displays a very similar profile in the absence of calcium, with medium length
saturates most easily hydrolysed while short-chain and olefinic substrates are more
resistant to lipase activity. The dataset for RmL also displays a similar overall profile
when calcium is excluded from the reaction. With medium chain saturates most
favoured while short chain saturate and olefinic acyl groups are hydrolysed more
slowly. It must be noted that the standard deviation obtained for C8:0 and C14:0
substrates are relatively high. Also when compared to the calcium included profile,
the activity against saturates >C14 in length is lower.
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0
20
40
60
80
100
120
Rela
tive
activ
ity (%
)
pNP-acyl ester
0
20
40
60
80
100
120
Rela
tive
activ
ity (%
)
pNP-acyl ester
Figure 6.2 Activity profiles depicting the relative activity of CalB, TlL and RmL against a
range of pNP-acyl esters with and without CaCl2. (A) CalB, (B) TlL and (C) RmL. The relative
activity of each substrate is normalised to 100% of the substrate for which the highest activity was
obtained. Blue and red bars represent the presence and absence of CaCl2, respectively. Substrates used
in screen are pNP-acetate (C2:0), pNP-butyrate (C4:0), pNP-octanoate (C8:0), pNP-dodecanoate
(C12:0), pNP-myristate (C14:0), pNP-palmitate (C16:0), pNP-palmitoleate (C16:1), pNP-stearate
(C18:0), pNP-oleate (C18:1), pNP-linoleoate (C18:2), pNP-linolenoate (C18:3), pNP-
eicosapentaenoate (C20:5) and pNP-docosahexaenoate (C22:6).
A
B
C
0
20
40
60
80
100
120
Rela
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activ
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)
pNP-acyl ester
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The comparison of profiles generated using the pNP-acyl ester library, both in the
presence and absence of calcium, showed two things clearly. First, when calcium is
present there is a significant increase in activity on all FAs tested by lipases that
possess cleft-type active sites. This was not observed with CalB which has a funnel-
type active site. Secondly, the inclusion of calcium in the assays does not appear to
alter the FA selectivity of any of the lipases tested, as similar overall profiles were
obtained regardless.
6.3 Effect of calcium on hydrolysis of TAG substrates by lipases
After observing the effect calcium had on the hydrolysis of pNP-acyl esters by TlL
and RmL, further work was required to investigate whether this was specific to pNP-
ester substrates (mono-FA substrates), or if it also occurred with TAG substrates. As
TAG is the natural substrate for lipases, understanding modifications to the
hydrolysis of these substrates is more relevant to real-world lipase applications. As
the most significant changes in activity occurred with TlL and RmL these lipases
were chosen for further testing. CalB was again used as a ‘reference lipase’ for
comparison. As stated previously, to our knowledge all of the lipases tested had been
characterised and shown to act in a calcium-independent fashion. Titration assays
were performed as described in Section 2.1.9.
6.3.1 Effect of calcium on tributyrin hydrolysis
Tributyrin was the first TAG substrate tested to investigate the influence of calcium
on lipase activity. The substrate mixture contains tributyrin, lecithin and sodium
caseinate (see Section 2.0.4). Table 6.3.1 displays the results for tributyrin hydrolysis
by the three lipases with and without CaCl2. CalB hydrolysed the tributyrin substrate
readily in both reactions. However, addition of CaCl2 to the reaction resulted in
activity dropping by approximately 30%. This would suggest that calcium may
inhibit the activity of CalB on tributyrin to some degree. In contrast, both TlL and
RmL hydrolysis of tributyrin was increased significantly in the presence of calcium.
The hydrolysis rate of TlL increased by 170 times while RmL increased by 32 times
when CaCl2 was added. Accordingly, these lipases had to be diluted to obtain
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reproducible results when calcium was present (as seen in Table 6.3.1). These data
support the fact the calcium was stimulating activity in the lipases with cleft-type
active sites, but not in the funnel-type active site of CalB.
Table 6.3.1 Activity of lipases on tributyrin in the presence and absence of CaCl2.
Lipase CaCl2 DF Lipase vol. (μL)
0.01M NaOH titrated (mL/min)/1 mL of lipase
CalB - 10 25 105.19 7.91 CalB + 10 25 67.75 2.43 RmL - 5 100 8.53 1.73 RmL + 25 25 273.92 13.07 TlL - 1 200 2.52 0.074 TlL + 50 20 428.63 29.74 Dilution factor (DF)
6.3.2 Effect of calcium on hydrolysis of fish oil by lipases
After observing the effect of calcium on tributyrin hydrolysis it was decided that
another TAG substrate should be trialled. Tuna oil was used as it is relevant to other
research in our laboratory involving omega-3 concentration; it is also more relevant
to real world applications where oils being processed contain a mixture of FAs. The
substrate contains tuna oil and gum arabic (Section 2.0.4). Results from the tuna oil
hydrolysis in the presence and absence of CaCl2 can be seen in Table 6.3.2. CalB
was found to hydrolyse this emulsion at much lower rates than achieved with
tributyrin. However, the activity of RmL and TlL was significantly higher with the
fish oil emulsion. These findings are consistent with the composition of tuna oil
(consisting primarily of longer unsaturated FA) and the activity profiles seen with the
pNP-acyl esters (Figure 6.2). CalB reacted more readily with pNP-C4:0 (butyrate)
while TlL and RmL hydrolysed it poorly and acted more favourably with medium-
long chain substrates. Changes in the rate of tuna oil hydrolysis with or without
CaCl2 in the reaction mixture were negligible, with no significant increases in
activity found in the presence of calcium.
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Table 6.3.2 Activity of lipases on tuna oil in the presence and absence of CaCl2.
Lipase CaCl2 DF Lipase vol. (μL)
0.01M NaOH titrated (mL/min)/1 mL of lipase
CalB - 1 100 1.22 0.19 CalB + 1 100 2.08 0.66 RmL - 25 25 326.12 50.07 RmL + 25 25 291.48 23.86 TlL - 1000 25 16300.00 1254.91 TlL + 1000 25 18908.40 2602.48 Dilution factor (DF)
This was contradictory to the observation of tributyrin hydrolysis being increased in
lipases with cleft-type active sites in the presence of calcium. This indicated that
there might be something in the emulsion mixture of the tributyrin substrate that was
working in conjunction with calcium to increase substrate hydrolysis. The tuna oil
emulsion only contained oil and gum arabic as an emulsifier, while the tributyrin
contained lecithin which is primarily composed of phospholipids. TlL and RmL have
been previously shown to be capable of hydrolysing phospholipid substrates and
their activity against TAG is stimulated in the presence of phospholipid (Gutierrez-
Ayesta et al. 2007). Subsequent investigation was undertaken to determine whether
the increase in activity against tributyrin was in fact due to the release of FAs from
lecithin in the assay mixture when calcium was present.
6.4 Effect of calcium on hydrolysis of lecithin by lipases
The hydrolysis of phospholipids and phospholipid/TAG mixtures by lipases with
cleft-type active sites has been previously reported by Gutierrez-Ayesta et al. (2007).
Despite the presence of a lid (which is usually associated with true lipases and
activity on TAG) these enzymes have been shown to be capable of hydrolysing a
number of phospholipids and galactolipids (Amara et al. 2013). However, the
influence of calcium on these reactions has not been described. A lecithin emulsion
was made that contained lecithin and sodium caseinate (Section 2.0.4). Results for
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the hydrolysis of lecithin by the three lipases with and without CaCl2 can be found in
Table 6.4. The hydrolysis of this substrate by CalB was negligible in all assays,
regardless of whether calcium was present. This was expected as CalB has been
shown previously not to hydrolyse lecithin substrates (Gutierrez-Ayesta et al. 2007).
In the absence of calcium RmL also reacted poorly with the substrate, which was
unexpected as Gutierrez-Ayesta et al. (2007) showed that lecithin was hydrolysed
well by this lipase.
Table 6.4 Activity of lipases on lecithin in the presence and absence of CaCl2.
Lipase CaCl2 DF Lipase vol. (μL)
0.01M NaOH titrated (mL/min)/1 mL of lipase
CalB - 1 20 1.03 1.37 CalB + 1 20 -1.19 0.75 RmL - 5 100 0.31 0.36 RmL + 5 100 5.47 0.60 TlL - 10 20 3.80 6.08 TlL + 10 20 174.29 21.82 Dilution factor (DF)
The low activity observed may be due to the low amount of lipase used in the
reaction. Nonetheless, when calcium was added to the reaction RmL activity
increased markedly, again suggesting that calcium was stimulating hydrolysis in this
reaction. The hydrolysis of this substrate by TlL without calcium again gave
negligible levels of hydrolysis; again this may be due to the low amounts of enzyme
used in the assays. As was seen with RmL, the addition of calcium led to a large
increase in substrate hydrolysis by TlL. These findings further support that the
activity of lipases with cleft-type active sites are stimulated by calcium.
Lecithin is a generic term used to describe lipid substances isolated from plant or
animal sources that may contain a range of lipid species, including phosphoric acid,
choline, FAs, glycerol, glycolipids, TAG and phospholipids. The primary constituent
is usually phospholipids such as phosphatidylcholine. The lipid composition of the
lecithin used in this work was unknown. It was important to analyse the lecithin and
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determine its lipid composition as this may influence the activity observed. This
meant that while calcium was affecting the activity of TlL and RmL, the lipases may
not have been acting on only phospholipid, as was intended by the assay. Further
work was required to assess the hydrolysis of the lecithin substrate and the
subsequent hydrolysis products.
6.5 Analysis of lecithin and lecithin hydrolysis products
The measurement of FA release in lecithin by TlL and RmL confirms that activity in
these enzymes is stimulated in the presence of calcium. However, having learned that
the lecithin used in this work may contain TAG, further investigation was required.
A hydrolysis of lecithin was performed and the reaction products analysed (Section
2.1.28). An Iatroscan was used to analyse the composition of the lecithin substrate
both before and after hydrolysis by lipases in the presence and absence of CaCl2.
Hydrolysis reactions were carried out at pH 8.0 at 30 C with stirring. Table 6.5
displays Iatroscan analysis of the peak area percentages of lipid classes found in the
lecithin substrate before and after hydrolysis.
The Iatroscan trace showed three main peaks that represented TAG, FFA and polar
lipid (PL, predominantly phospholipid), as determined by external standards. Before
hydrolysis the lecithin contained approximately 30% TAG, 2% FFA and 68% PL. It
should be noted that the results displayed here are only relative for each individual
lipase tested, as the amount of hydrolysis is dependent on lipase concentration.
Results are preliminary as enzyme concentration has not yet been optimised.
Hydrolysis of lecithin by RmL in the absence of calcium resulted in no increase in
FFA and little change in TAG and PL levels from that of the un-reacted lecithin. The
addition of calcium to the reaction also resulted in no significant hydrolysis by RmL.
The reactions carried out with TlL showed more activity against the lecithin
substrate. Without calcium the TAG peak was reduced to ~11% of total peak area of
lipids and FFA increased to ~9%. The PL peak did not decrease; this indicated that
the lipase was hydrolysing the TAG portion of the lecithin and not the phospholipid
as first thought. When calcium was added to the reaction with TlL the TAG was
almost completely hydrolysed, with TAG only accounting for 1% of lipid after
hydrolysis, while FFA increased to ~22% of all lipid post hydrolysis. Again this
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demonstrated that TAG was undergoing hydrolysis and that calcium was stimulating
the hydrolysis of TAG and not phospholipids in the lecithin substrate.
Table 6.5 Composition (%) of lecithin before and after reaction with lipases.
Lipase CaCl2 TAG FFA PL
No lipase n/a 30.18 1.82 68
RmL - 31.38 1.42 67.2
RmL + 28.2 1.05 70.74
TlL - 11.34 8.65 80.01
TlL + 0.98 21.62 77.4025
Triacylglycerol (TAG), free fatty acid (FFA) and polar lipid (PL)
This was checked by further titration assays with the lecithin substrate using TlL. A
titration assay was carried out over an hour using an emulsion not containing CaCl2.
The assay was allowed to proceed for 20 min before CaCl2 was added to a final
concentration of ~5 mM. Appendix 8.9, Figure 8.9 shows an example of a titration
trace obtained for the assay. As can be seen, the addition of the CaCl2 caused activity
to increase dramatically. Hydrolysis continued at this rate before gradually
decreasing to a stable slower hydrolysis rate. In combination with the Iatroscan data
it is possible to better follow what is occurring: (1) The calcium stimulates the
hydrolysis of TAG by TlL and (2) as the TAG concentration decreases in the
emulsion, the amount of available substrate declines and the hydrolysis rate drops
concurrently. (3) Once all TAG has been hydrolysed TlL then continues to hydrolyse
phospholipids at a lower rate (as it is known to do). This work validates that the
hydrolysis of TAG by these lipases with cleft-type active sites is stimulated by
calcium. However, stimulation only occurs when the emulsion also contains
phospholipids, as shown when no change in activity was observed in Section 6.3.2.
An exact mechanism for this is currently unknown and further work is required to
interpret whether the effect is a result of enzymatic function, the quality of the lipid
interface, or a combination of the two.
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6.7 Analysis of emulsion properties
Stimulation of hydrolysis by calcium was only observed in lipases with cleft-type
active sites. This suggests that lipases with this active site conformation were
influenced by calcium, or that the cleft-type active site acts more readily at the lipid
interface as a result of physical properties imparted by the calcium on the lipid.
Research has been carried out with pancreatic lipases showing that calcium is a
potent activator of these lipases (Golding et al. 2010). The impact of calcium ions on
lipase hydrolysis could be due to either a direct modification of the protein structure
and function, or to a substrate modification that facilitates improved access by the
lipase. Here we investigated whether or not modification of the physical properties of
micelles, or the so called ‘interfacial quality’ (Verger et al. 1973), was responsible
for the observed calcium-activation. Modifications might include altering micelle
size distribution and/or surface charge in a way that facilitates better interaction with
the lipases. Understanding this would aid in better determining why TAG hydrolysis
(in TAG-phospholipid mixtures) by lipases with cleft-type active sites was altered by
calcium.
A Malvern Nano Zetasizer (Malvern Instruments Ltd., Malvern, UK) was used to
determine the surface charge (zeta potential) of lecithin micelles (Section 2.1.29).
This provides insight into the physicochemical properties of the lipid-water interface
where the lipases act. The emulsions were prepared in duplicate for both samples
with and without CaCl2. The emulsion without CaCl2 gave an average zeta potential
value of -38.85 mV, while the sample with CaCl2 added generated a value of -14.75
mV. The addition of calcium resulted in a reduction in zeta potential. Due to the
zwitterionic head group of phospholipids, the surface charge of micelles in the
absence of surfactants is determined mainly by pH (Tadros & Vincent 1983). The pH
of emulsions was regulated by buffering at pH 8.0 to remove this variable. When
different compounds or ions, such as bile salts and calcium are added to the
emulsions, the micelle surface charge can be altered. In studies with pancreatic lipase
Wickham et al. (1998) found that addition of bile salts made the zeta potential of
TAG-phospholipid mixed micelles much more negative. They also analysed the
influence of calcium on zeta potential, finding that it reduced the zeta potential. The
results of Wickham et al. (1998) agree with the proposal that calcium ions act as a
shield of micelle surface charge, as described by Hunter (1981). The experiments
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performed here do not contain bile salts; however they do contain sodium caseinate.
Sodium caseinate carries a negative charge at pH above its isoelectric point (~4.6)
(Hu et al. 2003), thus explaining the negative value (-38.85 mV) despite the lack of
bile salt. The observation that calcium decreased the zeta potential of the lecithin
micelles in this research from -38.85 to -14.75 mV, is in agreement with the
shielding mechanism proposed. How or if this reduction of surface charge influences
lipase activity is unclear. Wickham et al. (1998) found no significant correlation
between zeta potential and pancreatic lipase activation/activity, still this cannot be
ruled out as a factor that influences activity with other types of lipase.
The influence of calcium on micelle size distribution was investigated by dynamic
light scattering using a Malvern Mastersizer 2000 (Malvern Instruments Ltd.,
Malvern, UK), (Section 2.1.30). Measurements of the micelle size distribution with
and without CaCl2 are displayed in Figure 6.7. The size distribution for both micelle
preparations was approximately 0.1-35 μm and presented as bimodal distributions.
For each emulsion the distribution values were obtained and the volume mean
diameter (D[4,3] value) was calculated. The solution without CaCl2 contained
micelles with d(0.1) = 0.32 μm, d(0.5) = 1.08 μm, d(0.9) = 12.32 μm and D[4,3] =
3.91 μm. When CaCl2 was added the distribution shifted towards larger sized
micelles, d(0.1) = 0.656 μm, d(0.5) = 6.083 μm d(0.9) = 21.474 μm and D[4,3] =
8.75 μm. The higher D[4,3] value emphasises the presence of larger micelles, which
can clearly be seen in Figure 6.7. The shift in size distribution appears to be due to
the aggregation of smaller micelles. It is possible this is occurring as a function of the
emulsion beginning to break down. Alternatively the change in micelle size may be
related to the sodium caseinate content of the micelles, and what is observed is the
beginning of micelle coagulation (Mueller-Buschbaum et al. 2007). It has been
suggested previously that a lipase binding substrate is dependent on the organisation
of substrate within the interface and that the enzyme will only bind to clusters of
substrate molecules (micelles) of sufficient size (Wickham et al. 1998). The shift in
micelle size distribution to predominantly larger sizes fits with this theory, and it is
possible that the increased activity is linked to this process. However, this can only
be speculated as the exact micelle structure and composition in these emulsions has
not been defined. The minimum micelle size for lipase activity to commence is also
unknown. Furthermore, the effect of calcium on micelles and its influence on lipase
activity may be different with different lipid substrate/micelle mixtures.
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Figure 6.7 Weighted-mean of micelle size distribution in lecithin emulsions with and without
CaCl2. (A) Emulsion prepared with 1.2 g sodium caseinate, 0.5 g lecithin in 200 mL MQ H2O; (B)
emulsion prepared as in A, with CaCl2 added to a final concentration of 5 mM.
It is still unknown why the lipases with cleft-type active sites are more influenced by
the presence of calcium. It is possible that calcium interacts with the cleft-type active
site topology to allow interaction at the interface more readily. Alternatively the
calcium may be influencing the substrate interface itself, allowing for more
favourable interaction with the lipase. Another possible mechanism of action is that
the calcium is binding the released FFA as calcium soaps, thereby removing them as
a potential source of lipase inhibition (Alvarez & Stella 1989). However, it would be
expected that this effect would also be seen with all other lipase active sites; which it
0
1
2
3
4
5
6
0.01 0.1 1 10 100 1000
Volu
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(%)
Micelle size ( m)
0
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Volu
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Micelle size ( m)
A
B
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was not. In addition, Appendix 8.9, Figure 8.9 demonstrated that the addition of
calcium had an immediate effect on the hydrolysis rates of a lecithin emulsion with
very little FFA present, thus removal of inhibition is implausible. As has been shown
here, there are definite alterations to micelle surface chemistry when calcium is
added. It remains to be investigated if the lipases with cleft-type active themselves
are influenced directly as a result of calcium. As suggested by Alvarez et al. (1989),
a combination of both substrate and enzyme may be effected and act in a cooperative
fashion resulting in increased hydrolysis rates. The analysis of lipase dynamics at the
lipid interface is difficult to investigate experimentally and previous analyses by
other groups have used molecular dynamic simulations to investigate these
mechanisms (Peters & Bywater 2001). However, such simulations did not investigate
the influence of divalent cations and further work is required to fully understand the
mechanisms at play.
6.8 Summary
Research involving the development and improvement of lipase catalysed reactions
is important in commercial settings. Maximising lipase activity requires gaining a
better understanding of the biochemical mechanisms by which lipases function. From
analyses in Chapter V, it was noted that the activity of lipases with cleft-type active
sites (TlL and RmL) was stimulated in the presence of CaCl2 (but not by its
concentration). It was found that the increase in activity did not significantly alter the
FA selectivity of these lipases when tested against a range of pNP-acyl esters.
Further analysis revealed that the stimulation of hydrolysis by calcium was also
achieved in the hydrolysis of TAG. However, stimulation only occurred when
phospholipid was present in the emulsion. The physicochemical properties of a
lecithin emulsion containing TAG were modified in the presence of calcium. These
included a reduction in the surface charge and a shift in the size distribution of
micelles towards larger micelles. The exact mechanism of the observed stimulation is
still unclear. Further work that may shed light on this is outlined in Chapter VII.
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CHAPTER VII
Conclusions and future directions
Lipases are highly versatile biocatalysts and are applied extensively in commercial
lipid and chemical processing. The use of lipases for the concentration of LC n-3
PUFAs has received increasing interest in recent years, with varying levels of
success achieved using well characterised commercial lipases. The isolation of
lipases with selective activities for or against the hydrolysis of EPA and DHA, could
lead to more efficient concentration of LC n-3 PUFAs. Marine microalgae represent
an untapped source of lipases, as there is currently no published data outlining the
isolation and characterisation of these enzymes. Improved methods for the detection
and characterisation of lipases are also required to aid in the discovery of lipases with
novel activities.
The underlying hypothesis for this project was that by monitoring the formation and
depletion of lipids during the growth of LC n-3 PUFA-synthesising microalgae, it
might be possible to identify, and subsequently isolate, PUFA-specific lipases.
Despite expectations, it was found that examination of the lipid class and FA
composition of Chaetoceros calcitrans, Isochrysis sp. (T.ISO) and Pavlova lutheri
throughout their growth phases (batch culture), did not indicate when lipases were
likely to be present. Analysis of the lipid data from this study did however provide a
more in-depth investigation of lipid metabolism in these microalgal species
throughout batch culture, than has been previously reported. Total lipid content was
highest in P. lutheri and lowest in C. calcitrans. The LC n-3 PUFA content of each
species varied greatly. EPA was the primary LC n-3 PUFA present in C. calcitrans,
while in Isochrysis sp. it was DHA. P. lutheri contained both FAs at relatively
similar levels. The LC n-3 PUFA content of all three species was highest in early
culture; between lag and logarithmic phases. Determining the FA composition and
LC n-3 PUFA content of each species may be of benefit when using these organisms
as feed components in aquaculture.
The duration of growth, from initial culture inoculation until cultures entered
regression phase, varied greatly between species. C. calcitrans, Isochrysis sp. and P.
lutheri took 12, 17 and 72 days, respectively. C. calcitrans and Isochrysis sp. growth
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was similar to previously reported growth trials in the literature. The duration of P.
lutheri growth in this study was longer than any previously recorded examples.
Subsequent cultures were grown until the late logarithmic growth stage, to obtain
maximal biomass in the shortest possible time for lipase extraction. It was found that
the recovery of biomass was very low and as a result, the small amount of
protein/enzyme that could be recovered made analysis and purification challenging.
Future investigations into these lipases would benefit from use of a transcriptomic
approach.
Once the analysis of microalgal lipid composition was completed, investigation of
lipases in the three microalgae species was undertaken. Due to the low levels of
protein recovery from the biomass, it was important to release as much enzyme as
possible from the algal cells. The highest degree of cell lysis and protein release was
achieved by a bead-beating method. Once cells were lysed, lipase activity was
detected in the crude protein homogenates from Isochrysis sp. and P. lutheri. Several
lipid-hydrolysing bands were observed when gel-overlay assays were performed on
crude algal extracts separated by native-PAGE. This indicated that multiple lipases
may be present in these species. The activity of lipases from both species was found
to be calcium dependent. Efforts to concentrate and purify the lipases using well-
established methods such as ammonium sulfate precipitation and ion-exchange
chromatography did not result in lipase enrichment. Acidic native-PAGE identified a
lipid hydrolase with a pI >7.0. However, no significant binding was observed by
cation-exchange chromatography of homogenates from either species.
Lipase from P. lutheri was adsorbed onto hydrophobic interaction resins. The
varying hydrophobicities of butyl- and octyl-Sepharose resins appeared to separate
different lipolytic enzymes when the protein homogenate was applied first to the
butyl resin (less hydrophobic), then the supernatant applied to the octyl resin (more
hydrophobic). A one-step adsorption onto butyl-Toyopearl suggested a separation of
lipase from other lipid hydrolases (such as esterases), as specific hydrolysis carried
out by the supernatant against pNP-myristate (lipase) decreased more than for pNP-
butyrate (esterase). This indicated that lipase was bound more readily than other
lipolytic enzyme species. Measurement of lipase activity on the resin, or removal of
lipase from the resin was not achieved. However, it was established that hydrophobic
supports selectively bind lipases and would be useful for the isolation of lipases from
microalgae; if the issues involving enzyme concentration were overcome.
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Preliminary biochemical characterisation of crude microalgal protein homogenates
provided an indication of whether lipases from these species were worth pursuing in
the future. Temperature optima of 55 C and 30 C were defined for Isochrysis sp. and
P. lutheri extracts, respectively. The relatively high optimum found for lipase
activity in Isochrysis sp. may be useful for industrial applications where enzymes are
required to act at relatively high temperatures. Thermostability studies for the
purified or mixed enzymes are needed. These findings suggest that the pH optima for
lipases from both species are between pH 5.0 to 7.0, which are similar to the majority
of lipases sources from eukaryotic sources. Investigation of FA selectivity of the
crude extracts required the development of new colorimetric assay substrates with a
variety of FAs that included LC n-3 PUFAs. The activity profiles indicated that short
chain substrates (≤ C4) were hydrolysed most readily. This may have been due to the
presence of other lipid-hydrolases such as esterases. When C2:0 and C4:0 were
excluded from the profile, maximum activity was achieved against C14:0, which is
consistent with the medium chain-length FA selectivity of many other lipases. The
activity of both homogenates against the longer unsaturated substrates was
approximately 70-80% of that obtained for C14:0. This suggests that the enzymes
from these species are active against a broad array of FAs. It is possible that the
activity observed resulted from a combination of different enzymes. However, the
fact that pNP-EPA and DHA were hydrolysed at relatively high levels is promising
for future studies, once the enzymes have been separated.
To resolve the major difficulties encountered in this part of the project, such as the
low amount of activity present, future efforts to isolate microalgal lipases will need
to be via a different approach. One solution is the use of recombinant expression
systems. While this approach is limited to species that currently have annotated
sequence data available, recombinant expression has significant advantages over
native isolation protocols. Improved yields of target protein and more
directed/specific purification schemes are two likely benefits. Recent work on
recombinantly expressing lipid hydrolases from microalgae has demonstrated the
move towards this approach (Godet et al. 2010; Kerviel et al. 2014). So far, this work
has used bacterial expression hosts. However, these organisms may not carry out
post-translational modifications satisfactorily. A more attractive methodology to
follow could involve using eukaryotic systems such as yeast, or microalgae
themselves.
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155
The second major area of this project involved extending the number of substrates
available for use in pNP assay, and investigating if this method could be used in
defining lipase FA selectivity. Methods for comprehensive biochemical
characterisation are relatively slow and therefore represent a bottleneck in the
progress of obtaining a full understanding of protein function. Consequently, the gap
between in silico data and data derived experimentally is growing and methods that
can be performed in a high-throughput fashion are of interest. The pNP activity assay
provided a well-established assay platform to build on. The synthesis of novel
olefinic pNP-acyl esters (pNP-C16:1 n-7, pNP-C18:1 n-9, pNP-C18:2 n-6, pNP-
C18:3 n-3, pNP-C20:5 n-3, pNP-C22:6 n-3), particularly LC n-3 PUFA derivatives,
extended the commercially available saturated substrates to provide a tool for the
rapid screening/profiling of lipase FA selectivity.
The methods for pNP-acyl ester synthesis and isolation provided yields in excess
85% and are simpler than those previously reported. The compounds were
successfully chemically characterised by UV spectroscopy, mass spectrometry,
ATR-FTIR spectroscopy and 1H and 13C NMR. All synthesised pNP-acyl esters were
found to be stable under assay and storage conditions for at least 6 months. The FA
selectivity of five commercial lipases (lipase B from Candida antarctica (CalB),
Aspergillus niger lipase (AnL), Thermomyces lanuginosus lipase (TlL), Rhizomucor
miehei lipase (RmL) and Candida rugosa lipase (CrL)) was probed using the newly
synthesised pNP derivatives in conjunction with commercial short and medium chain
pNP substrates. Results indicated that CalB and AnL have similar FA selectivity
profiles, possessing activity for a relatively broad range of substrates. Likewise TIL
and RmL have similar selectivity, with saturated FAs of C8 in length hydrolysed
much more rapidly than FAs containing double bonds and soluble short chain
substrates. CrL showed the most unique profile with best hydrolysis achieved against
pNP-C12:0, pNP-C16:1 n-7 and pNP-C18:3 n-3. This lipase also showed omega-3
selectivity, particularly toward the retention of DHA, indicating that CrL may be the
most appropriate of the screened lipases for concentrating LC n-3 PUFAs. Since
these FAs were discriminated against, they may be preferentially retained in
acylglycerol substrates, whilst other FAs are preferentially removed.
These findings demonstrate that the new comprehensive screening system with pNP-
acyl esters can be used to determine FA selectivity for lipases. Analyses of these
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156
lipases by other researchers with TAG substrates, indicate that the pNP-defined FA
selectivity correlates well to that seen with more natural substrates (Akanbi et al.
(2013), personal communication, Dr. Matthew Miller, Plant and Food Research, New
Zealand). Future developments to enhance the usefulness of the assay system should
include the following: (1) The extension of the ester library with other FA derivatives
such as LC n-6 PUFAs. (2) Conversion of the assay protocol to a 96-well plate
format to allow higher throughput analyses. (3) The investigation of alternative
labels that can increase sensitivity, such as fluorogenic methylumbelliferone
derivatives.
Understanding the physicochemical mechanisms of lipases enables the enzymes to
be utilised effectively for different applications. While the general mechanism of
lipase activity is defined, the mechanisms involving the modification of activity are
less well understood. The work undertaken here investigated the influence of calcium
on the activity of commercial lipases with different active site topologies on a range
of emulsified substrates. Using pNP-acyl esters, it was found that the activity of
some fungal lipases was stimulated by calcium. This was correlated with the active
site conformation of the lipases. Lipases with cleft-type active sites (TlL and RmL)
were stimulated, while tunnel-type (CalB and AnL) and funnel-type (CrL) topologies
showed no enhancement. Significant increases in hydrolysis rates were also observed
when calcium was added to tributyrin emulsions, with hydrolysis by TlL and RmL
increasing 170 and 32 times, respectively. This was not observed for lipases with
other active site topologies. However, emulsions containing both TAG and gum
arabic (as the emulsifier) were not affected by calcium, regardless of the active site
structure. Additional investigation demonstrated that the stimulation of activity was
only observed against emulsified substrates containing both TAG and phospholipid.
Analysis by Iatroscan demonstrated that TAG was the primary hydrolysis target and
not phospholipid. Further analysis of the physicochemical properties of the lecithin
(TAG and phospholipid) emulsions demonstrated that several physicochemical
effects were caused by the calcium. Firstly, the surface charge (zeta potential) of
micelles in emulsions containing calcium was decreased. Secondly, calcium caused a
shift in micelle size distribution towards larger micelles. It is unclear if these
modifications to substrate properties are the cause of increased activity by lipases
with cleft-type active sites, or if the calcium is influencing the lipase itself. The fact
that the calcium-stimulation is specific to a particular active site topology indicates
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157
that there is something unique in how this type of active site interacts with lipid at
the lipid-water interface. Calcium removing FFAs as calcium soaps is unlikely as this
mechanism could potentially occur with all lipases and yet it does not. Furthermore,
the addition of calcium during the reaction was found to have an immediate impact
on activity. Therefore, calcium either alters the lipid interface allowing lipases with
cleft-type active sites to interact more readily, or calcium is influencing the catalytic
mechanism itself.
Further experiments are required to determine if one or perhaps a combination of
these explanations is responsible. Future work should involve the analysis of lipase
activity against emulsions with more controlled lipid compositions, including the use
of a more pure phospholipid source so that substrate ratios (TAG/phospholipid) can
be better controlled. It would also be useful to investigate whether or not this
phenomenon is unique to calcium or whether other divalent cations, such as
magnesium, act in the same way. In addition to investigating the emulsion properties,
efforts also need to address changes to the enzyme itself.
This research project has investigated the lipid class and FA composition of three
microalgae over the duration of their growth period. This has provided an in-depth
picture of how the lipid in these organisms is utilised under batch culture conditions.
The analysis of lipases from these species has also been presented. Although the
isolation of these enzymes was challenging and their specificity towards LC n-3
PUFA unclear, preliminary data provided by this work has indicated a number of
useful properties possessed by the microalgal lipases. With the aid of recombinant
protein production in the future, the methods described in this work will be useful for
isolation and characterisation of microalgal lipases. The synthesis and application of
novel assay substrates has extended the capabilities of the pNP assay. It now
provides a practical and convenient tool for profiling lipase FA selectivity that can be
carried out in a relatively high-throughput fashion. The use of this method in
screening for lipases with LC n-3 PUFA-relevant activities is an exciting application.
The analyses of how lipases interact with different lipid substrates and the factors
(such as calcium) that influence the process are important. The uses of these enzymes
in different commercial applications will increase as we gain a more comprehensive
understanding of their function and the variables that affect it.
BIBLIOGRAPHY
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VIII APPENDICES
178
VIII
APPENDICES
Appendix 8.1 Fatty acid profiles of Chaetoceros calcitrans,
Isochrysis sp. and Pavlova lutheri and their lipid
classes
FA (%
) E
L
L
LL
E
S M
S L
S Σ
satu
rate
s 52
.4 (±
0.9
) 20
.4 (±
1.7
) 17
.3 (±
0.1
) 16
.5 (±
0.3
) 18
.5 (±
1.8
) 33
.5 (±
6.0
)
C14
:0
32.2
(± 1
.4)
13.4
(± 0
.8)
13.2
(± 0
.1)
12.0
(± 0
.1)
12.3
(± 0
.4)
13.0
(± 0
.3)
C15
:0
1.3
(± 0
.0)
n.d.
n.
d.
n.d.
n.
d.
0.6
(± 0
.0)
C16
:0
16.4
(± 0
.8)
5.2
(± 0
.4)
4.0
(± 0
.1)
4.5
(± 0
.2)
6.2
(± 1
.5)
19.6
(± 6
.2)
C18
:0
2.5
(± 0
.7)
1.8
(± 1
.0)
n.d.
n.
d.
n.d.
0.
3 (±
0.1
)
Σ m
onou
nsat
urat
es
5.1
(± 3
.3)
28.9
(± 1
.9)
31.6
(± 0
.2)
31.1
(± 0
.8)
28.6
(± 0
.7)
26.0
(± 1
.2)
C16
:1n-
7 5.
1 (±
3.3
) 27
.6 (±
2.1
) 30
.6 (±
0.1
) 30
.0 (±
0.9
) 28
.6 (±
0.7
) 25
.6 (±
1.6
)
C17
:1
n.d.
1.
3 (±
0.1
) 1.
0 (±
0.1
) 1.
1 (±
0.9
) n.
d.
n.d.
C
18:1
n-9
n.d.
n.
d.
n.d.
n.
d.
n.d.
0.
4 (±
0.6
) Σ
poly
unsa
tura
tes
42.5
(± 2
.4)
50.7
(± 2
.8)
51.2
(± 0
.2)
52.4
(± 1
.0)
53.0
(± 2
.0)
40.5
(± 5
.2)
C16
:2n-
6 3.
5 (±
0.8
) 6.
3 (±
0.2
) 7.
4 (±
0.1
) 8.
7 (±
0.5
) 8.
4 (±
0.8
) 5.
5 (±
0.8
)
C18
:2n-
6 n.
d.
n.d.
0.
6 (±
0.5
) 0.
5 (±
0.4
) 0.
8 (±
0.1
) 1.
0 (±
0.5
)
C20
:4n-
6 n.
d.
n.d.
n.
d.
n.d.
n.
d.
0.4
(±0.
0)
C22
:2n-
6 19
.5 (±
3.6
) n.
d.
n.d.
n.
d.
n.d.
n.
d.
C16
:2n-
4 n.
d.
3.8
(± 0
.1)
4.1
(± 0
.2)
4.2
(± 0
.4)
5.3
(± 0
.2)
5.0
(± 0
.9)
C16
:3n-
4 2.
4 (±
2.2
) 13
.3 (±
1.1
) 12
.7 (±
0.2
) 14
.5 (±
0.2
) 14
.5 (±
0.6
) 9.
8 (±
1.8
)
C18
:3n-
3 n.
d.
n.d.
1.
0 (±
0.1
) 1.
0 (±
0.0
) 1.
3 (±
0.1
) 1.
3 (±
0.1
)
C18
:4n-
3 n.
d.
1.1
(± 0
.2)
n.d.
0.
4 (±
0.3
) n.
d.
0.4
(± 0
.0)
C20
:5n-
3 1.
4 (±
1.3
) 23
.0 (±
1.8
) 23
.1 (±
0.2
) 21
.5 (±
0.2
) 20
.9 (±
0.8
) 15
.8 (±
2.0
)
C22
:6n-
3 15
.7 (±
2.2
) 2.
3 (±
0.5
) 1.
4 (±
0.1
) 1.
0 (±
0.1
) 1.
0 (±
0.1
) 1.
0 (±
0.1
)
C16
:4n-
1 n.
d.
0.8
(± 0
.1)
0.9
(± 0
.0)
0.7
(± 0
.0)
0.7
(± 0
.0)
0.4
(± 0
.0)
Tabl
e 8.
1.1
Cha
nges
in
to
tal
lipid
fa
tty
acid
co
mpo
sitio
n (%
) of
C
haet
ocer
os
calc
itran
s ov
er
grow
th.
Early
loga
rithm
ic (
EL),
loga
rithm
ic (
L), l
ate
loga
rithm
ic (
LL),
early
sta
tiona
ry (
ES),
stat
iona
ry (
S) a
nd la
te s
tatio
nary
(LS
) ph
ase.
Res
ults
pre
sent
ed a
s mea
n va
lues
(± S
D; n
=3),
n.d.
: not
det
ecte
d.
VIII APPENDICES
179
FA (%
) E
L
L
LL
E
S M
S L
S Σ
satu
rate
s51
.1 (±
5.3
) 28
.2 (±
4.9
) 22
.2 (±
1.0
) 14
.1 (±
2.6
) 15
.0 (±
2.4
) 36
.7 (±
5.8
)
C14
:0
25.0
(± 0
.3)
17.0
(± 2
.8)
15.4
(± 1
.0)
6.0
(± 2
.2)
8.0
(± 0
.8)
11.6
(± 0
.2)
C15
:0
n.d.
0.
7 (±
0.2
) 0.
4 (±
0.3
) n.
d.
n.d.
0.
6 (±
0.0
)
C16
:0
17.7
(± 5
.2)
7.0
(± 1
.4)
5.2
(± 0
.2)
5.7
(± 0
.3)
6.2
(± 1
.9)
23.8
(± 5
.8)
C18
:0
8.4
(± 1
.1)
3.5
(± 1
.5)
1.2
(± 0
.1)
2.4
(± 0
.3)
0.9
(± 0
.3)
0.8
(± 0
.2)
Σ m
onou
nsat
urat
es
10.4
(± 6
.5)
22.6
(± 1
.0)
23.3
(± 0
.4)
23.8
(± 0
.1)
25.3
(± 1
.2)
27.2
(± 2
.6)
C16
:1n-
7 10
.4 (±
6.5
) 22
.2 (±
1.3
) 23
.3 (±
0.4
) 18
.2 (±
4.8
) 25
.3 (±
1.2
) 27
.7 (±
2.6
)
C17
:1
n.d.
0.
3 (±
0.3
) n.
d.
n.d.
n.
d.
n.d.
C
18:1
n-9
n.d.
n.
d.
n.d.
5.
6 (±
5.0
) n.
d.
n.d.
Σ
poly
unsa
tura
tes
38.5
(± 1
.2)
49.2
(± 5
.9)
54.5
(± 1
.3)
62.1
(± 2
.5)
59.7
(± 3
.6)
35.6
(± 3
.4)
C16
:2n-
6 8.
3 (±
1.8
) 6.
5 (±
0.2
) 7.
8 (±
0.7
) 12
.2 (±
4.6
) 11
.5 (±
3.0
) 5.
5 (±
1.2
)
C18
:2n-
6 n.
d.
0.7
(± 0
.2)
1.3
(± 0
.1)
16.1
(± 1
2.3)
1.
5 (±
0.1
) 1.
2 (±
0.6
)
C20
:4n-
6 n.
d.
n.d.
0.
8 (±
0.0
.) n.
d.
0.4
(± 0
.4)
0.4
(± 0
.0)
C22
:2n-
6 10
.4 (±
7.8
) n.
d.
n.d.
n.
d.
n.d.
n.
d.
C16
:2n-
4 n.
d.
2.7
(± 0
.4)
3.1
(± 0
.1)
2.1
(± 0
.5)
3.8
(± 0
.3)
3.6
(± 0
.5)
C16
:3n-
4 2.
5 (±
2.2
) 4.
4 (±
0.8
) 4.
0 (±
0.2
) 4.
4 (±
0.2
) 7.
1 (±
0.5
) 5.
6 (±
0.5
)
C18
:3n-
3 n.
d.
0.3
(± 0
.3)
1.4
(± 0
.1)
0.6
(± 0
.6)
1.8
(± 0
.1)
1.3
(± 0
.1)
C18
:4n-
3 n.
d.
0.4
(± 0
.4)
n.d.
n.
d.
n.d.
0.
2 (±
0.2
) C
20:5
n-3
7.5(
±1.
8)
31.2
(± 4
.5)
33.4
(± 0
.8)
24.1
(± 4
.5)
30.3
(± 0
.3)
16.2
(± 1
.7)
C22
:6n-
3 9.
9 (±
5.3
) 1.
1 (±
0.2
) 1.
0 (±
0.1
) n.
d.
1.1
(± 0
.0)
0.8
(± 0
.1)
C16
:4n-
1 n.
d.
1.9
(± 0
.2)
1.8
(± 0
.1)
2.5
(± 1
.0)
2.3
(± 0
.7)
0.7
(± 0
.1)
Tabl
e 8.
1.2
Cha
nges
in
ne
utra
l lip
id
fatty
ac
id
com
posi
tion
(%)
of
Cha
etoc
eros
ca
lcitr
ans
over
gr
owth
. Ea
rly lo
garit
hmic
(EL
), lo
garit
hmic
(L)
, lat
e lo
garit
hmic
(LL
), ea
rly s
tatio
nary
(ES
), st
atio
nary
(S)
and
late
sta
tiona
ry (
LS)
phas
e. R
esul
ts p
rese
nted
as m
ean
valu
es (±
SD
; n=3
), n.
d.: n
ot d
etec
ted.
VIII APPENDICES
180
FA (%
) E
L
L
LL
E
S M
S L
S Σ
satu
rate
s 24
.6 (±
4.1)
14.4
(±1.
0)
15.1
(±0.
4)
17.6
(±1.
7)
19.4
(±2.
4)
28.9
(±6.
1)
C14
:0
17.1
(± 2
.6)
10.6
(± 0
.7)
11.8
(± 0
.3)
12.5
(± 1
.7)
13.4
(± 1
.2)
13.4
(± 0
.6)
C15
:0
n.d.
n.
d.
n.d.
0.
3 (±
0.3
) 0.
5 (±
0.1
) 0.
6 (±
0.0
)
C16
:0
7.5
(± 1
.6)
3.8
(± 0
.4)
3.3
(± 0
.2)
4.2
(± 0
.2)
5.5
(± 1
.2)
14.2
(± 5
.3)
C18
:0
n.d.
n.
d.
n.d.
0.
6 (±
0.1
) n.
d.
0.7
(± 0
.2)
Σ m
onou
nsat
urat
es
26.0
(± 8
.4)
30.3
(± 1
.8)
33.4
(± 0
.4)
32.9
(± 4
.4)
27.1
(± 1
.0)
21.3
(± 0
.8)
C16
:1n-
7 26
.0 (±
8.4
) 28
.2 (±
2.2
) 31
.7 (±
0.4
) 30
.7 (±
6.3
) 27
.1 (±
1.0
) 21
.3 (±
0.8
)
C17
:1
n.d.
2.
2 (±
0.3
) 1.
7 (±
0.1
) 1.
3 (±
1.1
) n.
d.
n.d.
C
18:1
n-9
n.d.
n.
d.
n.d.
0.
9 (±
0.9
) n.
d.
n.d.
Σ
poly
unsa
tura
tes
48.8
(± 8
.7)
55.3
(± 2
.8)
51.5
(± 0
.4)
49.5
(± 6
.1)
53.4
(± 2
.1)
49.7
(± 5
.9)
C16
:2n-
6 6.
1 (±
1.0
) 5.
9 (±
0.4
) 6.
6 (±
0.4
) 6.
9 (±
1.3
) 6.
9 (±
0.3
) 5.
4 (±
0.3
)
C18
:2n-
6 n.
d.
n.d.
n.
d.
2.9
(± 1
.9)
0.6
(± 0
.1)
0.7
(± 0
.3)
C20
:4n-
6 n.
d.
n.d.
n.
d.
n.d.
n.
d.
0.3
(± 0
.2)
C22
:2n-
6 4.
9 (±
3.7
) n.
d.
n.d.
n.
d.
n.d.
n.
d.
C16
:2n-
4 2.
8 (±
2.6
) 5.
0 (±
0.2
) 5.
2 (±
0.3
) 5.
4 (±
0.9
) 6.
5 (±
0.2
) 7.
3 (±
1.0
)
C16
:3n-
4 9.
8 (±
9.4
) 24
.7 (±
3.2
) 23
.7 (±
0.4
) 21
.3 (±
3.2
) 21
.0 (±
0.7
) 17
.7 (±
1.7
)
C18
:3n-
3 n.
d.
n.d.
n.
d.
0.9
(± 0
.2)
1.0
(± 0
.2)
1.2
(± 0
.2)
C18
:4n-
3 n.
d.
1.1
(± 0
.3)
0.8
(± 0
.0)
0.6
(± 0
.1)
0.5
(± 0
.0)
0.4
(± 0
.0)
C20
:5n-
3 13
.8 (±
4.6
) 16
.6 (±
0.4
) 14
.6 (±
0.2
) 11
.4 (±
9.8
) 16
.6 (±
2.3
) 16
.4 (±
2.8
)
C22
:6n-
3 11
.4 (±
4.6
) 2.
0 (±
0.9
) 0.
5 (±
0.5
) n.
d.
0.2
(± 0
.2)
0.3
(± 0
.3)
Tabl
e 8.
1.3
Cha
nges
in
gl
ycol
ipid
fa
tty
acid
co
mpo
sitio
n (%
) of
C
haet
ocer
os
calc
itran
s ov
er
grow
th.
Early
loga
rithm
ic (
EL),
loga
rithm
ic (
L), l
ate
loga
rithm
ic (
LL),
early
sta
tiona
ry (
ES),
stat
iona
ry (
S) a
nd la
te s
tatio
nary
(LS
) ph
ase.
Res
ults
pre
sent
ed a
s mea
n va
lues
(± S
D; n
=3),
n.d.
: not
det
ecte
d.
VIII APPENDICES
181
FA (%
) E
L
L
LL
E
S M
S L
S Σ
satu
rate
s 52
.1 (±
2.7
) 13
.8 (±
3.0
) 17
.1 (±
0.9
) 29
.8 (±
2.4
) 28
.6 (±
0.2
) 36
.1 (±
4.4
)
C14
:0
32.1
(± 5
.8)
8.8
(± 2
.1)
11.8
(± 0
.7)
19.8
(± 0
.8)
17.6
(± 1
.6)
18.6
(± 1
.2)
C16
:0
20.0
(± 5
.7)
4.9
(± 1
.0)
5.4
(± 0
.3)
10.0
(± 1
.6)
11.0
(± 1
.7)
17.5
(± 3
.2)
Σ m
onou
nsat
urat
es
n.d.
45
.7 (±
0.3
) 49
.7 (±
0.6
) 39
.8 (±
0.6
) 40
.6 (±
1.0
) 30
.9 (±
1.6
)
C16
:1n-
7 n.
d.
45.7
(± 0
.3)
49.7
(± 0
.6)
39.8
(± 0
.6)
40.6
(± 1
.0)
29.0
(± 3
.2)
C18
:1n-
9 n.
d.
n.d.
n.
d.
n.d.
n.
d.
1.9
(± 2
.1)
Σ po
lyun
satu
rate
s 47
.9 (±
2.7
) 40
.6 (±
3.1
) 33
.2 (±
0.6
) 30
.4 (±
2.5
) 30
.8 (±
1.1
) 33
.0 (±
3.4
)
C16
:2n-
6 n.
d.
6.4
(± 0
.7)
6.4
(± 0
.3)
5.8
(± 0
.4)
5.7
(± 0
.2)
5.3
(± 0
.5)
C18
:2n-
6 n.
d.
n.d.
2.
3 (±
0.2
) 3.
1 (±
1.1
) 2.
0 (±
0.2
) 2.
0 (±
0.4
)
C22
:2n-
6 27
.0 (±
1.7
) n.
d.
n.d.
n.
d.
n.d.
n.
d.
C16
:2n-
4 n.
d.
3.1
(± 0
.3)
3.2
(± 0
.0)
1.6
(± 1
.4)
3.1
(± 0
.2)
3.2
(± 0
.4)
C16
:3n-
4 n.
d.
3.2
(± 0
.5)
2.9
(± 0
.1)
3.1
(± 0
.3)
3.0
(± 0
.2)
4.2
(± 0
.4)
C18
:3n-
3 n.
d.
n.d.
n.
d.
n.d.
0.
8 (±
0.7
) 0.
8 (±
0.7
) C
18:4
n-3
n.d.
1.
8 (±
1.5
) n.
d.
n.d.
n.
d.
n.d.
C
20:5
n-3
n.d.
17
.0 (±
0.5
) 12
.6 (±
0.4
) 14
.9 (±
2.4
) 12
.9 (±
1.6
) 13
.1 (±
2.3
)
C22
:6n-
3 20
.9 (±
1.0
) 9.
0 (±
0.1
) 5.
7 (±
0.5
) 1.
9 (±
1.6
) 3.
3 (±
0.4
) 4.
3 (±
0.4
)
Tabl
e 8.
1.4
Cha
nges
in
po
lar
lipid
fa
tty
acid
co
mpo
sitio
n (%
) of
C
haet
ocer
os
clac
itran
s ov
er
grow
th.
Early
loga
rithm
ic (
EL),
loga
rithm
ic (
L), l
ate
loga
rithm
ic (
LL),
early
sta
tiona
ry (
ES),
stat
iona
ry (
S) a
nd la
te s
tatio
nary
(LS
) ph
ase.
Res
ults
pre
sent
ed a
s mea
n va
lues
(± S
D; n
=3),
n.d.
: not
det
ecte
d.
VIII APPENDICES
182
FA (%
) L
A
EL
L
L
L
ES
MS
LS
Σ sa
tura
tes
44.0
(± 1
.4)
41.1
(± 0
.5)
32.3
(± 0
.4)
33.9
(± 1
.1)
36.4
(± 3
.5)
37.5
(± 0
.2)
38.9
(± 0
.7)
C14
21
.8 (±
0.3
) 21
.9 (±
0.2
) 19
.3 (±
0.2
) 19
.2 (±
0.9
) 21
.3 (±
2.4
) 21
.5 (±
0.3
) 22
.1 (±
0.8
)
C15
n.
d.
0.3
(± 0
.0)
0.2
(± 0
.2)
n.d.
n.
d.
n.d.
n.
d.
C16
17
.9 (±
0.4
) 16
.4 (±
0.2
) 12
.4 (±
0.3
) 14
.8 (±
0.8
) 13
.8 (±
1.1
) 14
.7 (±
0.3
) 15
.4 (±
0.2
)
C18
4.
0 (±
0.6
) 2.
1 (±
0.2
) 0.
4 (±
0.4
) n.
d.
1.3
(± 0
.1)
1.2
(± 0
.1)
1.1
(± 0
.1)
C22
0.
4 (±
0.3
) 0.
4 (±
0.1
) n.
d.
n.d.
n.
d.
n.d.
0.
3 (±
0.3
)
Σ m
onou
nsat
urat
es
29.8
(± 0
.2)
26.3
(± 0
.6)
24.7
(± 0
.5)
28.6
(± 1
.5)
26.4
(± 0
.7)
29.6
(± 0
.7)
30.8
(± 0
.4)
C14
:1
n.d.
n.
d.
0.2
(± 0
.2)
n.d.
n.
d.
n.d.
n.
d.
C16
:1n-
7 1.
1 (±
0.2
) 1.
9 (±
0.2
) 4.
4 (±
0.2
) 3.
7 (±
0.3
) 2.
8 (±
0.4
) 2.
5 (±
0.4
)2.
5 (±
0.1
)
C18
:1
28.7
(± 0
.2)
24.5
(± 0
.6)
20.2
(± 0
.7)
24.9
(± 1
.8)
23.6
(± 1
.0)
27.1
(± 1
.0)
28.2
(± 0
.5)
Σ po
lyun
satu
rate
s 26
.1 (±
1.3
) 32
.6 (±
0.7
) 43
.0 (±
0.6
) 37
.4 (±
1.9
) 37
.2 (±
3.9
) 32
.9 (±
0.5
) 30
.4 (±
0.6
)
C16
:2n-
6 0.
7 (±
0.0
) 1.
1 (±
0.2
) 1.
9 (±
0.1
) 1.
6 (±
0.2
) 0.
9 (±
0.2
) 0.
7 (±
0.1
)0.
4 (±
0.3
)
C18
:2n-
6 5.
3 (±
0.1
) 5.
2 (±
0.1
) 9.
5 (±
0.4
) 8.
8 (±
1.2
) 6.
8 (±
1.9
) 6.
0 (±
0.7
)5.
5 (±
0.2
)
C18
:3n-
6 n.
d.
0.4
(± 0
.1)
1.0
(± 0
.4)
0.9
(± 0
.0)
0.9
(± 0
.3)
0.5
(± 0
.5)
0.3
(± 0
.3)
C20
:4n-
6 0.
6 (±
0.2
) 0.
4 (±
0.1
) 0.
3 (±
0.2
) n.
d.
n.d.
n.
d.
n.d.
C
16:2
n-4
0.7
(± 0
.3)
0.4
(± 0
.0)
n.d.
n.
d.
n.d.
n.
d.
n.d.
C
18:3
n-3
2.9
(± 0
.6)
4.4
(± 0
.8)
6.8
(± 0
.3)
6.6
(± 0
.8)
5.3
(± 1
.1)
4.3
(± 0
.4)
3.8
(± 0
.3)
C18
:4n-
3 7.
2 (±
0.2
) 10
.2 (±
0.2
) 11
.9 (±
0.1
) 10
.4 (±
0.4
) 12
.7 (±
0.4
) 12
.2 (±
0.2
) 11
.5 (±
0.2
)
C20
:5n-
3 0.
3 (±
0.2
) 0.
5 (±
0.0
) 0.
6 (±
0.0
) n.
d.
n.d.
n.
d.
n.d.
C
22:6
n-3
8.1
(± 0
.7)
9.5
(± 0
.2)
10.3
(± 0
.1)
9.1
(± 0
.6)
10.2
(± 1
.0)
8.5
(± 1
.3)
8.5
(± 1
.1)
C16
:4n-
1 0.
3 (±
0.3
) 0.
6 (±
0.0
) 0.
7 (±
0.0
) n.
d.
0.4
(± 0
.3)
0.5
(± 0
.1)
0.4
(± 0
.0)
Tabl
e 8.
1.5
Cha
nges
in
to
tal
lipid
fa
tty
acid
co
mpo
sitio
n (%
) of
Is
ochr
ysis
sp
. ov
er
grow
th.
Lag
(LA
), ea
rly l
ogar
ithm
ic (
EL),
loga
rithm
ic (
L),
late
log
arith
mic
(LL
), ea
rly s
tatio
nary
(ES
), st
atio
nary
(S)
and
lat
e st
atio
nary
(LS)
pha
se. R
esul
ts p
rese
nted
as m
ean
valu
es (±
SD
; n=3
), n.
d.: n
ot d
etec
ted.
VIII APPENDICES
183
FA (%
) L
A
EL
L
L
L
ES
MS
LS
Σ sa
tura
tes
45.5
(± 0
.8)
42.5
(± 0
.4)
32.4
(± 1
.2)
36.7
(± 1
.0)
38.7
(± 6
.7)
36.8
(± 5
.7)
43.4
(± 1
.3)
C14
22
.4 (±
0.2
) 22
.2 (±
0.2
) 18
.0 (±
0.3
) 18
.8 (±
0.8
) 23
.6 (±
5.4
) 21
.8 (±
4.7
) 26
.9 (±
0.6
)
C15
0.
3 (±
0.0
) 0.
3 (±
0.0
) 0.
2 (±
0.2
) n.
d.
0.4
(± 0
.1)
0.3
(± 0
.0)
0.4
(± 0
.0)
C16
19
.1 (±
0.4
) 17
.9 (±
0.1
) 12
.3 (±
0.8
) 15
.7 (±
0.9
) 13
.0 (±
1.9
) 13
.2 (±
1.1
) 14
.7 (±
0.5
)
C17
0.
2 (±
0.2
) 0.
4 (±
0.0
) n.
d.
n.d.
n.
d.
n.d.
n.
d.
C18
4.
9 (±
0.7
) 3.
4 (±
0.3
) 1.
6 (±
0.2
) 1.
4 (±
0.0
) 1.
0 (±
0.1
) 0.
9 (±
0.1
) 0.
6 (±
0.1
)
C20
0.
2 (±
0.0
) n.
d.
n.d.
n.
d.
0.1
(± 0
.1)
0.1
(± 0
.1)
0.2
(± 0
.1)
C22
0.
6 (±
0.1
) 0.
6 (±
0.1
) 0.
3 (±
0.3
) 0.
8 (±
0.2
) 0.
6 (±
0.3
) 0.
5 (±
0.1
) 0.
5 (±
0.1
)
Σ m
onou
nsat
urat
es
37.5
(± 0
.9)
37.1
(± 0
.9)
32.2
(± 1
.7)
37.3
(± 2
.1)
35.2
(± 1
.9)
35.0
(± 1
.4)
33.5
(± 0
.4)
C14
:1
n.d.
n.
d.
n.d.
n.
d.
0.3
(± 0
.0)
0.3
(± 0
.0)
0.3
(± 0
.0)
C16
:1n-
70.
6 (±
0.1
) 1.
0 (±
0.1
) 2.
7 (±
0.1
) 2.
1 (±
0.3
) 2.
1 (±
0.1
) 2.
1 (±
0.1
) 2.
1 (±
0.1
)
C17
:1n.
d.n.
d.n.
d.n.
d.0.
4 (±
0.1
)n.
d.0.
3 (±
0.0
)C
18:1
36
.2 (±
0.7
) 35
.2 (±
0.5
) 28
.8 (±
1.6
) 34
.4 (±
2.3
) 31
.6 (±
1.6
) 32
.0 (±
1.2
) 30
.2 (±
0.3
)
C22
:1
0.2
(± 0
.0)
0.3
(± 0
.0)
0.7
(± 0
.1)
0.8
(± 0
.0)
0.7
(± 0
.4)
0.6
(± 0
.1)
0.6
(± 0
.0)
Σ po
lyun
satu
rate
s 17
.0 (±
0.6
) 20
.4 (±
0.9
) 35
.4 (±
2.9
) 25
.9 (±
2.8
) 26
.1 (±
5.1
) 28
.2 (±
4.3
) 23
.1 (±
1.1
)
C16
:2n-
60.
7 (±
0.0
) 1.
3 (±
0.3
) 5.
0 (±
0.8
) 3.
0 (±
0.5
) 1.
8 (±
0.4
) 1.
5 (±
0.4
) 1.
3 (±
0.2
)
C18
:2n-
66.
2 (±
0.1
) 6.
6 (±
1.0
) 8.
3 (±
0.0
) 6.
6 (±
0.8
) 6.
6 (±
1.3
) 6.
2 (±
0.9
) 5.
3 (±
0.4
)
C18
:3n-
60.
1 (±
0.1
) 0.
2 (±
0.1
) 0.
3 (±
0.2
) n.
d.
0.5
(± 0
.2)
0.4
(± 0
.2)
0.3
(± 0
.1)
C20
:2n-
6n.
d.
n.d.
n.
d.
n.d.
n.
d.
0.1
(± 0
.1)
n.d.
C
20:4
n-6
0.4
(± 0
.1)
0.3
(± 0
.0)
n.d.
n.
d.
0.2
(± 0
.1)
0.2
(± 0
.0)
0.1
(± 0
.1)
C16
:2n-
4n.
d.
n.d.
n.
d.
n.d.
n.
d.
0.3
(± 0
.2)
0.7
(± 0
.3)
C18
:3n-
31.
0 (±
0.3
) 1.
3 (±
0.5
) 3.
8 (±
0.5
) 2.
8 (±
0.5
) 2.
7 (±
0.9
) 3.
1 (±
0.7
) 2.
5 (±
0.1
)
C18
:4n-
34.
1 (±
0.2
) 4.
9 (±
0.2
) 6.
0 (±
0.3
) 4.
3 (±
0.1
) 7.
3 (±
0.3
) 8.
7 (±
1.2
) 7.
5 (±
0.1
)
C20
:5n-
30.
3 (±
0.0
) 0.
3 (±
0.1
) 0.
2 (±
0.2
) n.
d.
0.3
(± 0
.1)
0.4
(± 0
.0)
0.3
(± 0
.0)
C22
:6n-
33.
2 (±
0.6
) 5.
1 (±
0.4
) 9.
6 (±
1.4
) 7.
9 (±
0.9
) 5.
3 (±
2.2
) 6.
2 (±
1.5
) 4.
2 (±
0.2
)
C16
:4n-
10.
8 (±
0.0)
1.
1 (±
0.0
) 2.
2 (±
0.2
) 1.
4 (±
0.2
) 1.
4 (±
0.2
) 1.
1 (±
0.1
) 0.
8 (±
0.1
)
Tabl
e 8.
1.6
Cha
nges
in
ne
utra
l lip
id
fatty
ac
id
com
posi
tion
(%)
of
Isoc
hrys
is
sp.
over
gr
owth
. La
g (L
A),
early
log
arith
mic
(EL
), lo
garit
hmic
(L)
, la
te l
ogar
ithm
ic (
LL),
early
sta
tiona
ry (
ES),
stat
iona
ry (
S) a
nd l
ate
stat
iona
ry (L
S) p
hase
. Res
ults
pre
sent
ed a
s mea
n va
lues
(± S
D; n
=3),
n.d.
: not
det
ecte
d.
VIII APPENDICES
184
FA (%
) L
A
EL
L
L
L
ES
MS
LS
Σ sa
tura
tes
42.7
(± 2
.1)
39.2
(± 0
.7)
31.8
(± 0
.2)
31.7
(± 0
.8)
29.1
(± 3
.9)
28.9
(± 4
.0)
32.0
(± 0
.6)
C14
21
.8 (±
0.6
) 22
.7 (±
0.1
) 19
.3 (±
0.2
) 18
.6 (±
0.7
) 18
.0 (±
2.7
) 17
.6 (±
3.3
) 20
.0 (±
0.4
)
C15
n.
d.
0.2
(± 0
.2)
0.4
(± 0
.0)
n.d.
0.
2 (±
0.2
) n.
d.
n.d.
C
16
16.3
(± 0
.7)
14.9
(± 0
.5)
11.7
(± 0
.2)
12.7
(± 0
.5)
10.5
(± 1
.2)
10.8
(± 1
.1)
11.7
(± 0
.5)
C18
4.
5 (±
1.1
) 1.
4 (±
0.3
) 0.
4 (±
0.1
) 0.
4 (±
0.3
) 0.
4 (±
0.1
) 0.
4 (±
0.4
) 0.
3 (±
0.3
)
Σ m
onou
nsat
urat
es
18.6
(± 0
.2)
17.5
(± 0
.3)
23.2
(± 0
.8)
22.1
(± 0
.3)
19.7
(± 1
.4)
17.4
(± 0
.4)
16.6
(± 0
.5)
C14
:1
n.d.
0.
1 (±
0.1
) 0.
4 (±
0.0
) n.
d.
0.3
(± 0
.0)
n.d.
n.
d.
C16
:1n-
7 2.
6 (±
0.2
) 3.
5 (±
0.4
) 6.
0 (±
0.2
) 5.
8 (±
0.3
) 5.
1 (±
0.2
) 5.
1 (±
0.3
) 5.
0 (±
0.5
)
C17
:1
n.d.
n.
d.
n.d.
n.
d.
0.4
(± 0
.2)
n.d.
n.
d.
C18
:1
16.0
(± 0
.4)
13.8
(± 0
.2)
16.9
(± 1
.0)
16.4
(± 0
.6)
13.8
(± 1
.1)
12.3
(± 0
.3)
11.6
(± 0
.8)
Σ po
lyun
satu
rate
s 38
.7 (±
2.3
) 43
.3 (±
0.5
) 44
.9 (±
0.9
) 46
.2 (±
1.0
) 51
.2 (±
2.8
) 53
.8 (±
4.1
) 51
.3 (±
0.9
)
C16
:2n-
6 0.
9 (±
0.2
) 0.
8 (±
0.3
) 0.
5 (±
0.0
) 0.
3 (±
0.3
) 0.
4 (±
0.0
) 0.
4 (±
0.0
) 0.
5 (±
0.1
)
C18
:2n-
6 4.
5 (±
0.3
) 4.
7 (±
0.3
) 11
.6 (±
0.6
) 11
.4 (±
1.4
) 7.
6 (±
3.3
) 6.
9 (±
2.4
) 4.
3 (±
0.5
)
C18
:3n-
6 0.
4 (±
0.3
) 0.
6 (±
0.1
) 1.
1 (±
0.0
) 1.
4 (±
0.1
) 1.
3 (±
0.5
) 1.
2 (±
0.6
) 0.
4 (±
0.3
)
C20
:4n-
6 1.
2 (±
0.5
) 0.
5 (±
0.1
) 0.
5 (±
0.0
) n.
d.
0.2
(± 0
.2)
n.d.
n.
d.
C16
:2n-
4 1.
1 (±
0.4
) 0.
7 (±
0.0
) 0.
3 (±
0.0
) n.
d.
0.5
(± 0
.1)
0.9
(± 0
.6)
1.8
(± 0
.3)
C18
:3n-
3 6.
9 (±
1.8
) 9.
4 (±
1.3
) 9.
5 (±
0.2
) 10
.7 (±
0.7
) 11
.8 (±
0.9
) 11
.2 (±
0.7
) 10
.8 (±
0.6
)
C18
:4n-
3 17
.3 (±
1.2
) 20
.3 (±
0.4
) 17
.4 (±
0.3
) 17
.6 (±
2.0
) 24
.1 (±
3.6
) 27
.2 (±
1.6
) 27
.9 (±
0.7
)
C20
:5n-
3 0.
4 (±
0.3
) 0.
8 (±
0.0
) 0.
7 (±
0.0
) 0.
4 (±
0.3
) 0.
5 (±
0.2
) 0.
6 (±
0.1
) 0.
5 (±
0.0
)
C22
:6n-
3 6.
0 (±
0.4
) 5.
6 (±
0.4
) 3.
3 (±
0.1
) 4.
5 (±
0.8
) 4.
7 (±
1.1
) 5.
4 (±
0.4
) 5.
2 (±
0.1
)
Tabl
e 8.
1.7
Cha
nges
in
gl
ycol
ipid
fa
tty
acid
co
mpo
sitio
n (%
) of
Is
ochr
ysis
sp
. ov
er
grow
th.
Lag
(LA
), ea
rly l
ogar
ithm
ic (
EL),
loga
rithm
ic (
L),
late
log
arith
mic
(LL
), ea
rly s
tatio
nary
(ES
), st
atio
nary
(S)
and
lat
e st
atio
nary
(LS)
pha
se. R
esul
ts p
rese
nted
as m
ean
valu
es (±
SD
; n=3
), n.
d.: n
ot d
etec
ted.
VIII APPENDICES
185
FA (%
) L
A
EL
L
L
L
ES
MS
LS
Σ sa
tura
tes
42.7
(± 1
.4)
42.3
(± 0
.3)
37.7
(± 1
.0)
39.1
(± 1
.7)
33.7
(± 2
.6)
39.6
(± 5
.6)
40.0
(± 1
.3)
C14
23
.2 (±
0.5
) 23
.7 (±
0.2
) 21
.4 (±
0.7
) 20
.0 (±
1.3
) 18
.1 (±
1.5
) 22
.4 (±
4.2
) 23
.4 (±
0.5
)
C16
19
.5 (±
0.9
) 18
.3 (±
0.4
) 16
.3 (±
0.3
) 19
.1 (±
0.4
) 15
.6 (±
1.1
) 17
.2 (±
1.4
) 16
.6 (±
0.8
)
C18
n.
d.
0.3
(± 0
.3)
n.d.
n.
d.
n.d.
n.
d.
n.d.
Σ
mon
ouns
atur
ates
13
.5 (±
1.0
) 12
.7 (±
0.8
) 13
.7 (±
0.8
) 15
.7 (±
0.9
) 17
.4 (±
1.4
) 18
.9 (±
1.2
) 14
.9 (±
0.9
)
C16
:1n-
7 n.
d.
0.5
(± 0
.0)
1.2
(± 0
.2)
1.0
(± 0
.9)
1.3
(± 0
.1)
2.2
(± 0
.5)
2.0
(± 0
.3)
C18
:1
13.5
(± 1
.0)
12.2
(± 0
.7)
12.5
(± 0
.6)
14.7
(± 0
.6)
16.2
(± 1
.3)
16.7
(± 0
.9)
12.9
(± 0
.6)
Σ po
lyun
satu
rate
s 43
.8 (±
2.0
) 45
.0 (±
1.1
) 48
.5 (±
1.8
) 45
.2 (±
1.7
) 48
.8 (±
4.0
) 41
.5 (±
4.9
) 45
.1 (±
0.5
)
C16
:2n-
6 n.
d.
0.5
(± 0
.4)
n.d.
n.
d.
n.d.
n.
d.
n.d.
C
18:2
n-6
3.7
(± 0
.6)
3.1
(± 0
.3)
5.1
(± 0
.6)
6.5
(± 0
.3)
4.5
(± 0
.7)
5.4
(± 1
.4)
3.6
(± 0
.4)
C16
:2n-
4 1.
4 (±
1.0
) n.
d.
n.d.
n.
d.
n.d.
n.
d 4.
4 (±
2.2
) C
18:3
n-3
3.4
(± 1
.5)
1.9
(± 0
.3)
2.3
(± 0
.3)
4.0
(± 0
.8)
2.7
(± 0
.4)
5.4
(± 1
.1)
3.8
(± 0
.7)
C18
:4n-
3 3.
1 (±
0.8
) 2.
8 (±
0.4
) 2.
6 (±
0.4
) 3.
6 (±
0.6
) 3.
8 (±
0.6
) 7.
5 (±
1.5
) 5.
9 (±
0.8
)
C20
:5n-
3 n.
d.
0.6
(± 0
.1)
0.6
(± 0
.5)
0.8
(± 0
.7)
n.d.
n.
d.
n.d.
C
22:6
n-3
32.2
(± 5
.8)
36.2
(± 2
.3)
38.0
(± 3
.3)
30.3
(± 3
.0)
37.8
(± 4
.8)
23.3
(± 4
.2)
27.5
(± 3
.8)
Tabl
e 8.
1.8
Cha
nges
in
po
lar
lipid
fa
tty
acid
co
mpo
sitio
n (%
) of
Is
ochr
ysis
sp
. ov
er
grow
th.
Lag
(LA
), ea
rly l
ogar
ithm
ic (
EL),
loga
rithm
ic (
L),
late
log
arith
mic
(LL
), ea
rly s
tatio
nary
(ES
), st
atio
nary
(S)
and
lat
e st
atio
nary
(LS)
pha
se. R
esul
ts p
rese
nted
as m
ean
valu
es (±
SD
; n=3
), n.
d.: n
ot d
etec
ted.
VIII APPENDICES
186
FA (%
) L
A
EL
L
L
L
ES
S L
S Σ
satu
rate
s 40
.1 (±
10.7
) 23
.3 (±
2.3
) 20
.6 (±
1.0
) 22
.8 (±
0.6
) 30
.1 (±
1.3
) 18
.9 (±
0.8
) 26
.3 (±
1.0
) C
14
12.2
(± 2
.8)
7.9
(± 0
.7)
7.4
(± 0
.3)
9.6
(± 0
.5)
8.7
(± 0
.5)
10.4
(± 0
.3)
9.0
(± 0
.3)
C15
n.
d.
n.d.
n.
d.
n.d.
0.
1 (±
0.1
) n.
d.
0.1
(± 0
.1)
C16
25
.2(±
7.1
) 14
.5 (±
1.5
) 12
.7 (±
0.7
) 13
.2 (±
0.3
) 21
.0 (±
1.1
) 8.
3 (±
0.4
) 17
.0 (±
0.9
) C
18
2.7
(± 0
.8)
0.9
(± 0
.2)
0.5
(± 0
.2)
n.d.
0.
3 (±
0.2
) 0.
1 (±
0.1
) 0.
2 (±
0.1
) Σ
mon
ouns
atur
ates
16
.6 (±
0.3
) 12
.9 (±
0.1
) 11
.5 (±
0.1
) 11
.7 (±
0.1
) 18
.3 (±
0.9
) 16
.1 (±
0.6
) 14
.9 (±
1.9
) C
16:1
n-7
16.6
(± 0
.3)
12.9
(± 0
.1)
11.2
(± 0
.1)
11.7
(± 0
.1)
17.3
(± 0
.7)
15.6
(± 0
.6)
14.3
(± 1
.7)
C18
:1
n.d.
n.
d.
0.3
(± 0
.0)
n.d.
0.
9 (±
0.2
) 0.
5 (±
0.1
) 0.
6 (±
0.2
) Σ
poly
unsa
tura
tes
43.3
(± 1
0.8)
63
.7 (±
2.3
) 67
.9 (±
0.9
) 65
.5 (±
0.7
) 51
.6 (±
2.0
) 65
.0 (±
1.3
) 58
.8 (±
2.9
) C
16:2
n-6
3.4
(± 0
.2)
3.7
(± 0
.1)
3.4
(± 0
.0)
3.2
(± 0
.3)
1.6
(± 0
.2)
1.4
(± 1
.0)
1.9
(± 0
.3)
C18
:2n-
6 1.
0 (±
0.9
) 2.
1 (±
0.1
) 1.
6 (±
0.1
) 1.
2 (±
0.1
) 2.
1 (±
0.2
) 2.
7 (±
0.2
) 1.
8 (±
0.1
) C
18:3
n-6
n.d.
1.
9 (±
0.1
) 1.
2 (±
0.1
) 0.
5 (±
0.0
) 0.
4 (±
0.0
) 0.
2 (±
0.0
) 0.
4 (±
0.0
) C
20:2
n-6
n.d.
n.
d.
n.d.
n.
d.
0.2
(± 0
.02)
n.
d.
n.d.
C
20:4
n-6
n.d.
n.
d.
0.4
(± 0
.0)
0.3
(± 0
.0)
0.6
(± 0
.1)
0.2
(± 0
.0)
0.5
(± 0
.0)
C16
:2n-
4 n.
d.
1.0
(± 0
.1)
1.0
(± 0
.0)
1.5
(± 0
.2)
0.7
(± 0
.2)
0.8
(± 0
.7)
0.9
(± 0
.2)
C16
:3n-
4 n.
d.
n.d.
0.
5 (±
0.0
) 0.
7 (±
0.2
) 0.
5 (±
0.1
) 0.
6 (±
0.2
) 0.
5 (±
0.0
) C
18:3
n-3
n.d.
1.
8 (±
0.2
) 2.
3 (±
0.1
) 2.
6 (±
0.8
) 1.
5 (±
0.4
) 0.
7 (±
0.1
) 1.
6 (±
0.2
) C
18:4
n-3
7.3
(± 2
.3)
9.3
(± 0
.5)
9.8
(± 0
.5)
8.2
(± 1
.0)
5.7
(± 0
.1)
9.7
(± 0
.6)
7.0
(± 0
.2)
C20
:5n-
3 19
.4 (±
7.4
) 32
.2 (±
2.0
) 36
.3 (±
0.4
) 37
.1 (±
0.6
) 28
.1 (±
1.5
) 27
.5 (±
0.5
) 32
.8 (±
3.0
) C
22:6
n-3
12.2
(± 1
.1)
11.2
(± 1
.1)
10.8
(± 0
.5)
9.7
(± 0
.1)
9.9
(± 0
.6)
20.5
(± 1
.8)
11.1
(± 1
.1)
C16
:4n-
1 n.
d.
0.6
(± 0
.1)
0.6
(± 0
.0)
0.6
(± 0
.0)
0.4
(± 0
.0)
0.6
(± 0
.0)
0.4
(± 0
.0)
Tabl
e 8.
1.9
Cha
nges
in
to
tal
fatty
ac
id
com
posi
tion
(%)
of
Pavl
ova
luth
eri
over
gr
owth
. La
g (L
A),
early
log
arith
mic
(EL
), lo
garit
hmic
(L)
, la
te l
ogar
ithm
ic (
LL),
early
sta
tiona
ry (
ES),
stat
iona
ry (
S) a
nd l
ate
stat
iona
ry (L
S) p
hase
. Res
ults
pre
sent
ed a
s mea
n va
lues
(± S
D; n
=3),
n.d.
: not
det
ecte
d.
VIII APPENDICES
187
FA (%
) L
A
EL
L
L
L
ES
S L
S Σ
satu
rate
s 28
.0 (±
6.8
) 15
.5 (±
0.5
) 13
.3 (±
0.4
) 19
.8 (±
0.7
) 35
.1 (±
1.0
) 15
.4 (±
0.9
) 31
.7 (±
1.4
) C
14
4.6
(± 1
.3)
4.0
(± 0
.4)
4.1
(± 0
.7)
6.0
(± 0
.1)
6.9
(± 0
.3)
7.4
(± 0
.2)
7.4
(± 0
.3)
C16
16
.7 (±
4.6
) 9.
2 (±
0.3
) 7.
4 (±
0.3
) 10
.0 (±
0.5
) 27
.5 (±
1.2
) 7.
5 (±
0.7
) 23
.4 (±
1.3
) C
18
6.8
(± 1
.3)
2.3
(± 0
.4)
1.8
(± 0
.6)
3.7
(± 0
.2)
0.7
(± 0
.1)
0.5
(± 0
.1)
0.9
(± 0
.2)
Σ m
onou
nsat
urat
es
21.4
(± 1
.7)
12.8
(± 1
.0)
10.8
(± 1
.0)
13.6
(± 0
.8)
29.6
(± 0
.4)
21.1
(± 1
.3)
28.6
(± 0
.1)
C16
:1n-
7 13
.9 (±
4.3
) 11
.3 (±
0.7
) 10
.8 (±
1.0
) 13
.6 (±
0.8
) 27
.7 (±
0.4
) 20
.3 (±
1.1
) 27
.0 (±
0.1
) C
18:1
7.
6 (±
6.0
) 1.
5 (±
0.4
) n.
d.
n.d.
1.
8 (±
0.2
) 0.
8 (±
0.2
) 1.
6 (±
0.0
) Σ
poly
unsa
tura
tes
50.6
(± 7
.6)
71.7
(± 1
.5)
75.9
(± 1
.3)
66.6
(± 1
.5)
35.3
(± 1
.2)
63.5
(± 2
.2)
39.7
(± 1
.3)
C16
:2n-
6 4.
9 (±
2.3
) 9.
8 (±
2.3
) 15
.6 (±
1.0
) 26
.1 (±
1.4
) 2.
6 (±
0.4
) 2.
5 (±
0.3
) 4.
5 (±
1.7
) C
18:2
n-6
11.9
(± 1
0.8)
4.
7 (±
0.7
) 2.
4 (±
0.7
) n.
d.
3.8
(± 0
.3)
3.8
(± 0
.1)
3.6
(± 0
.2)
C18
:3n-
6 0.
8 (±
0.1
) 1.
8 (±
0.1
) 0.
8 (±
0.7
) n.
d.
0.4
(± 0
.0)
n.d.
0.
4 (±
0.3
) C
20:2
n-6
n.d.
n.
d.
n.d.
n.
d.
n.d
n.d.
0.
3 (±
0.3
) C
20:4
n-6
n.d.
n.
d.
n.d.
n.
d.
n.d
n.d.
0.
7 (±
0.1
) C
22:2
n-6
n.d.
n.
d.
n.d.
n.
d.
n.d
n.d.
n.
d.
C16
:2n-
4 n.
d.
n.d.
n.
d.
n.d.
0.
3 (±
0.0
) 0.
4 (±
0.0
) 0.
3 (±
0.2
) C
16:3
n-4
n.d.
n.
d.
n.d.
n.
d.
n.d
0.6
(± 0
.0)
n.d.
C
18:3
n-3
n.d.
1.
6 (±
0.2
) 2.
0 (±
0.1
) n.
d.
0.6
(± 0
.1)
0.6
(± 0
.1)
0.7
(± 0
.0)
C18
:4n-
3 5.
9 (±
1.0
) 9.
6 (±
0.2
) 10
.0 (±
0.1
) 5.
4 (±
0.9
) 1.
9 (±
0.1
) 8.
8 (±
1.0
) 2.
3 (±
0.2
) C
20:5
n-3
15.9
(± 7
.1)
26.1
(± 1
.7)
26.2
(± 0
.1)
19.3
(± 1
.9)
14.9
(± 0
.4)
19.5
(± 0
.1)
15.1
(± 0
.2)
C22
:6n-
3 11
.2 (±
2.8
) 15
.9 (±
2.6
) 15
.6 (±
2.0
) 10
.0 (±
0.4
) 8.
4 (±
1.3
) 26
.1 (±
1.5
) 10
.4 (±
1.1
) C
16:4
n-1
n.d.
2.
1 (±
0.5
) 3.
3 (±
0.2
) 5.
8 (±
3.0
) 1.
1 (±
0.2
) 1.
2 (±
0.1
) 1.
6 (±
0.4
)
Tabl
e 8.
1.10
C
hang
es
in
neut
ral
lipid
fa
tty
acid
co
mpo
sitio
n (%
) of
Pa
vlov
a lu
ther
i ov
er
grow
th.
Lag
(LA
), ea
rly l
ogar
ithm
ic (
EL),
loga
rithm
ic (
L),
late
log
arith
mic
(LL
), ea
rly s
tatio
nary
(ES
), st
atio
nary
(S)
and
lat
e st
atio
nary
(LS)
pha
se. R
esul
ts p
rese
nted
as m
ean
valu
es (±
SD
; n=3
), n.
d.: n
ot d
etec
ted.
VIII APPENDICES
188
FA (%
) L
A
EL
L
L
L
ES
S L
S Σ
satu
rate
s 39
.2 (±
11.
4)
24.6
(± 1
.2)
22.1
(± 0
.7)
24.6
(± 0
.3)
27.7
(± 0
.2)
25.2
(± 1
.2)
24.2
(± 0
.4)
C14
14
.2 (±
3.7
) 8.
6 (±
0.1
) 8.
4 (±
0.2
) 9.
9 (±
0.5
) 11
.9 (±
0.8
) 15
.6 (±
0.3
) 11
.1 (±
0.9
) C
15
n.d.
n.
d.
n.d.
0.
3 (±
0.0
) 0.
3 (±
0.0
) n.
d.
0.2
(± 0
.2)
C16
24
.3 (±
7.5
) 14
.8 (±
0.9
) 13
.1 (±
0.7
) 13
.7 (±
0.2
) 14
.8 (±
0.6
) 8.
5 (±
0.6
) 12
.6 (±
1.1
) C
18
0.8
(± 0
.2)
1.3
(± 0
.2)
0.6
(± 0
.2)
0.7
(± 0
.1)
0.7
(± 0
.2)
1.1
(± 0
.4)
0.4
(± 0
.0)
Σ m
onou
nsat
urat
es
18.0
(± 0
.8)
14.8
(± 0
.4)
13.9
(± 0
.2)
12.9
(± 0
.0)
10.2
(± 0
.4)
10.7
(± 0
.6)
9.6
(± 0
.5)
C16
:1n-
7 16
.6 (±
0.5
) 14
.8 (±
0.4
) 13
.0 (±
0.6
) 12
.9 (±
0.0
) 9.
7 (±
0.4
) 10
.7 (±
0.6
) 9.
3 (±
0.3
) C
18:1
1.
4 (±
0.6
) n.
d.
0.8
(± 0
.8)
n.d.
0.
5 (±
0.0
) n.
d.
0.3
(± 0
.2)
Σ po
lyun
satu
rate
s 42
.8 (±
12.
2)
60.5
(± 0
.9)
64.1
(± 0
.7)
62.5
(± 0
.3)
62.1
(± 0
.6)
64.1
(± 1
.7)
66.2
(± 0
.6)
C16
:2n-
6 2.
3 (±
0.4
) 1.
9 (±
0.4
) 0.
9 (±
0.1
) 0.
7 (±
0.2
) 0.
7 (±
0.3
) 0.
5 (±
0.4
) 0.
7 (±
0.0
) C
18:2
n-6
2.1
(± 0
.6)
3.1
(± 0
.8)
2.0
(± 0
.2)
1.6
(± 0
.1)
2.7
(± 1
.1)
1.8
(± 0
.1)
1.2
(± 0
.2)
C18
:3n6
1.
0 (±
0.2
) 2.
3 (±
0.2
) 1.
6 (±
0.2
) 0.
6 (±
0.0
) 0.
4 (±
0.0
) 0.
8 (±
0.1
) 0.
4 (±
0.0
) C
20:4
n-6
n.d.
n.
d.
0.4
(± 0
.0)
0.3
(± 0
.0)
n.d
n.d.
0.
2 (±
0.2
) C
22:2
n-6
n.d.
n.
d.
n.d.
n.
d.
n.d
n.d.
n.
d.
C16
:2n-
4 0.
4 (±
0.4
) 1.
4 (±
0.1
) 1.
4 (±
0.0
) 1.
9 (±
0.2
) 1.
3 (±
0.2
) 0.
5 (±
0.4
) 1.
5 (±
0.1
) C
16:3
n-4
n.d.
0.
6 (±
0.1
) 0.
7 (±
0.1
) 0.
9 (±
0.2
) 0.
9 (±
0.2
) 1.
1 (±
0.1
) 0.
9 (±
0.0
) C
18:3
n3
0.8
(± 0
.2)
2.2
(± 0
.2)
3.0
(± 0
.1)
3.3
(± 1
.0)
2.8
(0±.
6)
n.d.
2.
7 (±
0.1
) C
18:4
n3
7.4
(± 3
.9)
11.6
(± 0
.6)
12.8
(± 0
.6)
10.7
(± 1
.2)
11.5
(± 1
.1)
16.0
(± 0
.7)
12.9
(± 1
.4)
C20
:5n-
3 17
.9 (±
7.9
) 34
.6 (±
1.8
) 40
.3 (±
0.3
) 41
.5 (±
0.5
) 42
.6 (±
0.3
) 41
.7 (±
1.6
) 45
.1 (±
0.7
) C
22:6
n-3
10.8
(± 0
.5)
2.9
(± 0
.6)
1.2
(± 0
.1)
1.1
(± 0
.1)
0.8
(± 0
.1)
1.8
(± 0
.2)
0.6
(± 0
.1)
Tabl
e 8.
1.11
C
hang
es
in
glyc
olip
id
fatty
ac
id
com
posi
tion
(%)
of
Pavl
ova
luth
eri
over
gr
owth
. La
g (L
A),
early
log
arith
mic
(EL
), lo
garit
hmic
(L)
, la
te l
ogar
ithm
ic (
LL),
early
sta
tiona
ry (
ES),
stat
iona
ry (
S) a
nd l
ate
stat
iona
ry (L
S) p
hase
. Res
ults
pre
sent
ed a
s mea
n va
lues
(± S
D; n
=3),
n.d.
: not
det
ecte
d.
VIII APPENDICES
189
FA (%
) L
A
EL
L
L
L
ES
S L
S Σ
satu
rate
s 56
.6 (±
15.
7)
35.0
(± 4
.7)
24.4
(± 3
.5)
26.1
(± 0
.2)
24.9
(± 1
.1)
29.5
(± 1
.1)
27.2
(± 1
.7)
C14
15
.2 (±
4.8
) 11
.1 (±
2.1
) 7.
1 (±
1.9
) 9.
7 (±
0.3
) 5.
5 (±
0.7
) 12
.6 (±
0.7
)8.
0 (±
1.0
) C
16
39.5
(± 1
1.0)
23
.1 (±
2.7
) 16
.9 (±
1.5
) 15
.9 (±
0.4
) 18
.4 (±
0.6
) 15
.9 (±
0.5
)18
.7 (±
0.7
) C
18
1.9
(± 0
.8)
0.8
(± 0
.1)
0.5
(± 0
.1)
0.5
(± 0
.4)
0.9
(± 0
.1)
1.0
(± 0
.1)
0.5
(± 0
.0)
Σ m
onou
nsat
urat
es
11.0
(± 4
.0)
9.6
(± 0
.4)
6.8
(± 1
.5)
8.1
(± 1
.4)
6.2
(± 1
.0)
6.8
(± 1
.0)
4.4
(± 0
.7)
C16
:1n-
7 7.
1 (±
2.0
) 8.
5 (±
0.2
) 6.
1 (±
1.4
) 6.
2 (±
0.3
) 3.
3 (±
0.3
) 4.
8 (±
0.2
) 3.
9 (±
0.6
) C
18:1
3.
9 (±
5.9
) 1.
1 (±
0.3
) 0.
8 (±
0.2
) 1.
9 (±
1.3
) 2.
1 (±
1.0
) 2.
1 (±
1.0
) 0.
5 (±
0.0
) C
20:1
n.
d.
n.d.
n.
d.
n.d.
0.
3 (±
0.2
) n.
d.
n.d.
Σ
poly
unsa
tura
tes
32.4
(± 1
3.0)
55
.4 (±
4.4
) 68
.7 (±
5.0
) 65
.8 (±
1.2
) 68
.9 (±
0.1
) 63
.7 (±
2.1
)68
.3 (±
2.3
) C
16:2
n-6
n.d.
0.
4 (±
0.4
) 0.
6 (±
0.2
) 0.
8 (±
0.7
) 0.
4 (±
0.4
) 1.
0 (±
0.1
) 0.
6 (±
0.2
) C
18:2
n-6
10.8
(± 1
5.2)
1.
7 (±
0.3
) 0.
8 (±
0.1
) 1.
7 (±
1.5
) 2.
7 (±
1.1
) 2.
4 (±
0.8
) n.
d.
C20
:4n-
6 n.
d.
n.d.
0.
3 (±
0.3
) n.
d.
0.7
(± 0
.1)
n.d.
0.
6 (±
0.1
) C
18:3
n-3
n.d.
n.
d.
0.7
(± 0
.2)
0.9
(± 0
.1)
0.5
(± 0
.1)
0.4
(± 0
.3)
0.5
(± 0
.1)
C18
:4n-
3 n.
d.
1.1
(± 0
.1)
0.6
(± 0
.1)
0.3
(± 0
.3)
0.3
(± 0
.2)
1.2
(± 0
.2)
0.2
(± 0
.2)
C20
:5n-
3 14
.5 (±
12.
6)
26.3
(± 2
.5)
27.8
(± 1
.1)
23.9
(± 0
.6)
24.3
(± 0
.3
26.6
(± 1
.0)
24.0
(± 0
.9)
C22
:6n-
3 7.
0 (±
3.5
) 26
.0 (±
2.2
) 37
.9 (±
4.4
) 38
.2 (±
2.0
) 40
.0 (±
0.7
) 32
.1 (±
1.9
)42
.4 (±
1.6
)
Tabl
e 8.
1.12
C
hang
es
in
pola
r lip
id
fatty
ac
id
com
posi
tion
(%)
of
Pavl
ova
luth
eri
over
gr
owth
. La
g (L
A),
early
log
arith
mic
(EL
), lo
garit
hmic
(L)
, la
te l
ogar
ithm
ic (
LL),
early
sta
tiona
ry (
ES),
stat
iona
ry (
S) a
nd l
ate
stat
iona
ry (L
S) p
hase
. Res
ults
pre
sent
ed a
s mea
n va
lues
(± S
D; n
=3),
n.d.
: not
det
ecte
d.
VIII APPENDICES
190
Appendix 8.2 Solvents trialled for gel-overlay assays with
4-methylumbelliferyl palmitate
Table 8.2 Solvents tested for use in 4-methylumbelliferyl palmitate gel-overlay assay
Solvent MUP soluble MUP emulsion formed
2-methoxyethanol - -
Dimethyl sulfoxide - -
Hexane - -
Chloroform + -
Acetonitrile - -
Acetone - -
Ethyl acetate - -
Isopropanol - -
Methanol - -
Dioxane + -
Dioxane +1% Triton X-100 + +
Appendix 8.3 Ion-exchange chromatography of Isochrysis sp.
protein homogenate
Figures 8.3.1 and 8.3.2 display the anion- and cation-exchange chromatography of
the protein homogenate from Isochrysis sp., respectively. The figures also show the
activity of the starting material (start), flow-through (ft) and fractions of interest
against pNPM (Figures 8.3.1B and 8.3.2B). Following this, the assays performed in
attempts to restore activity in the fractions of interest (by addition of starting
material) can be found in Figures 8.3.1C and 8.3.2C.
VIII APPENDICES
191
Figure 8.3.1 Anion-exchange chromatography of the protein homogenate from Isochrysis sp..
(A) Chromatogram of Isochrysis sp. protein homogenate, green line represents the absorbance at 280
nm, black line represents concentration of buffer B, red line indicates conductivity and light green
bars represent fractions of which the activity was investigated. (B) Activity ( A410/min) of starting
material (start), flow through (ft) and fractions of interest against pNPM. (C) Activity ( A410/min) of
fractions as in B, mixed with starting material (1:1) against pNPM.
00.005
0.010.015
0.020.025
0.030.035
start ft 6 7 9 10 11 12 13 16 17
Actv
iity
(A 4
10)
Fraction
0
0.002
0.004
0.006
0.008
0.01
0.012
start ft 6 7 9 10 11 12 13 16 17
Activ
ity (
A 410
)
Fraction
A
B
C
VIII APPENDICES
192
Figure 8.3.2 Cation-exchange chromatography of the protein homogenate from Isochrysis sp..
(A) Chromatogram of Isochrysis sp. protein homogenate, green line represents the absorbance at 280
nm, black line represents concentration of buffer B, red line indicates conductivity and light green
bars represent fractions of which the activity was investigated. (B) Activity ( A410/min) of starting
material (start), flow through (ft) and fractions of interest against pNPM. (C) Activity ( A410/min) of
fractions as in B, mixed with starting material (1:1) against pNPM.
0
0.005
0.01
0.015
0.02
0.025
0.03
start ft 3 6 7 8 9 12 14
Activ
ity (A
410)
Fraction
0.000
0.005
0.010
0.015
0.020
start ft 3 6 7 8 9 12 14
Activ
ity ( A
410)
Fraction
A
B
C
VIII APPENDICES
193
Appendix 8.4 Acidic and basic native-PAGE band analysis
Figure 8.4 MUB gel-overlay assay of P. lutheri protein homogenate separated by acidic and
basic native-PAGE. (1) Acidic native-PAGE, (2) basic native-PAGE. Gel load was 30 μl of P. lutheri
crude protein homogenate. Arrows indicate bands that were excised from each gel for subsequent
analyses by SDS-PAGE.
Band 1
1 2
Band 1
Band 2
VIII APPENDICES
194
Appendix 8.5 Sequences identified by MALDI TOF/TOF
The sequence matches identified by the Mascot database from the gel bands 3A and
3B are presented below. Ions score is -10*Log(P), where P is the probability that the
observed match is a random event. Individual ions scores >61 indicate identity or
extensive homology (p<0.05).
Sequence matches from band 3A:
1.
gi|61229242 Mass: 40989 Score: 312 Matches: 5(4) Sequences: 4(3)
S-adenosyl-L-homocysteine hydrolase [Pavlova lutheri] 2.
gi|21913671 Mass: 45107 Score: 220 Matches: 3(3) Sequences: 3(3)
ribulose-1,5-bisphosphate carboxylase/oxygenase [Pavlova gyrans] 3.
gi|21913673 Mass: 38073 Score: 206 Matches: 3(3) Sequences: 3(3)
ribulose-1,5-bisphosphate carboxylase/oxygenase [Pavlova lutheri] 4.
gi|3757503 Mass: 51598 Score: 106 Matches: 1(1) Sequences: 1(1)
ribulose-1,5-bisphosphate carboxylase/oxygenase large subunit [Heringia mirabilis]
Sequence matches from band 3B:
1.
gi|10645188 Mass: 38491 Score: 117 Matches: 3(0) Sequences: 3(0)
cytosolic aldolase [Fragaria x ananassa]
Appendix 8.6 Extinction coefficients of p-nitrophenol for
defining lipase pH optima
Table 8.6 Extinction coefficients of p-nitrophenol at different pH.
pH 6.0 7.0 7.5 8.0 8.5 9.0
M= (M-1 cm-1) 1607 8376 13294 16302 17104 17738 Values are averages of triplicates
VIII APPENDICES
195
Appendix 8.7 13C NMR of fatty acids used in pNP-acyl ester
synthesis
Table 8.7 The observed 13C NMR chemical shifts (δ) of free fatty acids.
Carbon Fatty acid
C16:1 n-9 C18:1 n-9 C18:2 n-6 C18:3 n-3 C20:5 n-3 C22:6 n-3
Carboxyl (C1) 180.71 180.63 180.47 180.46 180.25 179.56
130.16 θ 130.16 θ 130.37 θ 132.10 θ 132.17 θ 132.18 θ
129.37 θ 129.86 θ 130.15 θ 130.39 θ 129.17 θ 129.73 θ
34.27 34.25 128.22 θ 128.43 θ 129.03 θ 128.71 θ
31.94 32.06 128.05 θ 128.39 θ 128.88 θ 128.45 θ
29.88 29.92 34.22 127.90 θ 128.53 θ 128.41 θ
29.82 29.83 31.68 127.26 θ 128.30 θ 128.38 θ
29.29 29.68 29.73 34.21 128.29 θ 128.23 θ
29.21 29.47 29.50 29.71 128.22 θ 128.22 θ
29.18 29.47 29.29 29.29 128.01 θ 128.12 θ
29.14 29.29 29.22 29.22 127.17 θ 128.02 θ
27.37 ψ 29.18 29.18 29.17 33.54 127.68 θ
27.30 ψ 29.18 27.36 ψ 27.34 ψ 26.58 ψ 127.16 θ
24.80 27.36 ψ 27.33 ψ 25.76ǂ 25.76ǂ 34.11
22.81 27.30 ψ 25.78ǂ 25.68ǂ 25.76ǂ 25.77ǂ
14.24 24.80 24.80 24.79 25.76ǂ 25.77ǂ
22.83 22.73 20.70 ψ 25.67ǂ 25.77ǂ
14.25 14.22 14.42 24.60 25.73ǂ
20.69 ψ 25.68ǂ
14.40 22.62 ψ
20.70 ψ
Methyl (ω) 14.41
Olefinic carbon θ, allylic carbon adjacent to a single olefinic carbon ψ, allylic carbon adjacent to two
olefinic carbons ǂ.
VIII APPENDICES
196
0
20
40
60
80
100
120
Rela
tive
activ
ity (%
)
pNP-acyl ester
0
20
40
60
80
100
120
Rela
tive
activ
ity (%
)
pNP-acyl ester
0
20
40
60
80
100
120
140
Rela
tive
activ
ity (%
)
pNP-acyl ester
Appendix 8.8 pNP-acyl ester profiles for CalB, TlL and RmL
without CaCl2
Figure 8.8 Activity profiles of CalB, TlL and RmL against a range of pNP-acyl esters in the
absence of CaCl2. The relative activity of each substrate is normalised to 100% of the substrate for
which the highest activity was obtained. (A) CalB, (B) TlL and (C) RmL. Substrates used are pNP-
acetate (C2:0), pNP-butyrate (C4:0), pNP-octanoate (C8:0), pNP-dodecanoate (C12:0), pNP-myristate
(C14:0), pNP-palmitate (C16:0), pNP-palmitoleate (C16:1), pNP-stearate (C18:0), pNP-oleate
(C18:1), pNP-linoleoate (C18:2), pNP-linolenoate (C18:3), pNP-eicosapentaenoate (C20:5) and pNP-
docosahexaenoate (C22:6).
A
B
C
VIII APPENDICES
197
Appendix 8.9 Titrimetric assay trace of TlL against lecithin
Figure 8.9 STAT-pH trace from a titrimetric assay of TlL against a lecithin emulsion with
CaCl2 added after assay initiation. Plotted as volume (mL) of 0.01 M NaOH titrated over time. Red
arrow indicates time point at which CaCl2 was added at 1200 sec (20 min).
