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DIGESTIVE LIPASES FROM CHINOOK SALMON
(ONCORHYNCHUS TSHAWYTSCHA) AND NEW ZEALAND HOKI
(MACRURONUS NOVAEZELANDIAE) – PURIFICATION,
CHARACTERIZATION, APPLICATION AND IMMOBILIZATION
BY
IVAN KURTOVIC
Department of Animal Science
McGill University, Montreal
April 2011
A thesis submitted to McGill University in partial fulfillment
of the requirements of the degree of
DOCTOR OF PHILOSOPHY
©Ivan Kurtovic 2011
ii
ABSTRACT
Lipases from two New Zealand commercial fish species, Chinook salmon
(Oncorhynchus tshawytscha) and New Zealand hoki (Macruronus
novaezelandiae) were investigated. The lipases were extracted from the
pyloric ceca and purified by affinity chromatography and gel filtration.
Calcium ions and sodium cholate were absolutely necessary both for lipase
stability in a polyacrylamide gel and for optimum activity against p-
nitrophenol esters. Both fish lipases had a pI value of 5.8 ± 0.1, were most
active at 35°C, were thermally labile, had a pH optimum of 8-8.5, were more
acid stable compared to other fish lipases studied to date, and showed good
stability in several water-immiscible solvents. The salmon enzyme was an
overall better catalyst for the hydrolysis of p-nitrophenyl caprate based on its
higher turnover number and lower activation energy for the hydrolysis
reaction. Based on their chemical and catalytic properties, the salmon and hoki
enzymes were classified as carboxyl ester lipases. Chinook salmon and hoki
lipases were then evaluated as flavour modifying agents in dairy products.
Cream was either incubated with the fish lipases or two commercially
available lipases used in dairy flavour development. The fish enzymes were
more similar to calf pregastric esterase in terms of the total amount and types
of fatty acids released (mainly short chain) over the course of the reaction. The
highest specificity was towards the key dairy product flavour and odour
compounds, butanoic and hexanoic acids. Immobilization of the salmon lipase
was then carried out on two hydrophobic supports. Salmon lipase immobilized
on octyl-Sepharose had 40- and 10-fold higher activity (on a dry weight basis)
against a tributyrin emulsion than the same lipase immobilized on Lewatit VP
OC 1600 and a microbial lipase immobilized on Lewatit (Novozym 435),
respectively. Salmon lipase-octyl-Sepharose was highly active against both
ghee and fish oil emulsions, but salmon lipase-Lewatit and Novozym 435 had
very low activities against the fish oil emulsion.
The potential for flavour enhancement in dairy products with both fish lipases
was demonstrated based on the free fatty acid composition and sensory
characteristics of lipase-treated creams. In addition, the immobilized salmon
lipase showed potential for low temperature modifications of emulsified lipids.
iii
RÉSUMÉ
Les lipases de deux espèces de poissons commerciaux en Nouvelle-Zélande, le
saumon quinnat (Oncorhynchus tshawytscha) et le hoki de la Nouvelle-
Zélande (Macruronus novaezelandiae) ont été étudiées. Les lipases ont été
extraites des caeca pyloriques et purifiées par chromatographie d'affinité et gel
filtration. Les ions de calcium et cholate de sodium étaient absolument
nécessaires pour la stabilité de la lipase dans un gel de polyacrylamide et de
l'activité optimale contre les esters p-nitrophénol. Les deux lipases de poissons
avaient une valeur pI de 5.8 ± 0.1, ont été les plus actives à 35°C, ont été
thermolabiles, a un pH optimum de 8 à 8.5, ont été plus stables en milieu acide
par rapport à d'autres lipases de poissons étudiées à ce jour, et ont montré une
bonne stabilité dans plusieurs solvants miscibles à l'eau. L'enzyme du saumon
a été un catalyseur globalement meilleure pour l'hydrolyse de caprate p-
nitrophényl en fonction de son nombre de rotation élevé et faible énergie
d'activation pour la réaction d'hydrolyse. Sur la base de leurs propriétés
chimiques et catalytiques, les enzymes de saumon et hoki ont été classées
comme des lipases ester carboxylique. Les lipases du saumon quinnat et hoki
ont été aussi évaluées comme agents de modification de la saveur dans les
produits laitiers. La crème a été mise à incuber avec les lipases de poisson ou
avec deux lipases disponibles dans le commerce utilisées dans le
développement du goût des produits laitiers. Les enzymes de poissons avaient
plus des similitude avec l‘estérase prégastrique du veau en termes de montant
total et les types d'acides gras libérés (principalement à chaîne courte) au cours
de la réaction. La plus grande spécificité a été observée aux composés clés de
la saveur et odeurs des produits laitiers: les acides butanoïque et hexanoïque.
L‘Immobilisation de la lipase de saumon a ensuite été effectuée sur deux
supports hydrophobes. La lipase de saumon immobilisée sur octyl-Sépharose
avait une activité 40- et 10-fois plus élevée (sur la base du poids sec) par
rapport à une émulsion de tributyrine que la même lipase immobilisée sur
Lewatit VP OC 1600 et une lipase microbienne immobilisée sur Lewatit
(Novozym 435), respectivement. La lipase-octyl-Sépharose de saumon a été
très active à la fois contre le ghee et les émulsions d'huile de poisson, mais la
lipase-Lewatit et Novozym 435 de saumon avaient des activités très faible
iv
contre l'émulsion d'huile de poisson.
Le potentiel d'amélioration de la saveur dans les produits laitiers avec les deux
lipases de poisson a été démontré sur la base de la composition des acides gras
libres et les caractéristiques sensorielles des crèmes traitées avec lipases. De
plus, la lipase de saumon immobilisée a démontré un potentiel pour des
modifications au niveau des lipides émulsionnés à basse température.
v
ACKNOWLEDGEMENTS
First of all, I am extremely grateful to my supervisor, Prof. Xin Zhao of the
Department of Animal Science. Prof. Zhao has provided clear guidance and
has been a constant source of pragmatic advice.
My co-supervisor, Dr. Susan Marshall, Natural Extracts team leader at Plant
and Food Research, New Zealand, and Prof. Benjamin Simpson of the
Department of Food Science and Agricultural Chemistry played a crucial role
in getting this PhD project off the ground. Prof. Simpson‘s vast experience in
food enzymology and seafood biochemistry has been invaluable for my
understanding of enzyme purification and characterization. His advice is most
appreciated. I would like to express my deepest and most sincere gratitude to
Dr. Susan Marshall, for her continued guidance, encouragement, excellent
advice, (and patience) throughout my studies. Sue‘s tireless leadership of the
Natural Extracts team is a quality to be admired and I have learned a lot from
her over the years.
I want to thank TC Chadderton of Plant and Food Research for playing an
instrumental role in setting up this project and making the PhD possible.
I am indebted to The New Zealand Institute for Plant and Food Research
Limited for providing the financial means for my PhD studies.
A big ‗thank you‘ goes to everyone at Natural Extracts team, Plant and Food
Research for their ongoing support and help throughout the duration of my
studies.
I want to thank Maria van Noorden from Plant and Food Research Library
team for a superb help with obtaining journal papers.
Kathleen Hofman, Graham Fletcher and Kevin Sutton are acknowledged for
reviewing the manuscripts.
Thanks to Corina Tatar for helping with abstract translation into French.
To the fellow students at the Department of Food Science and Agricultural
Chemistry and Department of Animal Science, I thank for their support.
And of course, a huge ―Thank you‖ to my wife, Miaolin Li and both of our
families for their constant support, encouragement and patience throughout my
studies.
vi
CONTRIBUTION TO KNOWLEDGE
Chapter III: The spectrophotometric lipase assay using p-nitrophenyl
palmitate as substrate was improved significantly for measuring the activity of
fish bile salt activated lipases with the inclusion of calcium ions. Activities
obtained with the optimized assay were 100-fold higher than those obtained
with the original conditions (no calcium ions and sodium cholate). The
titrimetric lipase assay using tributyrin as substrate (presented in appendix I)
was also improved (three-fold, compared with the original assay) for
measuring the activity of fish bile salt activated lipases. The improvement was
due in part to the modification to the concentrations of the emulsifiers and
inclusion of calcium ions.
An extraction and purification buffer was optimized (20% (w/v) glycerol was
included in a phosphate buffer) for maximum yield and stability of lipases
extracted from the pyloric ceca of Chinook salmon and hoki.
Lyophilization in the presence of high glycerol concentration was used as a
means to improve the stability of the two fish enzymes.
This study was the first to report on a clear visualization of activity in a
polyacrylamide gel overlaid with a substrate solution (i.e. by zymographic
analysis) for any fish or bile salt activated lipase. The activity of purified
Chinook salmon and hoki lipases was visualized in this way.
Chapter IV: A potential for flavour enhancement in dairy products using
Chinook salmon and hoki digestive lipases was demonstrated as the fish
enzymes had the highest specificity towards key dairy product flavour and
odour compounds (butanoic acid and hexanoic acid) when dairy cream was
used as substrate.
Chapter V: Chinook salmon bile salt activated lipase was immobilized on two
hydrophobic supports: octyl-Sepharose and Lewatit VP OC 1600.
Immobilization of an active and functional bile salt activated lipase for
application purposes has not been reported to date. The high activity of salmon
lipase-octyl-Sepharose against emulsified lipid substrates containing a wide
range of fatty acids (tributyrin, ghee and fish oil) demonstrated the
immobilized enzyme‘s potential applications for lipid modifications in
aqueous media.
vii
STATEMENT ABOUT THE THESIS FORMAT
This thesis has been written in the format of manuscripts submitted to
scientific journals in accordance with the McGill University Faculty of
Graduate and Postdoctoral Studies ―Guidelines for Thesis Preparation, C:
Manuscript-based Thesis‖. Four manuscripts are included, one of which is a
review paper comprising the literature review (chapter II). The other three
manuscripts are research papers and constitute chapters III, IV and V.
Supplementary material for chapters III and V is provided in the appendices.
Connecting statements are given between chapters II to V to facilitate
continuity.
viii
CONTRIBUTIONS OF AUTHORS
Four co-authored manuscripts submitted for publication are included in this
thesis. In all manuscripts, Ivan Kurtovic planned and coordinated the research,
designed and carried out the experiments, interpreted and analyzed the results,
and wrote the manuscripts. Sue Marshall and Xin Zhao co-supervised and
directed all phases of the research, and reviewed and edited the manuscripts
presented in the thesis.
Manuscript 1: Lipases from mammals and fishes
I. Kurtovic carried out the literature review and wrote the manuscript.
S. Marshall, B. Simpson and X. Zhao reviewed and edited the manuscript.
Manuscript 2: Purification and properties of digestive lipases from Chinook
salmon (Oncorhynchus tshawytscha) and New Zealand hoki (Macruronus
novaezelandiae)
I. Kurtovic designed and carried out all experiments, analyzed the results, and
wrote the manuscript. S. Marshall supervised the research plan, experimental
design and reviewed and edited the manuscript. B. Simpson and X. Zhao
reviewed and edited the manuscript.
Manuscript 3: Flavour development in dairy cream using fish digestive
lipases from Chinook salmon (Oncorhynchus tshawytscha) and New Zealand
hoki (Macruronus novaezelandiae)
I. Kurtovic designed and carried out the experiments, analyzed the results, and
wrote the manuscript. S. Marshall supervised the experimental design and
reviewed and edited the manuscript. M. Miller assisted with experiments
involving GC-MS and manuscript review and editing. X. Zhao reviewed and
edited the manuscript.
Manuscript 4: Hydrophobic immobilization of a bile salt activated lipase
from Chinook salmon (Oncorhynchus tshawytscha)
I. Kurtovic designed and carried out the experiments, analyzed the results, and
wrote the manuscript. S. Marshall supervised the experimental design and
reviewed and edited the manuscript. X. Zhao reviewed and edited the
manuscript.
ix
TABLE OF CONTENTS
ABSTRACT ...................................................................................................... ii
RÉSUMÉ ........................................................................................................ .iii
ACKNOWLEDGEMENTS ............................................................................ v
CONTRIBUTION TO KNOWLEDGE ........................................................ vi
STATEMENT ABOUT THE THESIS FORMAT ...................................... vii
CONTRIBUTIONS OF AUTHORS ........................................................... viii
TABLE OF CONTENTS ............................................................................... ix
LIST OF ABBREVIATIONS ....................................................................... xv
LIST OF TABLES ...................................................................................... xviii
LIST OF FIGURES ....................................................................................... xx
CHAPTER I. INTRODUCTION AND RESEARCH OBJECTIVES ........ 1
CHAPTER II. LIPASES FROM MAMMALS AND FISHES –
LITERATURE REVIEW (MANUSCRIPT 1) .............................................. 4
2.1. Abstract .................................................................................................. 4
2.2. Lipases from terrestrial species (predominantly mammalian) ......... 5
2.2.1. Introduction....................................................................................... 5
2.2.2. Catalytic mechanism ......................................................................... 7
2.2.3. The lipase gene family ...................................................................... 9
2.2.3.1. Pancreatic lipase and pancreatic lipase-related proteins 1 and 2
................................................................................................................ 9
2.2.3.2. Hepatic lipase and lipoprotein lipase ...................................... 10
2.2.3.3. Endothelial lipase and phosphatidylserine phospholipase A1 10
2.2.4. Pancreatic lipase ............................................................................. 11
2.2.4.1. Three dimensional structure .................................................... 11
2.2.4.2. Interfacial activation ............................................................... 12
2.2.5. Carboxyl ester lipase....................................................................... 13
2.2.6. General properties ........................................................................... 15
2.2.7. Other mammalian lipases ............................................................... 15
2.2.7.1. Hormone-sensitive lipase ........................................................ 15
2.2.7.2. Pre-duodenal lipases ............................................................... 16
2.2.7.3. Phospholipases ........................................................................ 17
2.2.8. Lipase inhibition ............................................................................. 17
2.2.9. Purification methods ....................................................................... 18
2.2.10. Assay methods .............................................................................. 19
x
2.2.11. Applications of lipases .................................................................. 19
2.2.11.1. Biocatalysis in organic solvents ............................................ 22
2.3. Lipases from aquatic species .............................................................. 23
2.3.1. Introduction..................................................................................... 23
2.3.1.1. Fish lipases .............................................................................. 26
2.3.1.2. Lipases from other marine animals ......................................... 27
2.3.2. Fish digestive lipases ...................................................................... 28
2.3.2.1. Rainbow trout (Oncorhynchus mykiss) ................................... 28
2.3.2.2. Oil sardine (Sardinella longiceps) .......................................... 30
2.3.2.3. Colipase (various sp.) .............................................................. 30
2.3.2.4. Spiny dogfish (Squalus acanthius) ......................................... 30
2.3.2.5. Atlantic cod (Gadus morhua) ................................................. 31
2.3.2.6. Atlantic salmon (Salmo salar) ................................................ 32
2.3.2.7. Yellowfin tuna (Thunnus albacares) ...................................... 33
2.3.2.8. Red sea bream (Pagrus major) ............................................... 33
2.3.2.9. Nile tilapia (Oreochromis niloticus) ....................................... 34
2.3.2.10. Grey mullet (Mugil cephalus) ............................................... 34
2.3.2.11. Lipase activity in unpurified fish extracts ............................. 34
2.3.2.11-1. In vivo feeding trials and analysis of gut content .......... 35
2.3.2.11-2. Leopard shark (Triakis semifasciata) ............................ 35
2.3.2.11-3. Milkfish (Chanos chanos) ............................................. 35
2.3.2.11-4. Arctic charr (Salvelinus alpinus) ................................... 36
2.3.2.11-5. Juvenile turbot (Scophthalmus maximus/Psetta maxima)
.......................................................................................................... 36
2.3.2.12. Overview-properties of purified and partially purified fish
digestive lipases ................................................................................... 37
2.3.3. Fish tissue lipases ........................................................................... 38
2.3.3.1. Red muscle .............................................................................. 38
2.3.3.2. Adipose tissue ......................................................................... 38
2.3.3.3. Liver ........................................................................................ 39
2.3.3.4. Gene expression ...................................................................... 39
2.3.4. Comparison of fish digestive lipases with those from mammals and
birds .......................................................................................................... 40
2.3.4.1. Substrate and fatty acid specificity ......................................... 40
2.3.4.2. Temperature optimum and stability ........................................ 40
2.3.4.3. pH optimum and stability ........................................................ 41
2.3.4.4. Calcium activation and inhibitors ........................................... 41
xi
2.3.4.5. Isoelectric point ....................................................................... 42
2.3.4.6. Molecular weight .................................................................... 42
2.3.4.7. Amino acid composition ......................................................... 42
2.3.4.8. Catalytic efficiency ................................................................. 44
2.3.5. Potential applications of digestive lipases from marine sources .... 44
2.4. Outlook on lipases from marine animals .......................................... 45
2.4.1. Impact of biotechnology ................................................................. 45
2.4.2. Future trends ................................................................................... 46
2.5. Acknowledgements .............................................................................. 48
CONNECTING STATEMENT 1 ................................................................. 49
CHAPTER III. PURIFICATION AND PROPERTIES OF DIGESTIVE
LIPASES FROM CHINOOK SALMON (ONCORHYNCHUS
TSHAWYTSCHA) AND NEW ZEALAND HOKI (MACRURONUS
NOVAEZELANDIAE) (MANUSCRIPT 2) .................................................. 50
3.1. Abstract ................................................................................................ 50
3.2. Introduction ......................................................................................... 51
3.3. Materials and methods ........................................................................ 53
3.3.1. Biological materials ........................................................................ 53
3.3.2. Chemicals ....................................................................................... 54
3.3.3. Sample preparation ......................................................................... 54
3.3.4. Purification protocol ....................................................................... 55
3.3.5. Final preparation of lipase samples ................................................ 57
3.3.6. Protein determination...................................................................... 58
3.3.7. Lipase assays .................................................................................. 58
3.3.8. Electrophoresis and zymographic analysis ..................................... 58
3.3.9. Acyl-chain specificity ..................................................................... 59
3.3.10. Effect of pH on the activity and stability ...................................... 60
3.3.11. Effect of temperature on the activity and stability ....................... 60
3.3.12. Enzyme kinetics ............................................................................ 61
3.3.12.1. Inhibition with paraoxon ...................................................... 61
3.3.13. Thermodynamic parameters ......................................................... 62
3.3.14. Effect of different bile salts on activity ........................................ 62
3.3.15. Effect of selected chemicals on activity/stability ......................... 62
3.3.16. Stability in organic solvents.......................................................... 63
3.3.17. Amino acid composition and N-terminal sequencing .................. 63
xii
3.4. Results and discussion ......................................................................... 64
3.4.1. Sample preparation ......................................................................... 64
3.4.2. Lipase stability during purification and choice of buffer ............... 64
3.4.3. Lipase assay .................................................................................... 65
3.4.4. Purification of salmon and hoki lipases .......................................... 65
3.4.5. Preparation of SDL and HDL ......................................................... 68
3.4.6. Electrophoresis and zymographic analysis ..................................... 69
3.4.7. Acyl-chain specificity ..................................................................... 72
3.4.8. pH optimum and pH stability ......................................................... 73
3.4.9. Temperature optimum and thermal stability................................... 75
3.4.10. Kinetic parameters ........................................................................ 77
3.4.10.1. Inhibition by an organophosphate.........................................78
3.4.11. Thermodynamic parameters ......................................................... 79
3.4.12. Lipase activity with different bile salts ......................................... 80
3.4.13. Activity/stability with selected chemicals .................................... 81
3.4.14. Stability in organic solvents.......................................................... 84
3.4.15. Amino acid analysis ...................................................................... 85
3.5. Conclusion ............................................................................................ 87
3.6. Acknowledgements .............................................................................. 88
CONNECTING STATEMENT 2 ................................................................. 89
CHAPTER IV. FLAVOUR DEVELOPMENT IN DAIRY CREAM
USING FISH DIGESTIVE LIPASES FROM CHINOOK SALMON
(ONCORHYNCHUS TSHAWYTSCHA) AND NEW ZEALAND HOKI
(MACRURONUS NOVAEZELANDIAE) (MANUSCRIPT 3) ................... 90
4.1. Abstract ................................................................................................ 90
4.2. Introduction ......................................................................................... 91
4.3. Materials and methods ........................................................................ 93
4.3.1. Substrate and other chemicals ........................................................ 93
4.3.2. Enzymes .......................................................................................... 93
4.3.3. Standardized lipase assay using cream ........................................... 94
4.3.4. Hydrolysis of cream lipids .............................................................. 94
4.3.5. Sensory analysis.............................................................................. 94
4.3.6. SPME-GC-MS analysis .................................................................. 95
4.3.7. Lipid extraction and separation of free fatty acids ......................... 95
4.3.8. Gas chromatography ....................................................................... 96
xiii
4.3.9. Statistical analysis ........................................................................... 96
4.4. Results and discussion ......................................................................... 96
4.4.1. Lipase assay and properties of the enzymes ................................... 96
4.4.2. Reaction progress curves for cream lipid hydrolysis ...................... 97
4.4.3. Analysis of volatile compounds by SPME-GC-MS ....................... 99
4.4.4. Free fatty acid composition .......................................................... 103
4.5. Conclusion .......................................................................................... 106
4.6. Acknowledgements ............................................................................ 106
CONNECTING STATEMENT 3 ............................................................... 107
CHAPTER V. HYDROPHOBIC IMMOBILIZATION OF A BILE SALT
ACTIVATED LIPASE FROM CHINOOK SALMON
(ONCORHYNCHUS TSHAWYTSCHA) (MANUSCRIPT 4) ................... 108
5.1. Abstract .............................................................................................. 108
5.2. Introduction ....................................................................................... 109
5.3. Materials and methods ...................................................................... 111
5.3.1. General .......................................................................................... 111
5.3.2. Lipase extraction and purification with p-ABA-cellulose ............ 111
5.3.3. Immobilization .............................................................................. 112
5.3.4. Trypsin activity assay ................................................................... 113
5.3.5. Lipase activity assays during immobilization............................... 113
5.3.6. Immobilized lipase activity against natural substrates ................. 114
5.3.7. Protein determination.................................................................... 114
5.3.8. Properties of free and immobilized lipase .................................... 114
5.4. Results and discussion ....................................................................... 115
5.4.1. Lipase immobilization on hydrophobic supports ......................... 115
5.4.2. Immobilized lipase activity against natural lipid substrates ......... 118
5.4.3. Effect of immobilization on pH and thermal properties and bile salt
requirement of Chinook salmon lipase ................................................... 120
5.5. Conclusions ........................................................................................ 123
5.6. Acknowledgements ............................................................................ 124
CHAPTER VI. GENERAL DISCUSSION AND CONCLUSION .......... 125
BIBLIOGRAPHY ........................................................................................ 131
APPENDIX I. DETAILS OF METHODS AND RESULTS AND
ADDITIONAL EXPERIMENTS FOR CHAPTER III ............................ 156
xiv
APPENDIX II. DETAILS OF METHODS AND RESULTS AND
ADDITIONAL EXPERIMENTS FOR CHAPTER V ............................. 177
..............................................................................................................................
xv
LIST OF ABBREVIATIONS
ATP…………………………………………………Adenosine-5'-triphosphate
BA-p-NA.....................................................Benzoyl-DL-arginine-p-nitroanilide
CAL-B....................................................................Candida antarctica Lipase B
cAMP………………………………………..Cyclic adenosine monophosphate
cDNA……………………………………………………Complementary DNA
CEL……………………………………………………….Carboxyl ester lipase
CLA…………………………………………………...Conjugated linoleic acid
DEAE……………………………………………………….Diethylaminoethyl
DG…………………………………………………………….........Diglyceride
DHA……………………………………………………..Docosahexaenoic acid
DMSO………………………………………………………Dimethyl sulfoxide
EDAC……...N-Ethyl-N′-(3-dimethylaminopropyl)carbodiimide hydrochloride
EDTA…………………………………………Ethylenediaminetetraacetic acid
EL……………………………………………………………Endothelial lipase
EPA……………………………………………………...Eicosapentaenoic acid
FA…………………………………………………………………….Fatty acid
FAEE……………………………………………………...Fatty acid ethyl ester
FFA................................................................................................Free fatty acid
GC………………………………………………………Gas chromatograph(y)
GC-MS………………Gas chromatography combined with mass spectroscopy
GF…………………………………………………………………Gel filtration
HDL……………………………High density lipoprotein/Hoki digestive lipase
HL………………………………………………………………..Hepatic lipase
HPLC……………………………….High performance liquid chromatography
HSL…………………………………………………..Hormone-sensitive lipase
IDL……………………………………………Intermediate density lipoprotein
IE………………………………………………………………….Ion exchange
IEF…………………………………………….....................Isoelectric focusing
LDL................................................................................Low density lipoprotein
Lew.....................................................................................Lewatit VP OC 1600
LPL…………………………………………………………..Lipoprotein lipase
MG……………………………………………………………...Monoglyceride
xvi
MUF-butyrate……………………………….......4-methylumbelliferyl butyrate
MUF-oleate………………………………………..4-methylumbelliferyl oleate
Mw...........................................................................................Molecular weight
MWCO.........................................................................Molecular weight cut-off
Novo 435.......................................................................................Novozym 435
Oct-S..............................................................................Octyl-Sepharose CL-4B
p-ABA-cellulose…………………………...…...p-aminobenzamidine-cellulose
PAGE...........................................................Polyacrylamide gel electrophoresis
PBE20……………………………………………….100 mM phosphate buffer
(pH 7.8, containing 2 mM benzamidine, 1 mM EDTA and 20% (w/v) glycerol
PEG 1000……Polyethylene glycol with average molecular weight of 1000 Da
PGE.........................................................................................Pregastric esterase
PL……………………………………………………………..Pancreatic lipase
PLRP1………………………………………Pancreatic lipase-related protein 1
PLRP2………………………………………Pancreatic lipase-related protein 2
PMSF………………………………………….Phenyl methyl sulfonyl fluoride
p-NA………………………………………………………….......p-nitroaniline
p-NP................................................................................................p-nitrophenol
p-NPA…………………………………………………….p-nitrophenyl acetate
p-NPB……………………………………………….......p-nitrophenyl butyrate
p-NPC…………………………………………………….p-nitrophenyl caprate
p-NPP..............................................................................p-nitrophenyl palmitate
PUFA………………………………………………..Polyunsaturated fatty acid
QAE…………………………………………………….Quaternary aminoethyl
SDL…………………………………………………….Salmon digestive lipase
SDS..............................................................................Sodium dodecyl sulphate
SP…………………………………………………………………...Sulfopropyl
SPME......................................................................Solid phase micro extraction
TBE…………..25 mM Tris-HCl, 2 mM benzamidine and 1 mM EDTA buffer
TBE20...........................................................................100 mM Tris-HCl buffer
(pH 8.0, containing 2 mM benzamidine, 1 mM EDTA and 20% (w/v) glycerol
TG………………………………………………………………….Triglyceride
TLC...........................................................................Thin layer chromatography
U...................................................................................................Unit of activity
xvii
VLDL.....................................................................Very low density lipoprotein
xviii
LIST OF TABLES
Table 2.1 Properties of purified or partially purified fish digestive lipases. .... 37
Table 2.2 Amino acid composition of several purified CELs.......................... 43
Table 3.1 Purification scheme for Chinook salmon digestive lipase ............... 66
Table 3.2 Extinction coefficient of p-NP as a function of pH ......................... 73
Table 3.3 Thermodynamic parameters for lipase-catalyzed hydrolysis of p-
NPC at 35°C ..................................................................................................... 81
Table 3.4 Effect of selected chemicals on the stability of Chinook salmon and
hoki digestive lipases ....................................................................................... 82
Table 3.5 Effect of organic solvents on the stability of Chinook salmon and
hoki digestive lipases ....................................................................................... 85
Table 3.6 Amino acid composition of Chinook salmon and hoki digestive
lipases ............................................................................................................... 86
Table 4.1 Characteristics of the four lipase preparations ................................. 97
Table 4.2 Volatile organic compounds in lipase-treated creams and Parmesan
cheese ............................................................................................................... 99
Table 4.3 Sensory characteristics of the creams treated with different lipase
preparations .................................................................................................... 101
Table 5.1 Activities (U) against BA-p-NA (measurement of trypsin activity)
for the purification of Chinook salmon lipase on p-ABA-cellulose .............. 115
Table 5.2 Lipase activities (U) against p-NPP and tributyrin for the
immobilization of Chinook salmon lipase on two hydrophobic supports (Oct-S
and Lew) ........................................................................................................ 116
Table 5.3 Activity (U) of the immobilized Chinook salmon lipases and Novo
435 against ghee and fish oil emulsions......................................................... 119
Table A1.1 Effect of benzamidine concentration and slightly acidic pH
extraction on relative lipase activity (%) in the crude extract from hoki pyloric
ceca, with storage at 0°C for 72 h .................................................................. 160
Table A1.2 Effect of bile salts and glycerol on relative lipase activity (%) in
the crude extract from hoki pyloric ceca, with storage at 0°C for 24 h ......... 161
Table A1.3 Effect of EDTA and Na cholate concentration on relative lipase
activity (%) in the crude extract from hoki pyloric ceca, with storage at 0°C for
72 h ................................................................................................................. 161
xix
Table A1.4 Comparison of specific lipase activity (U/mg) in the crude extract
from Chinook salmon pyloric ceca obtained with TBE20 and PBE20 buffers,
with storage at 4°C for 96 h ........................................................................... 162
Table A1.5 Lipase activities of crude extract from hoki pyloric ceca obtained
with the titrimetric assay ................................................................................ 163
Table A1.6 Purification scheme for hoki digestive lipase ............................. 166
Table A1.7 Preparation of SDL and HDL ..................................................... 166
Table A2.1 a) Protein balance for the immobilization of Chinook salmon
lipase on Oct-S b) Protein balance for the immobilization of Chinook salmon
lipase on Lew ................................................................................................. 178
xx
LIST OF FIGURES
Fig. 2.1. Mechanism of triglyceride cleavage by lipase. ................................... 8
Fig. 3.1. Purification steps for salmon and hoki digestive lipases. .................. 55
Fig. 3.2. A representative elution profile of salmon lipase on Sephacryl S-300
HR…………………………………………………………………………….68
Fig. 3.3. Zymograms, (a) purified hoki and salmon digestive lipases (3.6 µg
HDL and 1.7 µg SDL), (b) 4.2 µg SDL (left) and 300 µg salmon crude extract
(right) ............................................................................................................... 69
Fig. 3.4. 7.5% acrylamide native-PAGE of purified hoki and salmon digestive
lipases. .............................................................................................................. 70
Fig. 3.5. 12% acrylamide SDS-PAGE, (a) 4.5 µg purified salmon digestive
lipase, (b) 10 µg purified hoki digestive lipase……………………………....71
Fig. 3.6. Acyl-chain specificity of Chinook salmon lipase with p-nitrophenyl
esters. ............................................................................................................... 73
Fig. 3.7. Activity of Chinook salmon (—) and hoki (– – –) lipase against p-
NPC as a function of pH. ................................................................................. 74
Fig. 3.8. Effect of pH on the stability of Chinook salmon (—) and hoki (– – –)
lipase.. .............................................................................................................. 75
Fig. 3.9. Activity of Chinook salmon (—) and hoki (– – –) lipase against p-
NPC as a function of temperature. ................................................................... 76
Fig. 3.10. Effect of temperature on the stability of Chinook salmon (—) and
hoki (– – –) lipase. ........................................................................................... 77
Fig. 3.11. Michaelis-Menten plot for paraoxon inhibition of Chinook salmon
(—) and hoki (– – –) lipases............................................................................. 79
Fig. 3.12. Arrhenius plot for the hydrolysis of p-NPC by Chinook salmon
lipase.................................................................................................................80
Fig. 3.13. Arrhenius plot for the hydrolysis of p-NPC by hoki lipase.............81
Fig. 3.14. Effect of bile salts on salmon lipase activity against p-NPC........... 82
Fig. 4.1. Changes in pH (—) and FFA% (– – –) for cream when treated with
four different lipase preparations: a) Palatase®; b) calf PGE; c) salmon; and d)
hoki .................................................................................................................. 98
Fig. 4.2. Peak area intensity for butanoic (C4:0) and hexanoic (C6:0) acids
after SPME-GC-MS analysis of creams treated with different lipases... ....... 102
xxi
Fig. 4.3. a) Changes in FFA composition of cream after treatment with
different lipases b) Changes in butanoic, hexanoic and octanoic acid
concentrations in cream after treatment with different lipases. ..................... 104
Fig. 5.1. Activity of free (– – –) and immobilized (—) Chinook salmon lipase
against tributyrin as a function of pH.. .......................................................... 121
Fig. 5.2. Activity of free (– – –) and immobilized (—) Chinook salmon lipase
against tributyrin as a function of temperature. ............................................. 122
Fig. 5.3. Effect of temperature on the stability of free (– – –) and immobilized
(—) Chinook salmon lipase. .......................................................................... 123
Fig. A1.1. Standard curve for the Lowry protein assay. ................................ 165
Fig. A1.2. A representative elution profile of hoki lipase on Sephacryl S-300
HR .................................................................................................................. 167
Fig. A1.3. IEF-PAGE of gel filtration concentrates: purified hoki (lane 1, 6.5
µg) and salmon digestive lipase (lanes 5 and 6, 7.5 and 2.6 µg, respectively)
........................................................................................................................ 167
Fig. A1.4. Acyl-chain specificity of hoki lipase with p-nitrophenyl esters.. . 168
Fig. A1.5. Effect of bile salts on hoki lipase activity against p-NPC. ........... 168
Fig. A1.6. Michaelis-Menten plot for the hydrolysis of p-NPC by Chinook
salmon lipase at 35°C.. ................................................................................... 169
Fig. A1.7. Michaelis-Menten plot for the hydrolysis of p-NPC by hoki lipase
at 35°C. .......................................................................................................... 169
Fig. A1.8. Lineweaver-Burke plot for the hydrolysis of p-NPC by Chinook
salmon lipase at 35°C. .................................................................................... 170
Fig. A1.9. Hanes plot for the hydrolysis of p-NPC by Chinook salmon lipase
at 35°C.. ......................................................................................................... 170
Fig. A1.10. Lineweaver-Burke plot for the hydrolysis of p-NPC by hoki lipase
at 35°C.. ......................................................................................................... 171
Fig. A1.11. Hanes plot for the hydrolysis of p-NPC by hoki lipase at 35°C.
........................................................................................................................ 172
Fig. A2.1. 12.5% acrylamide SDS-PAGE of fractions from the immobilization
of Chinook salmon lipase on Oct-S and Lew ................................................ 180
Fig. A2.2. TLC of the products of hoki oil hydrolysis with immobilized
salmon lipases and Novo 435 in hexane.. ...................................................... 182
1
CHAPTER I. INTRODUCTION AND RESEARCH OBJECTIVES
Much of the current knowledge about lipases stems from research conducted
on mammals, microorganisms and plants. The digestion of lipids in fish and
the properties of their lipase systems are at this time not well understood. The
research carried out to date on several marine fish species (including anchovy,
sardine, striped bass, pink salmon, leopard shark, dogfish, cod, red sea bream
and Atlantic salmon) indicates that fat digestion in these animals is restricted
to, or mostly carried out by homologues of mammalian bile salt activated
lipases (also known as carboxyl ester lipases; Patton et al. 1975; Patton et al.
1977; Mukundan et al. 1985; Rasco and Hultin 1988; Gjellesvik et al. 1992;
Gjellesvik et al. 1994; Iijima et al. 1998). However, it remains to be
established whether this applies to fish in general. There is very little (if any)
published information on the lipase systems of New Zealand commercial fish
species.
Cold water fish and shellfish enzymes, in general, have been shown to differ
from the majority of corresponding mammalian enzymes with the same
functions (e.g. lipases and serine proteases) in that they exhibit significant
reaction rates at lower temperatures, generally have lower thermal stabilities,
respond differently to changes in pH and possess dissimilar kinetic and
thermodynamic characteristics. The substrate and FA specificities further
contribute to the unique features of seafood lipases (Lopez-Amaya and
Marangoni 2000b).
As most of the commercially utilized lipases originate from microbial and a
few from limited mammalian sources, the lipases from an alternative source
(i.e. the marine environment) have the potential to expand the available
activities and specificities. Fish are a prospective source of lipases, with the
digestive organs of these animals being an abundant and underutilized by-
product of fish processing. Because of the environment in which different fish
live (e.g. at low temperatures) and in response to their diets, the lipases from
some fish may have higher catalytic efficiencies at lower temperatures than
either their microbial or mammalian counterparts. There is potential to exploit
the activity of fish lipases to reduce energy costs in industrial processes, to
2
purify specific FAs, to deliver structured lipids for nutraceutical use, and to
produce particular flavours in food products.
Despite their potential, fish lipases present challenges for extraction and use
that have so far made their commercial exploitation unattractive. The high
activity of some fish lipases at relatively low temperatures appears to be
coupled with a conformational instability that makes extraction and
stabilisation very difficult. My research aims to investigate the activities of
lipases from two commercial fish species in New Zealand, Chinook salmon
(Oncorhynchus tshawytscha) and New Zealand hoki (Macruronus
novaezelandiae) and to overcome the stability issues to allow any novel
activities of these enzymes to be exploited. Immobilization will be
investigated as one way to overcome the instability of the enzymes
(Fernandez-Lafuente et al. 1998; Hanefeld et al. 2009). Fish lipases are more
expensive to extract than their microbial counterparts but immobilization and
repeat uses may make them an economically viable option in industrial
processing.
This research undertaking is therefore motivated by: the paucity of
information about lipases from aquatic (marine) sources, especially New
Zealand commercial fish species – enzyme characteristics and lipase-type
classification; the potential applications of fish lipases for lipid modifications
in foods and during bioindustrial processing; and the need to immobilize fish
lipases for continuous and large scale industrial applications.
Research objectives
1. To develop a quick, sensitive, specific and reproducible activity assay
for fish digestive lipases
2. Purification of Chinook salmon and New Zealand hoki digestive
lipases
- to optimize the starting material and extraction buffer for highest yield
of lipase activity
3
- to develop methods of maintaining lipase stability (e.g. modifications
to buffer composition, a treatment that results in improved lipase
stability)
- to design a purification sequence through adaptation/modification of
existing techniques
3. Characterization of Chinook salmon and hoki digestive lipases
- to determine purity, molecular weight and isoelectric point
- to study the effects of pH and temperature on activity and stability
- to examine the effects of several activators and inhibitors
- to determine kinetic and thermodynamic properties
- to determine amino acid composition and N-terminal sequence
- to classify the lipases with reference to mammalian digestive lipase
types
4. Potential applications of Chinook salmon and hoki digestive lipases
- to test the performance of the digestive lipases with respect to lipid
modifications
- to demonstrate a potential application (e.g. in food products)
5. Immobilization of a fish digestive lipase
- to immobilize one digestive lipase on a suitable support (e.g.
hydrophobic support)
- to demonstrate the functionality of the immobilized lipase with several
lipid substrates in either aqueous (e.g. emulsion) or non-aqueous (e.g.
organic solvent) medium
4
CHAPTER II. LIPASES FROM MAMMALS AND FISHES –
LITERATURE REVIEW (MANUSCRIPT 1)
Published in Reviews in Fisheries Science, 2009, 17:18-40
Authors: Ivan Kurtovic1,2
, Susan N. Marshall2, Xin Zhao
1 and Benjamin K.
Simpson3*
Affiliations:
1Department of Animal Science, McGill University (Macdonald Campus)
21,111 Lakeshore Road, Ste. Anne de Bellevue (QC) Canada H9X 3V9
2Processed Foods Group – Seafood Research Unit, Crop & Food Research
Limited, Port Nelson, Nelson, New Zealand
3Department of Food Science and Agricultural Chemistry, McGill University
(Macdonald Campus): 21,111 Lakeshore Road, Ste. Anne de Bellevue (QC)
Canada H9X 3V9
*Corresponding Author: Benjamin K. Simpson
Department of Food Science and Agricultural Chemistry, McGill University
(Macdonald Campus): 21,111 Lakeshore Road, Ste. Anne de Bellevue (QC)
Canada H9X 3V9
E-mail address: [email protected]
2.1. Abstract
Lipases are a broad family of enzymes that catalyze the hydrolysis of ester
bonds in substrates such as triglycerides, phospholipids, cholesteryl esters and
vitamin esters. Lipases are receiving increasing interest due to their effects on
the quality of food products (e.g. the quality of post-harvest seafoods), and
their actual and potential applications in modified foods and industrial
processes. Lipases that catalyse specific reactions and that are active at
particular conditions of pH and temperature to suit the requirements of
industrial processes are of particular interest. This review focuses on lipases
that display predominantly triacylglycerol hydrolase activity. Section 2.2
presents an overview of lipases from terrestrial organisms, and the lipase gene
family members. Due to their unusual physiology, diet and habitat, fish lipases
may demonstrate novel activities that have potential applications for
5
bioindustrial catalysis. Section 2.3 discusses lipases from fish and compares
them with lipases from mammals (and birds). Purification strategies and
properties of the isolated enzymes are reviewed in detail.
Keywords: pancreatic lipase, carboxyl ester lipase, triglyceride, bile salt, fish,
protein engineering
2.2. Lipases from terrestrial species (predominantly mammalian)
2.2.1. Introduction
Lipases can be broadly defined as enzymes that catalyze the hydrolysis of
ester bonds in substrates such as triglycerides (TGs), phospholipids,
cholesteryl esters and vitamin esters (Wong and Schotz 2002). Although
lipases are water-soluble proteins, their substrates are insoluble in a purely
aqueous system. Within the above classification, the so called ‗true lipase‘ (EC
3.1.1.3 triacylglycerol lipase) is distinguished from other lipolytic enzymes
such as phospholipases, sterol esterase and retinyl-palmitate esterase. Lipid
esterases have traditionally been separated from lipases on the basis that their
substrates have higher solubility in water (Anthonsen et al. 1995).
In the early days of lipase research, mammals such as rat, sheep, cow and pig
were studied owing to a need to understand lipid digestion in humans. Some
relatively early studies include those of Bier (1955) and Sarda et al. (1958) on
porcine pancreatic lipase (PL); Gidez (1968) on rat PL; Julien et al. (1972) on
bovine PL and colipase; and Canioni et al. (1975) on ovine PL. The first
published lipase sequence was for porcine pancreatic triacylglycerol lipase (De
Caro et al. 1981). Mammalian digestive, systemic and tissue lipases continue
to be researched very extensively, especially for medical purposes; for
instance, to elucidate their roles in the process of atherosclerosis and obesity
(e.g. Goldberg 1996; Hui and Howles 2002; Jansen et al. 2002). For a long
time, the pancreas and serous glands of ruminants, especially young (kid, calf
and lamb) and the pancreas of pigs have been important sources of lipases for
flavour development in cheeses and other dairy products (see section 2.2.11).
6
Microbial lipases have received most of the attention in recent lipase research.
They have many commercial applications as a consequence of the
convenience of their production, ease of genetic manipulation and a huge
diversity in characteristics and specificities. A few examples are the studies of
Hiol et al. (2000), Kojima and Shimizu (2003) and Lima et al. (2004). A
comprehensive overview of microbial lipase production, purification,
characterization and applications was provided by Sharma et al. (2001). The
microbial lipases are the most significant commercially produced lipases.
Lipases have also been researched and sourced from plants, in which they are
widely distributed. Compared with mammalian and microbial lipases, less
information on plant lipases is available. Oilseeds, oily fruits and cereal grains
have been the most common sources for the study and extraction of plant
lipases. Besides the triacylglycerol lipase (EC 3.1.1.3), plants also contain
various phospholipases as well as glycolipase and sulpholipase. The
attractiveness of plant lipases is due to their special properties (e.g. cleavage of
all three fatty acids from TGs by oat lipase), unique substrate specificities and
abilities to synthesise ‗designed‘ esters (e.g. rape-seed lipase esterifies fatty
acids to primary alcohols only). Many special features of lipolytic activity
from plant extracts have been attributed to phospholipases (e.g. phospholipase
D from cabbage can be used to synthesize phosphatidylglycerols, one kind of
artificial lung surfactant). Understanding the properties of plant lipases is
important for maintaining the freshness of oilseeds, cereals and oily fruits, and
for the production of high-quality edible oils (Mukherjee and Hills 1994;
Mukherjee 2002).
Lipases belong to the α/β hydrolase-fold family of enzymes (Schrag and
Cygler 1997) that also includes esterases, thioesterases, certain proteases and
peroxidases, dehalogenases and several other intracellular hydrolases.
Members of this super-family are characterized by a certain α-helix and β-
sheet topology (e.g. a fan-like pattern of the centrally located, mostly parallel β
sheet). The catalytic triad of the α/β hydrolase fold proteins consists of a
nucleophile (serine), an acid (aspartic or glutamic) and a histidine group,
differing from that in serine proteases only in the order in which the catalytic
7
residues occur in the amino acid sequence (Wang and Hartsuck 1993; Wong
and Schotz 2002).
The different lipase groups exhibit low degrees of similarity with each other.
Only four major structural, functional or sequence homologies have been
recognized within almost all of the lipases: (i) the consensus sequence GxSxG
around the active site serine, where x can be any amino acid; (ii) the strand-
helix motif also around the active site serine; (iii) the buried active site
covered by a region known in different lipases as active site loop, lid domain
or flap. Although common to the lipases, this region varies extensively – it
consists of only 5 residues in guinea pig pancreatic lipase-related protein 2, 12
residues in human carboxyl ester lipase (CEL) and 23 in human pancreatic
lipase (PL). Residues in the lid domain play an important role in determining
the substrate selectivity of lipases; (iv) the order of the catalytic triad residues
– Ser…Asp/Glu…His (Svendsen 1994; Carriere et al. 1998; Hui and Howles
2002).
2.2.2. Catalytic mechanism
The proposed catalytic mechanism of lipases, summarized in Fig. 2.1, is
chemically analogous to that of the serine proteases, based on the presence of
a common catalytic triad. The difference between the two catalytic systems
lies in the handedness of the tetrahedral, hemiacetal intermediate (Winkler and
Gubernator 1994). The last step in the process, release of a fatty acid from the
active site cleft, is of special importance since a too tightly bound fatty acid
will inhibit the enzyme. Petersen et al. (2001) proposed a model for product
release called ‗the electrostatic catapult model‘. According to this model,
following the ester bond cleavage, the fatty acid is rapidly ejected from the
active site cleft which exhibits a negative electrostatic potential around neutral
and basic pH values.
8
Fig. 2.1. Mechanism of triglyceride cleavage by lipase (adapted from Winkler
and Gubernator 1994).
There are several categories of lipase specificity. Substrate-specific lipases
differentiate between esters such as TGs, diglycerides, monoglycerides, and
phospholipids. Fatty acid-specific lipases show preference for particular fatty
acids or a class of fatty acids (e.g. short-chain, polyunsaturated, etc). Lipases
can also be regioselective (i.e. show positional specificity) in that they
distinguish between the external, primary (sn-1 and sn-3 positions) and
internal, secondary (sn-2 position) ester bonds. Many microbial lipases, as
well as gastric and PL are specific for the external ester bonds. True sn-2
selective lipases are very rare. Most of the lipases show at least partial
stereospecificity, i.e. a preference for either the sn-1 or sn-3 position of TG.
The highest level of specificity that lipases exhibit is enantioselectivity – the
ability to differentiate between enantiomers of chiral molecules. This ability in
certain microbial lipases has been used recently for producing pure chiral
isomers during chemical synthesis (Matori et al. 1991; Anthonsen et al. 1995;
Hou 2002).
9
Kinetic models of lipase activity are complicated by the presence of several
phases. The most accepted model is that the lipase exists in two conformations
(inactive and active) in solution. In aqueous solution the equilibrium lies well
towards the inactive form and the enzyme displays very low activity towards
soluble substrates. At the lipid-water interface, however, the equilibrium shifts
towards the active form and insoluble substrates are hydrolyzed by the
enzyme. The active forms of the enzyme that are in solution and adsorbed to
the interface surface may have different conformation and there is a possibility
that several conformations of each active form exist (Martinelle and Hult
1994).
2.2.3. The lipase gene family
Initially made up of pancreatic lipase (PL), hepatic lipase (HL) and lipoprotein
lipase (LPL), the family has expanded and now includes PL-related proteins 1
and 2 (PLRP1 and PLRP2), phosphatidylserine phospholipase A1 and
endothelial lipase. Members of this family predominantly hydrolyze TGs,
diglycerides and phospholipids, and can be thought of as glycerol-sn-1-fatty
acid hydrolases. Many are glycoproteins (Choi et al. 2002; Wong and Schotz
2002). Lipase gene family members are typically 450-500 residues long and
have molecular weights in the approximate range of 50 to 70 kDa, depending
on the degree of glycosylation.
2.2.3.1. Pancreatic lipase and pancreatic lipase-related proteins 1 and 2
PL, also known as colipase-dependant lipase, is the main digestive lipolytic
enzyme in higher vertebrates and is secreted by the exocrine pancreas. The
enzyme has strong preference for acylglycerides over other lipids and may
contribute to the hydrolysis of retinyl esters in vivo. PLRP1 has little known
activity. PLRP2s are generally catalytically active against TGs, but can show
activity against a variety of lipid substrates, depending on the species. The
expression of PLRP1 and 2 differs between vertebrate species and their
function is still to be elucidated (Crenon et al. 1998; Lowe 2002).
10
2.2.3.2. Hepatic lipase and lipoprotein lipase
Hepatic lipase (HL) and lipoprotein lipase (LPL) are important in the
metabolism of circulating lipoproteins. The two enzymes are evolutionarily
more closely related to each other than either is to PL (Hide et al. 1992). LPL
may have evolved before HL, since the activity of the former is present in fish,
amphibians and birds. In contrast, these animals show very low levels or
complete absence of HL-like activity (Lindberg and Olivecrona 1995). HL is
synthesized by hepatocytes and functions extracellularly, attached to heparan-
sulphate proteoglycans at the surface of hepatocytes and endothelial cells of
liver sinusoid capillaries. The enzyme is also present (most probably via
circulation) in adrenals and ovaries. HL displays TG, diglyceride and
monoglyceride lipase, and phospholipase A1 activities. It functions to maintain
intracellular lipid homeostasis via its roles in HDL metabolism and reverse
cholesterol transport, IDL/LDL metabolism, and postprandial chylomicron-
remnant clearing. It may also assist with bridging of lipoproteins to lipoprotein
receptors or heparan sulphates on hepatocytes. Heparin strongly stimulates the
activity as well as expression of HL, while cholesterol inhibits its activity
(Connelly 1999; Jansen et al. 2002; Perret et al. 2002).
LPL has a high preference for TGs, hydrolyzing these from circulating
chylomicrons and VLDLs. The enzyme is produced in many tissues, with
adipose and muscle tissues containing the highest levels. It is bound to
capillary endothelium through heparan-sulphate proteoglycans. LPL is
inhibited by protamine, and by molar concentrations of NaCl, and requires a
cofactor (apolipoprotein C-II) for activity. Besides the central roles in
chylomicron catabolism and initiation of conversion of VLDL to IDL, LPL
may have several roles in subendothelial space including a non-enzymatic role
as a ligand, similar to HL‘s bridging ability (Goldberg 1996; Lopez-Amaya
and Marangoni 2000b; Merkel et al. 2002; Wong and Schotz 2002).
2.2.3.3. Endothelial lipase and phosphatidylserine phospholipase A1
The two most recent additions to the lipase gene family, endothelial lipase
(EL) and phosphatidylserine phospholipase A1, both show phospholipase A1
activity, with the latter having an absolute preference for phosphatidylserine
and lysophosphatidylserine. Apart from being produced by platelets, little else
11
is known about phosphatidylserine phospholipase A1. EL has been detected in
several tissues such as liver, lung, thyroid and reproductive organs, but differs
from HL and LPL in that its synthesis and function (intra- and extracellular)
both occur at the same place (endothelial cells). The expression of EL is
regulated by physical forces and cytokines and the lipase may be involved in
HDL metabolism; however, the real physiological role of this enzyme is not
yet understood (Choi et al. 2002; Wong and Schotz 2002).
2.2.4. Pancreatic lipase
2.2.4.1. Three dimensional structure
The structure of human PL was revealed in 1990 by Winkler et al. (1990) and
was the first published 3-D organization of any PL. The 449 residue-long
enzyme has two distinct domains: an N-terminal catalytic domain with the α/β
hydrolase fold structure, and a smaller, C-terminal ancillary domain that has a
β-sandwich structure. The following elements in the N-terminal domain are of
particular functional significance: the active site (Ser, Asp, His triad)
containing the oxyanion hole, the lid domain covering the active site in the
enzyme‘s closed conformation, β5 and β9 loops that form contacts with the lid
domain and assist in the enzyme‘s interaction with the lipid-water interface,
and Ca2+
-binding and glycosylation sites. The C-terminal domain binds
colipase and contains the β5′ loop that is likely to participate in lipid binding.
The surface of colipase, lid domain, β5 and β9 loops form a large hydrophobic
plateau predicted to interact with the interface. Other members of the lipase
gene family like LPL and HL have very similar structure to PL, but instead of
binding colipase, the C-terminal domain in the former two enzymes contains
regions that bind to heparin, apolipoprotein C-II (in LPL) and a receptor
(Winkler et al. 1990; Winkler and Gubernator 1994; Wong and Schotz 2002).
The structural basis for the substrate selectivity of PL and other members of
the lipase gene family is likely to be encoded in the degree of hydrophobicity
of the surface loops (β5 and β9 loop and lid domain) surrounding the active
site (Carriere et al. 1998).
12
2.2.4.2. Interfacial activation
The interfacial activation of PL has been well documented – when the
substrate concentration exceeds its solubility in water, a dramatic increase is
seen in the lipase activity. While the esterases display a hyperbolic, Michaelis-
Menten relationship between the enzyme activity and substrate concentration,
in lipases this relationship is sigmoidal (Anthonsen et al. 1995). Upon binding
to the emulsified substrate at the lipid-water interface, the activity of PL
greatly increases due to the conformational change in the enzyme: the lid
domain ‗peels back‘, exposing the active site and the β5 loop also assumes a
new orientation that further facilitates improved substrate access to the active
site (van Tilbeurgh et al. 1993; van Tilbeurgh et al. 1999). Bile salts and
colipase must both be present if the enzyme is to transform to optimal
conformation for activity. Bile salts are thought to anchor the enzyme to the
surface of the interface and may alter the properties of the interface by
preventing compounds with emulsifying capability (e.g. proteins and
oligosaccharides) from binding to the interface, which could potentially
interfere with lipase-substrate interactions. Bile salts can also bind and carry
away fatty acids thus preventing product inhibition (Lowe 2002). However, in
the absence of colipase, bile salts above their critical micellar concentration
are inhibitory to PL activity. This is due to their coating of the substrate
globules at the interface, which hampers lipase-TG interaction (Carriere et al.
1998). Critical micellar concentration ranges from < 1 mM to > 4 mM,
depending on the bile salt (or its conjugate) (Borgstrom and Erlanson 1973).
Colipase interacts with the bile salt (micelles) to allow PL to adsorb at bile
salt-covered interfaces and stabilizes the lid domain of PL in an open and
active conformation via hydrogen bonds (Carriere et al. 1998; Lowe 2002).
An important question arises from the above discussion: what exactly triggers
the movement of the lid domain in the lipases? There is no current consensus
(Carriere et al. 1998). Originally, the interface itself or the emulsified
substrate, in the presence of colipase, was thought to be the trigger. This mode
of activation is still supported by some researchers (references in Carriere et
al. 1998; Lowe 2002). Alternatively, Hermoso et al. (1996) and Pignol et al.
(1998) concluded that PL activation is not an interfacial phenomenon but that
a detergent micelle, bile salt micelle or mixed micelle (bile salt, phospholipid
13
and a detergent) together with colipase mediate PL activation (i.e. lid opening)
in aqueous media. In that model, the micelle is bound to the concave face
created by colipase and the distal tip of the PL C-terminal domain. In a study
on the activity of a true lipase against soluble, short-chain substrates (i.e.
esterase activity), Nini et al. (2001) reported that small multimolecular
substrate aggregates in the aqueous phase trigger the structural transition to the
active conformation in many microbial lipases. In that case, the enzyme
activation could be due to the presence of the small lipid-water interface
created by the substrate aggregates. The work of Nini et al. thus also highlights
the difficulties in distinguishing true lipases from esterases. Furthermore,
Ferrato et al. (1997) reviewed many interfacial activation studies and found
that reaction conditions and physico-chemical parameters have a significant
impact on the interfacial activation process. They caution against the use of
interfacial activation as a defining feature of lipases.
2.2.5. Carboxyl ester lipase
The pancreas produces three major lipolytic enzymes – PL, phospholipase A2
and carboxyl ester lipase (CEL). Of the three, CEL has by far the broadest
substrate specificity. The lipase is capable of hydrolyzing tri-, di- and
monoglycerides, cholesteryl esters, esters of vitamins A and E, aryl esters,
phospholipids, lysophospholipids, wax esters and ceramide. Owing to its wide
substrate specificity, CEL has in the past been referred to as non-specific
lipase, cholesterol esterase, carboxylesterase, lysophospholipase and most
often as bile salt-activated, -stimulated or -dependant lipase. The last three
names reflect the enzyme‘s requirement for primary (7α hydroxylated) bile
salts for activity against bulkier substrates, like medium and long chain TGs
and diglycerides, cholesteryl esters and phospholipids. The enzyme has basal
activity against shorter chain TGs (≤ 8 carbon fatty acids), monoglycerides and
lysophospholipids (Wang and Hartsuck 1993; Hui and Howles 2002).
CEL has been isolated from many mammals (Wang and Hartsuck 1993; Hui
and Howles 2002) as well as from fish (see section 2.3.2). This lipase is
synthesized primarily in the pancreas and lactating mammary gland (except in
the mammary gland of cow and rat), but can also be found in liver,
14
macrophages, endothelial cells and heart. CEL has undisputed roles in
enhanced lipid digestion in the infant (where PL activity is low and CEL is
provided in milk) and hydrolysis of dietary cholesteryl esters, as it is the only
enzyme secreted by the pancreas capable of doing so. Other roles of CEL are
still not clear. Due to its broad fatty acid specificity it is likely to assist PL in
digestion of certain glycerides, most notably those containing long-chain
PUFAs. CEL is also likely to have an important role in hydrolyzing
monoglycerides. Phospholipid digestion and absorption, vitamin A and E
absorption and cholesterol absorption processes may all be influenced by CEL
activity, giving the enzyme compensatory or supplementary roles to other
lipolytic enzymes in the digestive tract. Through its ceramidase capability,
CEL appears to control the assembly and secretion of normal-sized
chylomicrons by enterocytes. Still further functions of CEL have been
postulated in hepatic lipoprotein metabolism and in lipid metabolism in the
vasculature, possibly affecting atherosclerosis (Hui and Howles 2002).
Human CEL from either mammary gland or pancreas is made up of 722 amino
acid residues and is the longest mammalian CEL. The amino acid identity
between human, cow and rat homologues is over 75%. Two intra-chain
disulfide bridges are present in human CEL. C-terminal domain of the enzyme
contains proline-rich tandem repeats whose number differs between species.
This proline-rich region is important for protein stability and provides O-
glycosylation sites. The proline-rich region could also give the enzyme an
open and flexible conformation. The molecular weight of CEL in general
varies between 60 and 100+ kDa and is significantly affected by the degree of
glycosylation (Wang and Hartsuck 1993; Hui and Howles 2002).
The structure of CEL is referred to as the ―left handed oven mitt‖. The enzyme
has a characteristic α/β hydrolase fold motif and contains a lid covering the
active site (Ser194, Asp320, and His435 in the human variant) known as the
active site loop. Primary bile salts bind to the lid, causing a conformational
change that exposes the active site (Blackberg and Hernell 1993; Wang et al.
1997; Moore et al. 2001; Hui and Howles 2002). Bile salts may also bind to a
region away from the active site loop and by doing so protect the lipase from
15
protease attack or allow the lipase to bind to the lipids at the interface
(Blackberg and Hernell 1993; Moore et al. 2001). Arginine residues (e.g.
Arg63 in the human variant) were found to be important for bile salt binding.
N-glycosylation and heparin binding sites are also present on the enzyme
(Moore et al. 2001; Hui and Howles 2002).
2.2.6. General properties
Mammalian digestive lipases produced by pancreas (PL and CEL) have the
following general catalytic properties: temperature optima around 37°C (Bier
1955), e.g. bovine PL (Shahani et al. 1976) and human CEL (Gjellesvik et al.
1992); pH optima in the neutral to moderately alkaline range of 6.5-9.5 and
stabilities between pH 6 and 10 (Bier 1955; Borgstrom and Erlanson 1973;
Vandermeers et al. 1974; Shahani et al. 1976; Momsen and Brockman 1977;
Lombardo et al. 1978; Iizuka et al. 1991).
The molecular weight of PL ranges between 45 and 52.5 kDa (Brockman
1981; De Caro et al. 1981; Winkler and Gubernator 1994; Steiner et al. 2003),
whereas CELs range between 60 and 100 kDa (Wang and Hartsuck 1993). The
isoelectric point of PL is typically between 4.5 and 7.5, i.e. the lipase is
anionic under physiological conditions (e.g. 5.5 for bovine PL – Shahani et al.
1976; 5.2 for porcine PL – Brockman 1981; 4.8 for dromedary PL – Mejdoub
et al. 1994; 6.0–6.2 for canine PL – Steiner and Williams 2002) and (7.4–7.5
for human PL – Iizuka et al. 1991; Miled et al. 2005). CEL is also anionic,
with an isoelectric point between 4 and 6 (e.g. 5.0–6.0 for bovine CEL – van
den Bosch et al. 1973; 5.0 for rat CEL – Erlanson 1975; 4.0 for porcine CEL –
Rudd et al. 1987).
2.2.7. Other mammalian lipases
2.2.7.1. Hormone-sensitive lipase
Another mammalian lipase with broad substrate specificity is an intracellular,
neutral lipase, known most commonly as hormone-sensitive lipase (HSL).
HSL catalyzes the hydrolysis of TGs, diglycerides, monoglycerides at 1 and 3
position, cholesteryl and retinyl esters, lipoidal esters of steroid hormones, as
well as several water-soluble substrates, but has no activity against
16
phospholipids. The importance of this enzyme is revealed in its roles as a
major provider of fatty acids (stored as TGs and other lipids in adipose and
muscle tissues) for energy generation, and as a significant provider of fatty
acids for steroidogenesis, spermatogenesis and possibly other vital cellular
processes. The enzyme is also expressed in macrophages and pancreatic islets.
Although HSL activity against TGs and cholesteryl esters is regulated by
reversible phosphorylation, this mechanism has no effect on the lipase‘s
activity against other substrates. Two things that make HSL quite unique are
its amino acid sequence, which is unrelated to any of the other known
mammalian lipases, and its N-terminal, ~320-residue domain, which has no
primary or secondary structural similarity with known proteins. The C-
terminal domain contains the active site triad (Ser, Asp, His), the regulatory
module with phosphorylation sites, and has α/β hydrolase fold structure
similar to that of CEL. HSL has been most extensively studied in humans and
rats. The enzyme exists as a functional dimer (of identical subunits). The
isoform from human adipose tissue is 775 residues long, with a molecular size
of approximately 84 kDa (Kraemer and Shen 2002).
2.2.7.2. Pre-duodenal lipases
Lipases secreted by the pancreas are the dominant, but not exclusive, enzymes
involved in lipid digestion in animals. Pre-duodenal (lingual, pharyngeal and
gastric) lipases are especially important in situations in which the pancreatic
secretions are or have become minimal, for instance in premature infants or in
the advanced stages of a pancreatic disease (Lopez-Amaya and Marangoni
2000b). In addition to gastric lipases from several sources such as human,
leporine and canine (references in Carriere et al. 1994), lingual lipases of
caprine, bovine, ovine, murine and human origin have also been purified (Lai
et al. 1998 and references therein). O‘Connor and colleagues have extensively
studied caprine lingual lipases (e.g. Lai et al. 1998; Lai and O'Connor 1999;
O'Connor and Manuel 2000).
Unlike PL and CEL, pre-duodenal lipases are stable and highly active in acidic
conditions. The majority of pre-duodenal lipases are glycoproteins and have
molecular weights of 43-51 kDa. The amino acid sequences of gastric and
17
lingual lipases are unlike those of other mammalian, plant or microbial lipases
(Lai et al. 1998; Lopez-Amaya and Marangoni 2000b).
2.2.7.3. Phospholipases
Phospholipases are a broad class of enzymes, catalyzing the hydrolysis of
mainly water-insoluble substrates and involved in digestive, regulatory and
signal transduction pathways, among others. Their nomenclature
(phospholipase A1, A2, B, C and D) is based on which phospholipid ester bond
they attack. The majority of these enzymes require Ca2+
for (maximal)
catalysis, have many disulfide bonds, and are relatively small with molecular
weights from 13 to 30 kDa, but some are above 80 kDa (Lopez-Amaya and
Marangoni 2000a). The amino acid sequences of most phospholipases are
totally different from those of triglyceride lipases (Svendsen 1994). Like lipase
activity, phospholipase activity has been associated with the deterioration of
seafood post harvest (Lopez-Amaya and Marangoni 2000a).
2.2.8. Lipase inhibition
Lipases (like serine proteases) are generally inhibited by certain
organophosphorous compounds (e.g. diethyl p-nitrophenyl phosphate) through
irreversible phosphorylation of active site serine. High concentrations of both
cationic surfactants (e.g. quaternary ammonium salts) and anionic surfactants
(e.g. SDS) affect the quality of the interface and are inhibitory to lipolytic
activity. Other general inhibitors of lipases can include metal ions like Fe2+/3+
,
iodine, boronic acids, sulfhydryl reagents and mercury derivatives. Calcium
and zinc ions tend to activate lipases or produce no effect (Patkar and
Bjorkling 1994; Anthonsen et al. 1995; Gargouri et al. 1997). The effect of
bile salts on the enzyme activity depends on the identity of the enzyme and
presence of cofactors, in addition to the type of bile salts (e.g. colipase restores
bile salt inhibition of PL, and CEL is activated by primary, but not secondary
bile salts).
The use of gastric and pancreatic lipase inhibitors is an important tool in the
control of human obesity (Mukherjee 2003). A common drug that acts by
blocking pancreatic lipase action is tetrahydrolipstatin (Orlistat, Xenical),
18
which was shown to render a third of the consumed fat undigested (Halford
2006). Besides the synthetic chemicals, potential natural inhibitors of
pancreatic lipase have been discovered. Protamine, a very basic polypeptide,
was shown to interfere with lipid absorption by inhibiting the catalytic
functions of PL and CEL (Tsujita et al. 1996). Several proteins, including
lipoxygenase-1, capable of inhibiting PL and some microbial lipases, were
isolated from soybeans (Widmer 1977; Gargouri et al. 1984; Satouchi et al.
1998; Satouchi et al. 2002). The inhibitory capacity of soybean proteins is
believed to be due to the interactions with lipids and ability to modify the
quality of the substrate emulsion. Other prospective natural inhibitors are also
emerging, such as extracts from medicinal plants (Sharma et al. 2005),
polyphenolic compounds, e. g. resveratrol from red wine (Armand 2007) and
green tea catechins (Koo and Noh 2007).
2.2.9. Purification methods
The topic of isolation and purification of lipases has been well reviewed by
Taipa et al. (1992), Aires-Barros et al. (1994) and Palekar et al. (2000). In
summary, non-specific purification techniques involving a precipitation step,
most commonly with ammonium sulphate, acetone or ethanol, and
chromatographic approaches such as gel filtration and ion-exchange
chromatography have been used routinely for purification of animal and
microbial lipases. Adsorption (hydroxyl apatite) chromatography and
hydrophobic interaction chromatography were also used during the
purification sequence of some mammalian lipases (Hyun et al. 1969;
Brockman 1981). Affinity chromatography techniques have been developed in
the last 25-30 years, resulting in faster purification. Examples of ligands used
in the selective separation of CELs include cholate and specifically raised
antibodies (Wang 1980; Abouakil et al. 1988). Mammalian lipases generally
required fewer steps than microbial lipases, because plasma or tissue extracts
tended to contain lower concentrations of protein than fermentation media
(Taipa et al. 1992).
19
2.2.10. Assay methods
Excellent overviews of in vitro lipase and esterase assays are provided by
Gupta et al. (2003b) and Gilham and Lehner (2005). Traditionally, lipase
activity has been measured titrimetrically either directly by pH-stat or by a pH
indicator. The substrates used most often are triolein (or olive oil as a cheaper
alternative) and tributyrin. This is a most reliable and commonly used
procedure. Some other techniques that measure released fatty acids are
spectrophotometry with TGs as substrate (Kwon and Rhee 1986), and TLC,
GC or HPLC, also using TGs as substrate. Other spectrophotometric
techniques measure the release of β-naphthol from its fatty acid conjugates and
para-nitrophenol from its esters. While convenient, the last two methods have
the disadvantages of spontaneous substrate hydrolysis (especially at extreme
pH) and lack of absorbance of para-nitrophenol at acidic pH. Highly sensitive
fluorescence assays, like the one measuring the levels of 4-
methylumbelliferone released from its fatty acid esters (references in Gupta et
al. 2003b), and a turbidimetric method based on the precipitation of fatty acids
with calcium or copper (von Tigerstrom and Stelmaschuk 1989), are also
available. Fluorescence assays, however, require very expensive substrates. A
lipase assay that measures glycerol released from triolein was described by
Young et al. (1988). Even though the method can be very sensitive, detecting
picomole quantities of glycerol, it is not widely applicable since lipases in
general seldom cleave all three fatty acyl chains from a TG within a short time
frame.
Although there is a good choice of lipase assay methods, the search continues
for simpler, faster, cheaper and more specific techniques.
2.2.11. Applications of lipases
More than a decade ago, lipases were touted as having a huge potential for
applications in a multitude of industrial fields (Vulfson 1994). Some progress
has been made and today many of these applications have been made, with
many more promising uses of lipases still to be realized. The following is a
brief overview of current and potential lipase applications, with more detailed
descriptions to be found in the cited reviews and articles.
20
Lipases are very versatile catalysts owing to their various specificities. This
makes them potentially valuable in many applications. Novozyme (Bagsvaerd,
Denmark) markets a range of microbial lipases for various industrial purposes.
Lipases have found extensive applications in the dairy industry for the
hydrolysis of milk fat, contributing to flavour enhancement in cheeses, creams
and other milk products and accelerating cheese ripening. Many other products
can also be given specific dairy flavours by the addition of lipase-hydrolysed
milk fat, using lipases from pancreatic and pre-gastric glands of ovine, bovine,
caprine and porcine origin (Vulfson 1994; Lai et al. 1998). A whole range of
microbial lipases has allowed the production of dairy products suitable for
vegetarians (Vulfson 1994). Besides uses in dairy products, microbial lipases
can be used to ferment many other foods, improving flavour and digestibility.
Some examples are fermented sausages, tempeh (fermented soybeans) and
pickled vegetables (Pandey et al. 1999).
Commercially, the most important field of application for lipases (especially
those of microbial origin) is in household detergents, washing powders and
general cleaners. Lipases in detergents reduce environmental chemical load,
can save power, and remove lipid stains more efficiently than conventional
cleaning chemicals. To be effective in detergents, lipases need to be
alkalophilic and must not be inhibited by various components of the product
formulation (Vulfson 1994; Pandey et al. 1999). Egmond (2002) reviewed
some examples of microbial lipases in detergents.
Lipases can also be used in the oleochemical industry to great advantage for
energy saving and to minimize thermal degradation during processes such as
hydrolysis, glycerolysis and alcoholysis. Lipase-based bioreactors have
become widespread (Vulfson 1994).
Another very important area of lipase application is in the synthesis of
structured lipids (TGs). Lipases can be naturally selected or bioengineered to
provide the necessary substrate-, fatty acid-, regio- and stereospecificity to
selectively enhance TGs with omega-3 fatty acids and conjugated linoleic acid
21
(CLA), as well as to produce many other kinds of desired glycerolipids and
fatty acid concentrates. Higher value fats, such as cocoa butter substitutes, can
be produced from cheaper oils such as palm (Gupta et al. 2003a). Reports of
lipid manipulations using lipase-catalyzed ester bond cleavage, direct
esterification and transesterification are numerous (e.g. Breivik et al. 1997;
Robles Medina et al. 1999; Haraldsson et al. 2000; Halldorsson et al. 2001;
Torres et al. 2001; Akoh et al. 2002; Rocha-Uribe and Hernandez 2004;
Kojima et al. 2006).
Organic synthesis using lipases has been trialled and applied in both laboratory
and commercial settings. Various surfactants/emulsifiers (e.g. diglycerides,
monoglycerides and sorbitan esters) can be more economically produced using
lipases than via traditional chemical pathways, as can wax esters and other
specialty esters used in personal-care products, cosmetics and perfumes. As an
example, Croda Universal Ltd. (Hull, United Kingdom) uses enzymatic
processes to manufacture wax esters. Enantiomerically pure pharmaceuticals,
agrochemicals (e.g. pesticides) and polymers are also produced through bio-
organic synthesis using lipases (Kotting and Eibl 1994; Vulfson 1994; Pandey
et al. 1999; Patel 2002).
Biodiesel (fatty acid alkyl esters) can be conveniently produced via
transesterification reaction between a TG and a short chain alcohol using
lipases as biocatalysts, instead of more traditional alkali or acid catalysts.
Lipases offer several advantages, like the possibility of multiple use of the
same enzyme and ease of product recovery, both due to the immobilization
approach (Marchetti et al. 2007). Immobilization of whole lipase-producing
cells in a packed-bed reactor (Hama et al. 2007), lipase catalyzed methanolysis
(Nie et al. 2006) and use of immobilized lipase in a solvent free system (Shah
and Gupta 2007) are some of the many examples of biodiesel production using
lipases.
As discussed in section 2.2.1, several plant lipases have unusual properties
compared with mammalian and microbial lipases. Consequently, these plant
lipases are used as biocatalysts for synthesis and transformation of novel lipids
(Mukherjee and Hills 1994; Mukherjee 2002).
22
Lipases have the potential to be used in many other applications, such as
environmental management, as biosensors, in the leather industry (Pandey et
al. 1999; Haas et al. 2002), in the pulp and paper industry (Sharma et al.
2001), in tea processing (Ramarethinam et al. 2002) and cake manufacture
(Guys and Sahi 2006).
2.2.11.1. Biocatalysis in organic solvents
Although lipases usually operate in an aqueous environment in vivo, they are
also able to catalyze reactions in organic solvents. This ability is used in
particular for interesterification of industrial fats and oils and synthesis of
enantiomerically pure lipids (Torres and Castro 2004). Aquatic lipases are not
yet in large-scale commercial use. However, their different specificities and
chemical properties suggest there may be potential for their application in
organic solvents.
The field of enzymatic catalysis in ‗non-conventional media‘ (media which are
not purely aqueous) has been developing progressively in the last three
decades (Castro and Knubovets 2003). Organic solvents represent one of the
non-conventional media (besides reversed micelles, supercritical fluids and
ionic liquids) and enzymes are being used increasingly in these systems.
Catalysis in almost exclusively hydrophobic solvent media is possible due to
several important effects on enzymes, which include greater structural rigidity
(high kinetic barriers prevent the native-like conformation from unfolding) and
‗pH memory‘. These effects originate from the presence of a water monolayer
around the enzyme that shields the enzyme from direct interaction with the
bulk solvent. The enzyme remains catalytically active as long as this essential
layer of water is present. Hydrophobic solvents that do not strip the water
monolayer and lyophilization of enzymes prior to addition to organic solvent
are important in this regard (Zaks and Klibanov 1988; Klibanov 1989).
Despite several disadvantages of using organic solvents – high price, solvent
toxicity and flammability, higher equipment costs, mass transfer limitations in
viscous solvents and reduced enzyme activity relative to purely aqueous
systems (Castro and Knubovets 2003; Bezbradica et al. 2007) – there are
23
numerous benefits available to the organic chemist. These advantages include
the improved solubility of lipophilic substrates in organic solvents, reduced or
absent microbial contamination, efficient recovery and reusability of the
suspended enzyme, increased enzyme thermostability, suppression of
undesirable water-dependent side reactions, a kinetic shift towards synthetic
reactions instead of hydrolysis, and altered substrate specificity,
regiospecificity and enantioselectivity (Gupta 1992; Wescott and Klibanov
1994; Torres and Castro 2004). Lipases, in particular, favour the reactions of
esterification, transesterification, alcoholysis and acidolysis, over hydrolysis,
in homogenous organic solvent systems (Pencreac'h and Baratti 1996; Lima et
al. 2004). These reactions are important for preparative purposes in organic
synthesis, such as the synthesis of pharmaceutical precursors and drugs, pure
enantiomers, single isomers and biopolymers (Torres and Castro 2004). The
synthesis of biosurfactants (e.g. monoglycerides), as well as the
interesterification of oils and fats, has been achieved using lipases in organic
solvents (Gupta 1992). Lipase synthesis of structured lipids containing CLA
and alkyl esters of fatty acids, in hydrophobic media, has also been reported
(Hazarika et al. 2002; Rocha-Uribe and Hernandez 2004). Various
modifications have been applied to lipases in order to improve their solubility,
activity and stability in organic solvents: attaching soluble modifiers like
aldehydes, polyethylene glycols and imidoesters, adsorption onto support
particles, gel entrapment, immobilization, and protein engineering (Gupta
1992; Salleh et al. 2002; Hult and Berglund 2003; Mateo et al. 2007). Further
developments in the area of lipase modification and media engineering will
lead to improved biocatalysis in organic solvents and production of novel
structured lipids, emulsifiers and intermediates in organic synthesis.
2.3. Lipases from aquatic species
2.3.1. Introduction
Fish tissue by-products are a potentially rich source of numerous
biochemicals, including enzymes such as lipases. These by-products can
include the head, frame, viscera and sometimes the skin and scales. The
digestive organs and glands are the most important by-products in terms of the
24
quantity of enzymes. Fish digestive organs such as stomach, pyloric cecum,
pancreas and intestines are relatively unexplored sources of lipolytic enzymes
that could have unique characteristics. The demand for lipases with novel
properties is ongoing. Considering the evolutionary pathways, unusual diets
and habitats of fish, piscine lipases may present certain qualities that
complement those of the lipases from conventional mammalian and microbial
sources.
The underutilized digestive organs from the large volume landings of many
fish species worldwide are available as an exploitable and relatively cheap
source of lipases. As an example, Alaska pollock is one of the world‘s largest
fisheries and its viscera are a potential source of useful lipases (FAO 2007;
McGinnis and Wood 2007), especially as the diet of the fish contains a high
percentage of phospholipids and wax esters (Lee et al. 1970; Saito et al. 2002).
Another possible large source of novel marine lipases are Antarctic krill. Wax
esters can account for more than half of total storage lipids in this organism
(Ju and Harvey 2004) suggesting the presence of possibly unique wax-ester
specific lipases. Before these lipases can be extracted and products developed,
a full understanding of their characteristics needs to be gained. To this end,
more fundamental research into fish digestive lipases is required, especially
from the commercially important species such as wild-harvested pollock,
herring, sardine, tuna, and anchovy as well as aquacultured tilapia, salmon and
various species of carp (FAO 2007).
The prospect of extracting enzymes from fish wastes is attractive from an
economic viewpoint. Currently, most fish by-products are turned into low
value fish meal and crude oil fractions. Economic returns could be improved if
specialty biochemicals were extracted and marketed on a commercial scale.
Fish digestive organs represent an abundant raw material as most fish are
gilled and gutted before sale. Furthermore, as wild fish stocks are a limited
natural resource, it is of general importance to use the available raw material
as wisely as possible.
25
In order to determine whether fish can be a possible source of lipase(s) for
industrial applications, Nayak et al. (2003) prepared crude enzyme extracts by
centrifuging tissue homogenates from rohu (Labeo rohita), oil sardine
(Sardinella longiceps), mullet (Liza subviridis) and Indian mackerel
(Rastrelliger kanagurta). With the titrimetric method using tributyrin as
substrate, rohu had the highest lipolytic activity in all of the tissues (intestine,
stomach/pyloric ceca, liver/hepatopancreas, red muscle) compared with the
other fish. In general, the highest lipase activities were observed in intestine
and stomach/ceca. Lipolytic activity in the muscle was the lowest among
tissues for all fish, except mackerel, in which it was the highest. Rohu
intestinal and hepatopancreatic extracts were also able to hydrolyze fish oil.
The authors concluded that fish digestive organs might be a good source for
industrial lipase production.
Some fish that inhabit cold and temperate environments have enzymes with
relatively high catalytic efficiencies, as a result of adaptation to their
environments. Thus, marine species could be a novel source of unique, high-
activity lipases that offer the benefits of lower thermal requirements for
catalysis, and deactivation by mild heat treatment. The adaptation strategies of
enzymes produced by psychrophilic organisms in general are beginning to be
understood (Georlette et al. 2004). These enzymes show highly flexible
catalytic regions. Marine lipases operating at cold/temperate environments
could show similar features.
The reactivity of marine digestive lipases towards fatty acid classes differs
from that of digestive lipases from other sources, most notably mammals, but
also from those of plants and micro-organisms. Whereas mammalian digestive
lipases such as CEL most actively cleave fatty acids with ≤ 20 carbons and a
low degree of unsaturation, the preference in fish is towards long-chain
polyunsaturated fatty acids (PUFA). Gjellesvik (1991) demonstrated this with
CELs from Atlantic cod and of human origin. The high reactivity towards
long-chain polyenoic fatty acids could be very useful in many applications
(see section 2.3.4).
26
The lipolytic systems in fish differ from those in mammals. Examples of
differences include the potentially exclusive or predominant role of CEL-like
enzyme in lipid digestion compared with PL-like enzyme (Lopez-Amaya and
Marangoni 2000b), a likely absence of HL-like enzyme (Lindberg and
Olivecrona 1995) and a translated last exon of LPL (Arnault et al. 1996).
2.3.1.1. Fish lipases
Chesley (1934) presented one of the earlier investigations regarding the
presence and quantity of lipases in digestive tissues of several marine species.
Active fishes, mackerel and scup, had markedly higher lipase activities than
more sluggish species, with the upper intestine, pyloric ceca and pancreas
containing most of the enzyme.
Later research in fish digestive lipolysis was inconclusive as to which types of
lipases fish have. Some workers reported PL-like enzyme and colipase in
rainbow trout (Leger et al. 1970; Leger et al. 1977; Leger et al. 1979). Patton
and his group did not detect PL-like activity in the leopard shark pancreas, but
instead partially characterized CEL-like enzyme from this species (Patton
1975; Patton et al. 1977). Still other researchers, such as Mukundan et al.
(1985), purified a lipase from the hepatopancreas of oil sardine but did not test
the effects of bile salts and colipase on the lipolytic activity, and hence this
lipase has remained unclassified. Some other studies on fish digestive lipases
were performed using either crude extracts (e.g. Borlongan 1990; Das and
Tripathi 1991) or were directed at the products of lipid digestion (e.g. Patton et
al. 1975) and thus gave indirect information about the enzymes involved.
Some new insights into the nature of fish digestive lipases have been provided
in the last 15 years with the purification of first non-mammalian CEL from the
pyloric ceca/pancreas of Atlantic cod by Gjellesvik et al. (1992). This was
followed by another breakthrough study in 1994 by Gjellesvik et al., who
reported the cDNA sequence of pancreatic CEL from Atlantic salmon, the first
such sequence of a non-mammalian origin. Since then, another piscine CEL
has been purified to near homogeneity from the hepatopancreas of red sea
bream (Iijima et al. 1998). Many other investigations into the occurrence and
27
specificities of digestive lipases, as well as the developmental aspects of
lipases in larval and juvenile fish, have also been carried out in the past decade
(e.g. Bolasina et al. 2006). While useful, these investigations only concerned
crude enzyme preparations or analyses of digesta.
In addition to digestive lipases, tissue lipases have also been studied in fish,
but to a lesser extent. Lipases in red muscle, adipose tissue and liver of
different fish species have been studied to better understand the mobilization
of depot fat in fish and to compare it to the same process in mammals (see
section 2.3.3).
2.3.1.2. Lipases from other marine animals
There are far fewer reports on lipase systems in other marine animals.
Digestive lipases are mentioned in a few studies conducted on marine
crustaceans, e.g. American lobster (Homarus americanus) larvae (Biesiot and
Capuzzo 1990). Such studies generally examined larval/young animals,
sometimes for ontogenic purposes.
A few bivalves and a cephalopod have been examined. A crude lipase
preparation from the crystalline styles of surf clam (Spisula solidissima)
hydrolyzed triolein most efficiently at pH 8 and 20°C. The clam lipase had
very low specific activity against triolein compared with porcine pancreatic
lipase, and was unable to split long-chain PUFA methyl esters (Patton and
Quinn 1973). A preliminary investigation into the digestive enzymes of the
green mussel (Perna viridis) revealed very low lipase activity in this species
(Teo and Sabapathy 1990). Itabashi and Ota (1994) found that delipidated
powders from scallop hepatopancreas can hydrolyze the sn-1 and sn-3 ester
bonds of TGs. Lipid breakdown in shellfish during cold storage has been
associated with the hydrolytic action of lipases (Kaneniwa et al. 2004).
A lipase was partially purified from the hepatopancreas of neon flying squid
(Ommastrephes bartramii) by anion-exchange chromatography and gel
filtration of the delipidated tissue powder. The apparent molecular weight of
the enzyme, 33 kDa, is rather low compared with either PL or CEL from
28
mammalian species. The squid lipase displayed optimum activity at pH 7.0,
and was stable between pH 6.0 and 9.0 for 6 h at 25°C, while thermal
optimum and stability were achieved at 25°C and up to 37°C (for 10 min),
respectively. The cephalopod lipase did not show particular regioselectivity
and, akin to land-based mammalian lipases, hydrolysed monounsaturated and
saturated chain TGs most readily. A colipase was isolated in the same study
and when added to the hydrolytic system it increased the initial activity of the
lipase approximately ten times (Sukarno et al. 1996).
2.3.2. Fish digestive lipases
2.3.2.1. Rainbow trout (Oncorhynchus mykiss)
Very few PL-like enzymes have been reported in fish. One such enzyme was
isolated from the inter-cecal pancreatic tissue of rainbow trout (Salmo
gairdneri, now called O. mykiss) by Leger et al. (1977). A few years earlier,
Leger‘s group reported on the in vitro effects of pH and temperature on the
lipolytic activity of a crude preparation from the diffuse pancreas of rainbow
trout (Leger et al. 1970). Leger and Bauchart (1972) presented further studies
on the pancreatic lipase from this salmonid that showed selectivity towards the
two exterior ester bonds on the glycerol backbone of TGs and preference for
the detachment of oleic acid from the trialcohol backbone, irrespective of
position. In the same year, Leger (1972) published a method for the partial
purification of this enzyme that encompassed defatting, ammonium sulphate
precipitation and gel filtration steps. This method was used by Leger et al.
(1977) in their isolation of the enzyme. The trout lipase hydrolyzed tributyrin
in the absence of Ca2+
, but required mM concentrations of the ion for the
lipolysis of triolein. The enzyme was shown to have higher affinity towards
tributyrin than porcine PL. The addition of mixed bile salt conjugates
increased the activity against triolein of the fish enzyme several-fold, and a
linear dependence was observed between the lipase activity and the amount of
bile salts in the range of 25-250 mM. Considering the lipase activity relative to
the bile salt concentration, the authors suggested the existence of a colipase,
which was detected in 1979 (Leger et al. 1979). These researchers employed
techniques such as anion-exchange chromatography and ultrafiltration in
29
addition to gel filtration. In the 1979 study, the molecular weights of rainbow
trout PL-like enzyme and colipase were estimated to be 57 and 11 kDa
respectively.
The conclusions of Leger and co-workers that the lipolytic system in trout
pancreas is of the pancreatic lipase-colipase type have been debated by other
researchers in the field such as Gjellesvik et al. (1992). The latter group
pointed to the very low specific activity of the trout enzyme versus the porcine
homologue characterized by Verger et al. (1969) and questioned the purity and
identity of the trout lipase. The trout lipase activity data, taking in account bile
salt concentration, and the estimated molecular weight of 57 kDa, could also
suggest the presence of a fish CEL-like enzyme. Thus, in the absence of a
more highly purified lipase extract from trout pancreas, it can be argued that
this fish has a PL-like enzyme, or a CEL-like enzyme, or both.
Another study on the digestive lipolytic activity in rainbow trout was carried
out by Tocher and Sargent (1984) using a desalted infranatant from a
suspension of pyloric ceca. These workers were interested in the variations of
TG, wax ester and sterol ester hydrolase activities in the ceca with different
diets. It appears that CEL-like enzyme may be present in the trout ceca, owing
to the observation that primary bile salts were a prerequisite for any lipolytic
activity. The authors of the study suggested the presence of two different
lipolytic proteins in the salmonid‘s ceca, one predominantly hydrolyzing TGs
and the other predominantly attacking wax and sterol esters. The apparently
different observations concerning the identity of the lipase seen between this
study and the ones of Leger‘s group may be related to the source of the
enzyme – Tocher and Sargent (1984) looked at the lipolytic activity in the
pyloric ceca, whereas Leger and co-workers (1977) investigated this in the
diffuse pancreas of rainbow trout. Nevertheless, no reliable conclusion can be
reached about the full nature of the lipase from the very simple enzyme
preparation of Tocher and Sargent (1984).
30
2.3.2.2. Oil sardine (Sardinella longiceps)
Mukundan et al. (1985) described the purification of a lipase from the
hepatopancreas of oil sardine using anion-exchange chromatography and gel
filtration. The glycosylated enzyme (6.1% carbohydrate) showed a molecular
weight of 54-57 kDa (by gel filtration), was marginally activated by ~1 mM
Ca2+
, was completely inhibited by ~1 mM Fˉ and hydrolyzed tributyrin most
efficiently amongst C2 to C18 monoacid TGs. The sardine lipase exhibited
optimum activity against tributyrin at pH 8-8.5 and 37°C. The enzyme was
stable up to 45°C for 15 min and in the pH range 5-9.5 after 1 h incubation
period. The regioselectivity of the enzyme was towards the exterior, α-ester
bonds of TGs.
2.3.2.3. Colipase (various sp.)
Colipase is a critical cofactor for the reversal of PL inhibition by bile salts and
has been mainly studied in mammals (e.g. Erlanson et al. 1973). In addition to
the partial characterization of colipase from rainbow trout intercecal pancreatic
tissue by Leger et al. (1979), evidence for the presence of the cofactor was
reported in several other fish, including hagfish, ratfish, rayfish and Greenland
shark (Sternby et al. 1983). Dogfish (Squalus acanthius) pancreatic colipase
was purified by Sternby et al. (1984) using an ion-exchange chromatography
step followed by gel filtration, and is the only piscine colipase fully purified to
date. The amino acid composition of this colipase was at least 57% identical to
those of human and galline cofactors. The dogfish colipase consisted of 86-90
residues and its molecular weight was calculated at ~9.2 kDa. The specific
activity with the human pancreatic lipase was 4-5 times lower than the pure
human complex and this was explained as potentially relating to the
interspecies differences in the cofactor, which includes the very high pI of the
dogfish colipase (10.2 vs. 5.8 for the human colipase).
2.3.2.4. Spiny dogfish (Squalus acanthius)
The presence of colipase in dogfish pancreas ought to indicate that PL-like
enzyme is involved in lipid digestion in this elasmobranch. A lipase was
isolated from the pancreas of dogfish by Rasco and Hultin (1988) using
preparative isoelectric focusing, with pre-treatments including tissue defatting
31
and drying, ammonium sulphate precipitation, desalting and ultrafiltration.
The partially purified enzyme had a pH optimum of 8.5, temperature optimum
of 35°C and seemed quite stable at 45°C up to 1 h. Above 45°C, however, at
least half of the activity was lost after 1 h incubation. The enzyme was able to
hydrolyse a wax ester more efficiently than a commercial porcine PL, but did
not show high rates of cleavage of low molecular weight fatty acid ester bonds
present in dairy cream TGs. The activity of the enzyme was increased several-
fold by addition of sodium taurocholate up to about 20 mM, but not by
deoxycholate. From the effect of bile salt concentration on the activity of the
enzyme, Rasco and Hultin hinted that the lipase may be of a PL-type and that
colipase may have been co-isolated with it. Further purification and
characterization are needed to establish the identity of the dogfish lipase.
2.3.2.5. Atlantic cod (Gadus morhua)
After the work of Overnell (1973) in which no lipolytic activity was detected
in the pyloric ceca and associated mesentery of Atlantic cod, Lie et al. carried
out further investigations on the digestive lipolytic enzymes in this gadiform
(Lie and Lambertsen 1985; Lie et al. 1987; Lie and Lambertsen 1991). By
withdrawing the lipid digesta and the digestive juice from various segments of
the digestive tract, Lie‘s group discovered that lipolysis in Atlantic cod was
stimulated by bile salt and that the lipolytic activity was preferential towards
esters of longer, PUFAs. Gjellesvik et al. (1989) partially purified and
characterized a TG lipase from the pyloric ceca of G. morhua and also found
that it has an absolute requirement for bile salts for the hydrolysis of olive oil.
Furthermore, Gjellesvik et al. (1989) were unable to demonstrate the presence
of colipase in the cecal extracts. The findings of the groups of Lie and
Gjellesvik indicated that mammalian CEL-like enzyme may be secreted by the
digestive organs/glands of Atlantic cod. This hypothesis was confirmed with
the purification of pancreatic/pyloric ceca CEL from cod in 1992 by Gjellesvik
et al. Cholate-Sepharose affinity chromatography and gel filtration were used
to isolate the enzyme in a pure form. The enzyme‘s molecular weight was
estimated at 60 kDa by SDS-PAGE. Bile salts were an absolute requirement
for the hydrolysis of insoluble fatty acid esters (TGs – triolein and those in
olive oil, and cholesteryl oleate). CEL from cod appears to have a pH optimum
32
of 6.5-7.5 and a temperature optimum in the range of 25 to 35°C, and the
enzyme is rapidly inactivated above 40°C. The amino acid composition of the
cod enzyme was compared to several mammalian CELs (further discussed in
section 2.3.3). Cod CEL had highest rate of hydrolysis with diglyceride as
substrate, 10 times faster than against TG, and esters of long chain PUFAs
(except DHA) were most efficiently hydrolysed (Gjellesvik 1991; Gjellesvik
et al. 1992). Contrary to the general expectation of enzymes from cold-adapted
poikilotherms, cod CEL did not show significantly higher activity at the
temperature of the fish‘s habitat (0 to 8°C) than the human pancreatic
counterpart, and the catalytic efficiency of the cod enzyme at 25°C was
considerably lower than that of the human enzyme. The cod lipase may have
rather flexible structure, shown by lower cysteine content and lower
hydrophobicity compared with human CEL, although it is not reflected in the
catalytic efficiency of this enzyme. Gjellesvik et al. (1992) did not detect any
colipase-dependent PL-like activity in the cod tissue isolates and postulated
that CEL could be the only pancreatic/cecal enzyme involved in lipid digestion
in this species.
2.3.2.6. Atlantic salmon (Salmo salar)
Gjellesvik was also part of a research team who published the cDNA sequence
of pancreatic CEL from Atlantic salmon. This enzyme consists of 527 amino
acids, does not contain the C-terminal proline-rich repeats, and has
approximately 58% primary structure identity with the mammalian analogues.
The authors used the tertiary structure of acetylcholine esterase from Torpedo
californica as a template for the construction of a computer-assisted model of
the 3-D organization of the salmon enzyme and located putative bile salt-
binding and lipid-binding sites on it. Using the location of the ‗flap‘ (residues
between Cys64 and Cys80) in fungal lipases from Geotrichum candidum and
Candida rugosa as a guide, the authors concluded that their model of salmon
CEL did not contain an active site ‗lid‘ domain (Gjellesvik et al. 1994).
However, the present knowledge of the location of active site loop in
mammalian CELs, such as those of human (Moore et al. 2001; Hui and
Howles 2002) and bovine (Wang et al. 1997) origins, points to residues in the
range 115-126. This region is highly conserved between salmon and
33
mammalian CELs and may well contain the active site loop in the fish
enzyme.
2.3.2.7. Yellowfin tuna (Thunnus albacares)
In 1996, Maurin and Gal reported on the characterization of a partially purified
60 kDa enzyme from the pyloric ceca of yellowfin tuna. Ethanol fractionation
and immobilized metal-ion affinity chromatography were used to prepare the
enzyme sample. Highest esterolytic activities were obtained at pH 8.0. The
enzyme was thermally unstable, losing more than 30% of its original activity
following incubation at 30°C for 30 min. Synthetic ester substrates with acyl
chains exceeding four carbons were not hydrolyzed by the tuna enzyme. Based
on the characteristics of the enzyme, especially its acyl chain length
specificity, the authors concluded that the isolated tuna enzyme was a
hydrophobic esterase and not a lipase (Maurin and Gal 1996).
2.3.2.8. Red sea bream (Pagrus major)
A CEL from the hepatopancreas of red sea bream was purified to near
homogeneity and partially characterized by Iijima et al. (1998). The enzyme
was extracted by a combination of anion-exchange, hydrophobic interaction
and molecular sieve chromatography. Although only 2.3% of the total activity
in the crude extract was recovered after the gel filtration step, this lipase
fraction was purified about 340-fold and had specific activity of 75 U/mg
protein with para-nitrophenyl myristate as the substrate. Preliminary
investigations by the authors showed that the lipase binds tightly to a cholate-
Sepharose column, and this strong interaction was confirmed through the
finding that the enzyme had an absolute requirement for primary (7α-
hydroxylated) bile salts for activity against esters of long chain fatty acids
(para-nitrophenyl myristate and triolein). The estimated molecular weight (64
kDa) and the pH optimum (7.0-9.0) of red sea bream CEL are both similar to
the cod CEL (see above in this section). The lipolytic activity of the bream
enzyme is Ca2+
-independent. As was the case with the cod homologue, the
bream lipase had highest reactivity towards esters of 20:4n-6 and 20:5n-3 fatty
acids, and 22:6n-3 was also cleaved from the alcohol backbone. Thus, red sea
34
bream lipase hydrolyzes substrates that are poorly utilized by the PL family of
enzymes.
2.3.2.9. Nile tilapia (Oreochromis niloticus)
Taniguchi et al. (2001) reported on the purification of intestinal lipase from
tilapia (Tilapia nilotica, now called O. niloticus). The lipase was purified
using ion-exchange, chromatofocusing and gel filtration chromatography
techniques. The temperature optimum was 35°C and it was stable below 40°C.
The pH stability fell within the relatively narrow range of 6.5-8.5, and the
highest activity was at pH 7.5, typical of corresponding mammalian lipases
and those from other fish species. The inclusion of bile salts caused an
approximate 9-fold increase in the enzyme activity. The tilapia lipase‘s
molecular weight of 46 kDa, pI of 4.9 and efficient hydrolysis of soybean and
coconut oils are characteristics similar to mammalian PL.
2.3.2.10. Grey mullet (Mugil cephalus)
A lipase was partially purified from the viscera (pyloric ceca, intestines and
associated mesenteries) of grey mullet by ammonium sulfate fractionation,
ultrafiltration and cholate-Sepharose affinity chromatography (Aryee et al.
2007). Optimum activity against para-nitrophenyl palmitate was displayed at
pH 8, akin to most other fish and mammalian lipases. The temperature
optimum against the same substrate was 50°C, which was also the stability
limit. Grey mullet lipase is completely stable in several water-immiscible
organic solvents, suggesting that it may have potential applications for
synthesis reactions in organic media. The classification of this lipase awaits
further purification, determination of molecular weight, and amino acid
composition/sequence analyses.
2.3.2.11. Lipase activity in unpurified fish extracts
Whereas the majority of the studies described in sections 2.3.2.1–10 were
conducted on at least partially purified lipase extracts, other studies of fish
digestive lipolysis used lipid-containing digesta or very crude enzyme
preparations. Some of these studies are reviewed briefly below.
35
2.3.2.11-1. In vivo feeding trials and analysis of gut content
The specificity of digestive lipases in several fish was assessed by Patton et al.
(1975) via two approaches. Northern anchovies (Engraulis mordax) were used
in in vivo feeding experiments and the postprandial contents of their
alimentary canal were examined. The intestinal and pyloric ceca contents of
pink salmon (Oncorhynchus gorbuscha), striped bass (Morone saxatilis), jack
mackerel (Trachurus symmetricus), pacific mackerel (Scomber japonicus) and
northern anchovy were centrifuged and then supernatant was used as the
source of the lipase for in vitro activity assays. The results from both
approaches indicated that: (i) these fish have enzyme(s) that hydrolyze(s) wax
esters more efficiently than mammalian PL; (ii) long-chain PUFAs such as
20:4n-6 and 20:5n-3 were cleaved faster than less unsaturated ones such as
18:2 and 18:3 and (iii) the sn-2 ester bond in TGs was attacked by the lipase(s)
in the sample. From these findings, Patton et al. suggested that an enzyme
other than PL may contribute significantly to the lipid digestion process in the
fish. At the time of this study little was known about CEL, and the alternative
enzyme was referred to as non-specific lipase or carboxylic ester hydrolase.
2.3.2.11-2. Leopard shark (Triakis semifasciata)
The supernatant from a suspension of acetone/diethyl ether delipidated
pancreatic powder in pH 8.5 Tris buffer was used to partially characterize a
lipase from leopard shark. The temperature optimum of the enzyme was 36°C,
but the enzyme was rapidly inactivated at 50°C even in the presence of bile
salts. Ca2+
increased the lipolytic activity of the sample. The authors classed
the lipase as CEL type based on an absolute requirement for trihydroxy bile
salts for activity, as well as the hydrolysis of all three ester bonds of TG
substrates and PL-resistant ester bonds formed by long-chain PUFA and
glycerol. PL-like activity was not detected (Patton et al. 1977).
2.3.2.11-3. Milkfish (Chanos chanos)
The digestive lipase activity along the digestive tract of milkfish, a widely
aquacultured species in Southeast Asia, was evaluated by homogenizing and
then centrifuging the tissue of interest to obtain a clear supernatant as the
crude enzyme extract. Highest lipolytic activities were found in the intestines,
36
pancreas and pyloric ceca, where the temperature and pH optima of the
enzymes were 45-50°C and around 6.5 and 8.5 respectively. The temperature
optimum represents adaptation to the environment, as milkfish can live in
pond and lagoon waters exceeding 40°C (Borlongan 1990). The presence of
maximal activity at two pH values may indicate that this species has two kinds
of digestive lipase.
2.3.2.11-4. Arctic charr (Salvelinus alpinus)
By analyzing the contents of digesta from various gut segments, Olsen et al.
(1998) suggested that non-specific lipolytic activity predominates in the
pyloric ceca and intestinal tract of Arctic charr. Substrates containing PUFAs
and medium-chain saturated fatty acids were well digested and absorbed,
while longer chain saturated and monounsaturated fatty acids tended to remain
in the undigested glycerides.
2.3.2.11-5. Juvenile turbot (Scophthalmus maximus/Psetta maxima)
The posterior digestive tract comprising the hindgut and rectum appears to be
the major site for lipolysis and lipid absorption in juvenile turbot (Koven et al.
1997). Lipid digestion was studied both in vivo (analysis of digesta post
feeding, Koven et al. 1994b) and in vitro (centrifuged digesta as the crude
enzyme source, Koven et al. 1994a). The lipolysis in this flatfish was
positionally non-specific and PUFA were preferentially cleaved and absorbed.
In another study, Izquierdo and Henderson (1998) successfully demonstrated
the use of fluorimetric assays for the analysis of lipase and phospholipase
activities in small quantities of tissue in turbot. Hoehne-Reitan et al. (2001a)
measured an increase in CEL activity with growth in turbot larvae. Hoehne-
Reitan et al. (2001b) established the positive correlation between the level of
CEL in larval turbot and the quantity of the prey, but found that the lipid
content of the prey did not affect the enzyme levels in the early feeding phase.
The synthesis and secretion of neutral lipase and phospholipase in developing
turbot larvae were also influenced by diet (Hoehne-Reitan et al. 2003). Many
other studies have looked at lipid and fatty acid requirements of marine fish
larvae and a few have examined the changes in lipolytic enzymes in
37
developing fish larvae (references in Hoehne-Reitan et al. 2001a; Hoehne-
Reitan et al. 2003).
2.3.2.12. Overview-properties of purified and partially purified fish digestive
lipases
There are few studies where fish lipases have been fully purified. A summary
of known data for fully and partially purified lipases (see sections 2.3.2.1–10
for references) is given in Table 2.1. Fish digestive lipases are either of CEL
or PL types. CEL has broader substrate specificity and can hydrolyze a wide
range of lipid classes. PL is more specific, favouring triglycerides. From the
very limited available data, there are indications that freshwater fish may favor
PL-type lipase as the dominant enzyme, and marine fish CEL-type lipase. It is
likely that these differences are a response to differences in diet, i.e. freshwater
fish are primarily omnivorous (energy derived from lipids and carbohydrates),
whereas marine fish are primarily piscivorous, consuming less carbohydrate,
and a wider variety of lipid classes that are not hydrolysed by PL (e.g. wax
esters).
Table 2.1
Properties of purified or partially purified fish digestive lipases (see text for
references)
Source –
common name
Latin name Fresh-
water or
marine
CEL-type
or PL-type
Mol.
weight
(kDa)
pI pH
optimum
pH
stability
Temp.
optimum
(°C)
Temp.
stability
(°C)
Nile tilapia Oreochromis
niloticus
F PL-like 46 4.9 7.5 6.5-8.5 35 <40
Rainbow trout Oncorhynchus
mykiss
F PL-like 57 8.4-8.7
Atlantic cod Gadus morhua M CEL 60 6.5-7.5 25-35 ≤40
Atlantic
salmon
Salmo salar M CEL <60
Red sea bream Pagrus major M CEL 64 7-9
Yellowfin tuna Thunnus
albacares
M 60 8 <30
Oil sardine Sardinella
longiceps
M 54-57 8-8.5 5-9.5 37 ≤45
Spiny dogfish Squalus
acanthius
M PL-like? 5.4 8.5 35 ≤45
Grey mullet Mugil cephalus M 8 4-10 50 ≤50
38
2.3.3. Fish tissue lipases
2.3.3.1. Red muscle
Red muscle is one of the lipid storage organs in fish and was examined for
lipolytic activity towards both long-chain and short-chain TGs (Bilinski and
Lau 1969 and references therein). It is not yet established whether the same
enzyme breaks down both classes of substrate in this tissue. The finely cut
lateral line muscle of rainbow trout hydrolyzed longer chain TGs (e.g.
tripalmitin) most efficiently at pH 7.3 and required the dispersal of substrate
by phospholipids for lipolysis. Sodium fluoride and protamine inhibited the
lipolytic activity, whereas albumin and epinephrine, typical activators of
hormone sensitive lipase (HSL) in mammals, had little effect on the lipolysis
in the salmonid muscle. It appears that some differences exist between the
neutral, fat-mobilizing lipase in trout red muscle and that in mammalian
adipose tissue (Bilinski and Lau 1969).
A crude enzyme extract of red muscle from rohu (L. rohita) had higher muscle
lipase activity than oil sardine (S. longiceps), mullet (L. subviridis) and Indian
mackerel (R. kanagurta). In the same study, Indian mackerel and mullet had
equivalent lipolytic activities in the liver (Nayak et al. 2003).
2.3.3.2. Adipose tissue
Sheridan and Allen (1984) partially purified a TG lipase from the adipose
tissue (i.e. mesenteric fat) of juvenile rainbow (or steelhead) trout. Heparin-
Sepharose affinity chromatography was used to separate the enzyme of
interest from lipoprotein lipase. The salmonid TG lipase was akin to
mammalian HSL based on the pH optimum of 7.5 and activation by cAMP-
mediated phosphorylation of the fish enzyme. The trout lipase showed the
highest activity at 25°C, as expected from a temperate environment fish, and
was estimated to be a 48 kDa protein (Sheridan and Allen 1984; Michelsen et
al. 1994). HSLs from different animals vary in their molecular weights –
human and rat enzymes are 80-85 kDa in size (Kraemer and Shen 2002 and
references therein), whereas chicken HSL is 42 kDa (Berglund et al. 1980). It
is unclear if hormonal stimulation of adipose tissue lipolysis is present in trout
(Michelsen et al. 1994).
39
2.3.3.3. Liver
Another organ that accumulates lipid reserves in many fish is the liver. A
hepatic TG lipase was partially purified from juvenile rainbow trout using
size-exclusion chromatography (Harmon et al. 1991). Heparin-Sepharose
affinity chromatography was used to remove lipoprotein lipase, which was
present in very low amounts. The hepatic TG lipase had a pH optimum of 7.0,
a temperature optimum of 15°C and a molecular weight of 40-43 kDa. These
are all somewhat lower than the corresponding properties of adipose tissue TG
lipase from the same species (see the previous paragraph). Like the adipose
tissue enzyme, the hepatic cytosolic enzyme was activated by phosphorylation
in the presence of cAMP/ATP-Mg2+
(Harmon et al. 1991). The
phosphorylation state of the hepatic TG lipase was shown to be altered by
insulin and glucagon (Harmon et al. 1993). In conclusion, the biochemical
properties of the trout hepatic enzyme are similar to those of mammalian HSL
and unlike those of mammalian hepatic lipase (HL).
In Notothenioid fishes found in Antarctic waters, the lipid reserves play an
important role as a primary fuel for energy production, and lipids stored in
muscles help to increase the buoyancy of those fish that lack swim bladders.
Sidell and Hazel (2002) assessed the lipolytic abilities of heart, liver, oxidative
skeletal muscle and adipose tissue from four Notothenioid species using tissue
homogenates as crude enzyme sources and triolein as the substrate. The liver
displayed the greatest lipolytic capacity per 100 g body weight in each of the
studied species.
2.3.3.4. Gene expression
The molecular characterization and nutritional regulation of gene expression
of LPL, HL and PL was investigated in red sea bream, P. major. The study
concluded that LPL, HL and PL gene expression in this marine teleost is under
different regulatory mechanisms with respect to the tissue-specificities and
their nutritional regulation – expression of LPL genes varied during the fasting
and re-feeding process, whereas expression of HL and PL genes was not
significantly affected (Oku et al. 2006).
40
2.3.4. Comparison of fish digestive lipases with those from mammals and
birds
Between animals, comparisons between digestive lipases are restricted to
those from mammals, fish and birds due to the paucity of information from
other sources. Comparing the biochemical and thermodynamic properties of
digestive lipases from dissimilar (and even similar) sources is not a simple
task due to numerous discrepancies (e.g. enzyme purity, identity, activity
assay method used) between the studies. Because of parallel physiological
roles and evolutionary relationships, the main post-gastric digestive lipases
considered in this section are of piscine and mammalian origin.
2.3.4.1. Substrate and fatty acid specificity
A significant distinction between the digestive lipases from fish and pancreatic
lipases from mammals and birds lies in the specificity of substrate and fatty
acid class. From numerous examples given in section 2.3.2, it is clear that in
fish PUFAs are preferentially cleaved over other fatty acids, especially long-
chain saturated fatty acids, and that fish can metabolize wax esters, albeit
slower than TGs and phospholipids. In contrast, digestive lipolysis in
mammals is suited to glycerolipids containing saturated and monounsaturated
fatty acids. Wax ester hydrolysis is not very efficient in mammals (Hargrove et
al. 2004). It must still be noted that human CEL can cleave long-chain PUFAs,
although not as efficiently as piscine CEL (Gjellesvik 1991; Hui and Howles
2002 and references therein). It is yet to be established whether CELs from
other mammals are also able to split PUFA ester bonds.
2.3.4.2. Temperature optimum and stability
Temperature optima and stability of fish digestive lipases vary with habitat. In
temperate water fish, optimum activity is typically between 20 and 40°C and
the enzymes are rapidly deactivated above approximately 40-45°C (see
references in section 2.3.2). Thus, digestive lipases from temperate fish can be
considered slightly more thermally vulnerable than the homologues in land-
based animals that have optima around 37°C and stabilities up to
approximately 50°C (section 2.2.6). An exception to the above general
41
observation was noted with the lipase isolated from dogfish pancreas, where
the enzyme (without substrate) was subjected to temperatures of 45 to 60°C
and showed consistently higher activities over 1 h than commercial porcine PL
(Rasco and Hultin 1988).
2.3.4.3. pH optimum and stability
pH optima in the neutral to moderately alkaline range (pH 6.5-8.5) are a
shared feature of post-gastric digestive lipases from both fish and mammals
(references in section 2.3.2 and Bier 1955 respectively). Avian pancreatic
digestive lipases also share this characteristic. For example, turkey PL has
maximum activity at pH 8.5 (Sayari et al. 2000).
Data on pH stability of fish and mammalian digestive lipases are scarce. A
purified lipase from the hepatopancreas of oil sardine was stable between pH 5
and 9.5 (Mukundan et al. 1985) and intestinal lipase from Nile tilapia was
stable between pH 6.5 and 8.5 (Taniguchi et al. 2001). Mammalian post-
gastric digestive lipases are also generally stable in this pH range (section
2.2.6).
2.3.4.4. Calcium activation and inhibitors
Ca2+
activates mammalian PL by minimizing product inhibition and through
direct interaction with the enzyme (Bier 1955; Winkler and Gubernator 1994).
However, mixed results were reported for the effect of Ca2+
on fish digestive
lipases (refer to examples in section 2.3.2). The effects of other known
activators of lipolytic activity in mammals (e.g. divalent cations other than
Ca2+
, albumin) were rarely studied in fish.
It appears that most of the compounds that inhibit mammalian enzymes, such
as halogen ions and organophosphates, have the same effect on fish lipolytic
activity (e.g. Mukundan et al. 1985; Gjellesvik et al. 1992).
2.3.4.5. Isoelectric point
Isoelectric points were not routinely measured in studies of fish digestive
lipases. A lipase isolated from the pancreas of dogfish had an estimated pI of
5.4, similar to 5.0 of porcine PL (Rasco and Hultin 1988), whereas the pI of
42
Nile tilapia intestinal lipase was 4.9 (Taniguchi et al. 2001). Mammalian PLs
and CELs are also anionic under physiological conditions (section 2.2.6).
2.3.4.6. Molecular weight
One observation is that the few fish CELs that have been purified tend to have
lower molecular weights than their mammalian counterparts. As an example,
Atlantic cod CEL has a molecular weight in the vicinity of 60 kDa (Gjellesvik
et al. 1992), whereas those of human, bovine, murine and porcine CELs,
among others, are in the range of 65 to 100 kDa (Wang and Hartsuck 1993).
CELs isolated from other fish also have molecular weights not exceeding 65
kDa (refer to examples in section 2.3.2). The lower molecular weight of CELs
in fish is related to their shorter primary structure and somewhat different
amino acid composition (see section 2.3.4.7). Pancreatic CEL from mature
Atlantic salmon has 527 amino acids (Gjellesvik et al. 1994). In contrast,
human CEL secreted by both the mammary glands and the pancreas contains
722 amino acids (Baba et al. 1991; Reue et al. 1991 respectively), whereas rat
and bovine pancreatic enzymes are 592 and 579 residues long (Han et al.
1987; Kyger et al. 1989 respectively). The degree of glycosylation further
contributes to the different molecular weights seen between mammalian and
fish CELs, and amongst mammalian CELs. Proline-rich repeats near the end
of the C-terminus in CELs are the site of O-glycosylation (Hui and Howles
2002). These tandem repeats are presumably absent in piscine CELs, as was
found in the Atlantic salmon enzyme (Gjellesvik et al. 1994).
2.3.4.7. Amino acid composition
The amino acid composition (Table 2.2) of Atlantic cod CEL is similar to that
of mammalian CELs, with a few notable exceptions. The cod lipase has very
high glycine content (19%), and high serine and glutamate/glutamine content,
but low proline and lysine content. Low proline is likely to correlate with the
absence of proline-rich repeats at the end of the C-terminal domain in this
enzyme (Gjellesvik et al. 1992; Wang and Hartsuck 1993). An increased
number of glycine residues may provide local mobility, and low proline
content may provide enhanced chain flexibility between secondary structures,
43
reflecting the structural alterations in this relatively psychrophilic lipase
(Georlette et al. 2004).
Table 2.2
Amino acid composition of several purified CELs; the values represent molar
percentages
Amino
acid
Human
pancreasa
Rat
pancreasb
Bovine
pancreasc
Porcine
pancreasd
Canine
pancrease
Atlantic cod
pancreas/pyloric
cecaf
Lys
His
Arg
Asx
Thr
Ser
Glx
Pro
Gly
Ala
Cys
Val
Met
Ile
Leu
Tyr
Phe
Trp
5.0
1.3
2.6
10.6
8.0
5.6
6.3
12.7
9.9
9.3
0.7
7.2
1.7
3.7
6.1
3.7
3.6
1.8
6.3
2.0
3.4
10.5
5.8
5.1
10.8
5.3
8.8
8.9
1.4
8.5
1.4
5.0
9.5
2.1
3.8
1.3
6.0
1.7
3.1
12.0
7.2
6.4
7.7
5.7
8.2
8.2
0.9
5.7
1.9
5.0
8.8
5.2
4.3
2.1
5
1
3
12
7
8
8
10
9
12
0
6
1
4
7
3
4
nd
4.4
1.9
3.5
12.8
5.8
7.1
7.6
10.7
10.5
9.9
nd
6.0
1.8
4.4
6.8
3.6
3.3
nd
1.6
1.4
4.0
9.8
5.0
10.5
11.8
3.9
19.0
8.3
0.4
4.8
1.2
3.3
7.8
3.5
3.8
nd
a(Lombardo et al. 1978);
b(Albro et al. 1985);
c(van den Bosch et al. 1973);
d(Rudd et al. 1987);
e(Abouakil et al. 1988);
f(Gjellesvik et al. 1992); nd = not
determined.
2.3.4.8. Catalytic efficiency
The catalytic efficiency of cod CEL is lower than that of the human enzyme
(Gjellesvik et al. 1992). One explanation for this observation could be related
to the slower rates of digestion in fish than in mammals, i.e. a food mass
44
spends more time in the pyloric ceca and intestine of fish than it does in the
small intestine of mammals. The pyloric cecum in particular provides an
extensive surface area for substrate-enzyme contact and nutrient absorption
(Lopez-Amaya and Marangoni 2000b). Thus, cod and other teleosts containing
pyloric ceca may be able to ‗afford‘ to secrete digestive lipases with relatively
lower catalytic efficiencies.
2.3.5. Potential applications of digestive lipases from marine sources
Due to their high affinity for long chain fatty acids, specificity for particular
fatty acids and regiospecificity (Diaz-Lopez and Garcia-Carreno 2000), fish
digestive lipases may find applications in the synthesis of structured lipids.
Using lipases, TGs could be enriched with certain beneficial fatty acids, like
EPA, DHA or CLA, at a specific position along the glycerol backbone.
Microbial lipases (e.g. from Candida spp.) have been used by several
researchers to improve PUFA content and to selectively extract PUFAs from
fish oils (Hou 2002). Enzymatic synthesis of modified lipids using lipases
could be economically more attractive than chemical synthesis, because of the
lower energy requirements and specificity. In addition, long-chain PUFAs are
highly labile and lipase-catalyzed lipid modifications could prevent
detrimental oxidation and cis-trans isomerization processes. Marine lipases are
likely to have advantages over their microbial or mammalian counterparts
because they may operate more efficiently at lower temperatures. Termination
of enzymatic reactions could be achieved by smaller changes in temperature as
marine lipases often have lower temperature optima and stabilities than lipases
from other sources (section 2.3.4.2). The lipases do not require potentially
hazardous high or low pH media, in contrast to the requirements of many
chemically driven reactions. Fish digestive lipases could also carry out lipid
interesterification reactions (e.g. transesterification) with minimal or no by-
products, a problem often encountered in chemical transformations.
The screening for fish digestive lipases with alkaline pH optima would allow
the identification of suitable enzymes for use in cleaners and household
detergents. Many fish lipases operate most efficiently under slightly alkaline
conditions (section 2.3.4.3). Lipases (mostly of microbial origin) are exploited
45
at an increasing rate by many biotechnological companies (e.g. Novozyme,
Bagsvaerd, Denmark) to provide low temperature, better cleaning
performance.
Some fish digestive lipases are stereospecific, as well as enantiospecific, and
these features could allow the lipases to be used in the synthesis of specialty
pharmaceuticals and agrochemicals, where enantiomerically pure products are
desired. Specialty esters for personal-care products and environmentally-safe
surfactants for applications in detergents and as food emulsifiers may also
await the application of fish digestive lipases.
2.4. Outlook on lipases from marine animals
2.4.1. Impact of biotechnology
Biotechnology has played a significant role in lipase research and
commercialisation, including the cloning and expression of desirable enzymes
in foreign hosts. Genetic modification and/or fermentation is already used in
the production of microbial lipases for commercial purposes. Almost all
lipases currently sold at a commercial level are of microbial origin.
Recombinant DNA technology has made it possible to specifically alter the
DNA sequence of a lipase enzyme in order to study the effects on the primary
structure (and sometimes secondary, tertiary and quaternary) and on enzyme
activity, stability, specificity and many other parameters. Protein engineering
tools have been used successfully to improve the properties of many microbial
lipases (Svendsen 2000; Bornscheuer 2002; Lutz 2004; Olempska-Beer et al.
2006) through either application of recombinant DNA technology, i.e. pre-
translational enzyme modification, or post-translational alteration of the
enzyme, such as glycosylation. Protein engineering can be used to create
lipases with properties that include increased hydrophobicity or hydrophilicity,
improved thermostability, wider substrate specificity, selectivity towards
certain fatty acids, particular enantioselectivity, and alkalophilicity (increase in
net positive charge and pI) (Bott et al. 1994).
Studies of various structural and functional aspects of mammalian, microbial
and plant digestive lipases have all been deepened through protein
46
engineering. This biotechnological technique will likely be applied to marine
lipases in the future in order to facilitate our understanding of fundamental
aspects of these enzymes and allow exploitation of their unique catalytic
properties.
To our knowledge there are no purified marine lipases currently extracted for
large-scale commercial use. As with other marine enzymes, this is because of
a variety of factors such as low extraction yields, variable supply of raw
materials, and environmental and political restrictions (Simpson 2000).
Cloning of marine lipases into microorganisms is a means to overcome some
of these hurdles, and to make these enzymes available at sufficient volumes
and low enough cost to allow their widespread use as industrial enzymes. An
alternative approach is to extract the lipases from a marine raw material, then
immobilize them onto long-lived and re-usable insoluble carriers, as has
already been achieved for several microbial lipases (Christensen et al. 2003).
The immobilization process itself can also be used to further enhance the
stability of the enzymes, extending their catalytic ―life‖ even when used under
harsh conditions of pH, temperature, and in organic solvents (Cao 2005).
2.4.2. Future trends
Researchers and enzyme-producing companies worldwide are actively seeking
―new‖ enzymes from the natural environment to meet growing market
demands for industrial enzymes, including lipases. Thus far, lipases from
aquatic animals have been less well studied compared with their counterparts
from microbial, plant, and terrestrial animals. Furthermore, most of the fish
lipases that have been characterized so far have involved crude or semi-
purified extracts. To allow exploitation of marine lipases and to fully
understand their properties, effective purification methods to produce highly
purified, active and stable forms of these enzymes are required. Such highly
purified forms could then be properly characterized to discover distinct
properties (e.g. superior stability and activity in organic solvents, high
catalytic activity at reduced reaction temperatures, etc.) with these enzymes
that could make them better suited for particular industrial applications. For
large-scale extraction, development of affinity chromatography using a cholate
47
ligand (Wang and Johnson 1983) is a likely route to commercial production,
although at present this has not gone beyond a laboratory scale process. Novel
purification technologies such as reversed micellar systems, aqueous two-
phase systems, enhanced membrane filtration processes, and
immunopurification using highly specific antibodies may also see more
widespread use in future (marine) lipase research (Taipa et al. 1992). High-
throughput activity assays such as acetic acid assay or ―adrenaline test‖
(Schmidt and Bornscheuer 2005) used for routine screening of mutant
lipase/esterase libraries from microorganisms could also be applied to fish
lipase screening once the desired characteristics of these enzymes for
particular applications are established. X-ray crystallography, nuclear
magnetic resonance, and liquid chromatography/mass spectrometry may help
resolve the problems of lipase structure and identity (Anthonsen et al. 1995).
Inexpensive and quick assays for testing the enantioselectivity of lipases are
needed to allow rapid screening of potentially useful enzymes (Gupta et al.
2003b). In this connection, development of lipase-based biosensors would be
useful to enable rapid analysis of samples that would not require pre-treatment
steps to alter and/or contaminate the sample. Large scale extraction and
application of seafood enzymes, sometimes labeled as ―value added‖ products,
is expected to continue to grow as the demand for specialty enzymes increases.
The predominant seafood enzymes in current commercial use are proteases
applied for the production of fish sauces and flavorings and as fish processing
aids for scale and skin removal (Diaz-Lopez and Garcia-Carreno 2000;
Gildberg et al. 2000). Seafood lipases represent the next group of marine
enzymes that are likely to be harnessed for industrial use. In situations where
large-scale recoveries and purification of unique lipases from particular fish
species is impractical, such lipases may be produced in microorganisms by
genetic engineering techniques.
The future of lipase applications is likely to center on the continued functional
improvement and demonstration of their catalytic efficiency, specificity,
stability, and versatility in organic solvents and other nonconventional media.
To this end, protein engineering, immobilization, selection of reaction
conditions, and screening of new enzymes will play pivotal roles.
Environmental concerns will also have an increasing effect on the direction of
48
lipase application research, where energy-intensive processes such as chemical
lipid esterification will be replaced with lipase-catalysed alternatives. Further
growth in lipase applications is expected in food production (e.g. new flavors),
detergents (e.g. greater temperature range), modifications of fats and oils (e.g.
esterification, transesterification, prevention of generation of trans-fats in
margarines, and low-calorie or lipotropic TGs), synthesis of organic
intermediates, and production of biodiesel fuels. The scope of lipase
applications is constantly broadening and this trend will certainly continue.
2.5. Acknowledgements
This work was supported with funds provided by the New Zealand Foundation
for Research, Science, and Technology (FRST) under contract C02X0301, and
the Natural Sciences and Engineering Council (NSERC) of Canada.
49
CONNECTING STATEMENT 1
The next chapter reports on purification and comprehensive evaluation of the
characteristics of Chinook salmon and New Zealand hoki digestive lipases.
Lipase activity assay development, extraction buffer formulation trials and
selection of the most reliable protein assay were carried out prior to the fish
lipase studies and these are reported on in Appendix I. Several purification
techniques were evaluated which allowed the purification sequence to be
established. The techniques that were not used in the ultimate purification
protocol are described as additional experiments in Appendix I.
50
CHAPTER III. PURIFICATION AND PROPERTIES OF DIGESTIVE
LIPASES FROM CHINOOK SALMON (ONCORHYNCHUS
TSHAWYTSCHA) AND NEW ZEALAND HOKI (MACRURONUS
NOVAEZELANDIAE) (MANUSCRIPT 2)
Published in Fish Physiology and Biochemistry, 2010, 36:1041-1060
Authors: Ivan Kurtovic1,2
, Susan N. Marshall2*, Xin Zhao
1 and Benjamin K.
Simpson3
Affiliations:
1Department of Animal Science, McGill University (Macdonald Campus)
21,111 Lakeshore Road, Ste. Anne de Bellevue (QC) Canada H9X 3V9
2The New Zealand Institute for Plant & Food Research Limited, PO Box 5114,
Port Nelson, Nelson, New Zealand
3Department of Food Science and Agricultural Chemistry, McGill University
(Macdonald Campus). 21,111 Lakeshore Road, Ste. Anne de Bellevue (QC)
Canada H9X 3V9
* Corresponding Author: Dr. Susan N. Marshall
The New Zealand Institute for Plant & Food Research Limited
PO Box 5114, Port Nelson, Nelson, New Zealand
Tel: + 643 9897611, Fax: + 643 5467049,
E-mail: [email protected]
3.1. Abstract
Lipases were purified from delipidated pyloric ceca powder of two New
Zealand sourced fish, Chinook salmon (Oncorhynchus tshawytscha) and hoki
(Macruronus novaezelandiae), by fractional precipitation with polyethylene
glycol 1000, followed by affinity chromatography using cholate-Affi-Gel 102,
and gel filtration on Sephacryl S-300 HR. For the first time, in-polyacrylamide
gel activity of purified fish lipases against 4-methylumbelliferyl butyrate has
been demonstrated. Calcium ions and sodium cholate were absolutely
necessary both for lipase stability in the gel and for optimum activity against
caprate and palmitate esters of p-nitrophenol. A single protein band was
present in native polyacrylamide gels for both salmon and hoki final enzyme
51
preparations. Under denaturing conditions, electrophoretic analysis revealed
two bands of 79.6 kDa and 54.9 kDa for salmon lipase. It is proposed that
these bands correspond to an uncleaved and a final form of the enzyme. One
band of 44.6 kDa was seen for hoki lipase. pI values of 5.8 ± 0.1 and 5.7 ± 0.1
were obtained for the two salmon lipase forms. The hoki lipase had a pI of 5.8
± 0.1.
Both lipases had the highest activity at 35°C, were thermally labile, had a pH
optimum of 8-8.5, and were more acid stable compared to other fish lipases
studied to date. Both enzymes were inhibited by the organophosphate
paraoxon. Chinook salmon and hoki lipases showed good stability in several
water-immiscible solvents. The enzymes had very similar amino acid
composition to mammalian carboxyl ester lipases and one other fish digestive
lipase. The salmon enzyme was an overall better catalyst based on its higher
turnover number (3.7 ± 0.3 s-1
vs. 0.71 ± 0.05 s-1
for the hoki enzyme) and
lower activation energy (2.0 ± 0.4 kcal/mol vs. 7.6 ± 0.8 kcal/mol for the hoki
enzyme) for the hydrolysis of p-nitrophenyl caprate. The salmon and hoki
enzymes are homologous with mammalian carboxyl ester lipases.
Keywords: bile salt; calcium; Chinook salmon; enzyme inhibition; hoki;
lipase; organic solvents; zymography
3.2. Introduction
Interest in lipases has been growing steadily in the past two decades because
of the actual and potential uses of these enzymes in applications ranging from
cleaning products to modified foods, flavour development, and large-scale
molecular transformations of lipids in processes such as hydrolysis,
transesterification/biodiesel production, and synthesis of structured lipids.
Mammalian lipases have had traditional uses in applications such as dairy-
product flavour development but it is the enzymes produced by micro-
organisms that have received most attention in recent times. Commercial
microbial lipases are produced extracellularly in high yield from cultures,
helping to make recovery convenient and relatively economical. In addition,
the microbial lipases can be genetically manipulated more easily than those
52
from animals and they tend not to require additional factors such as calcium
and bile salts for activity (as indicated in the review by Kurtovic et al. 2009b).
Despite the apparent advantages of commercial microbial lipases, fish and
shellfish are a largely unexplored source of potentially useful enzymes that
may have different substrate specificities and activity characteristics from the
microbial and mammalian lipases. Fish that inhabit cold and temperate waters
have enzymes with relatively high catalytic efficiencies at low temperatures,
that can be denatured under milder thermal conditions (Georlette et al. 2004),
attributes that may allow lower temperature processing, protection of
substrates from thermal damage and reduced energy costs. Limited studies of
marine digestive lipases indicate that they may also have different reactivities
towards fatty acid classes than their mammalian, plant and microbial
counterparts, with the fish enzymes most actively cleaving long-chain
polyunsaturated fatty acids. Gjellesvik (1991) has demonstrated this with bile
salt dependent digestive lipases from humans and Atlantic cod. A greater
understanding of fish lipases and lipid digestion may also be of benefit in the
development of more effective aquaculture feeds.
Studies of lipolytic and esterolytic enzymes from marine species have been
carried out for many years but few publications have reported on purified
lipases, their characteristics and activities. Challenges associated with the
proteolytic degradation of lipases and conformational instability during
extraction of these enzymes can make purification difficult (Derewenda et al.
1994; Palekar et al. 2000). Despite these challenges, some marine lipases have
been purified, including those from the pyloric ceca/pancreatic tissue of
Atlantic cod (Gadus morhua) (Gjellesvik et al. 1992), intestinal tissue of
tilapia (Tilapia nilotica) (Taniguchi et al. 2001); and hepatopancreas of red sea
bream (Pagrus major) (Iijima et al. 1998) and oil sardine (Sardinella
longiceps) (Mukundan et al. 1985). Studies both of purified and partially
purified lipases (reviewed by Kurtovic et al. 2009) indicate that carboxyl ester
lipase (CEL, also known as bile salt activated lipase or bile salt dependent
lipase) is the dominant digestive lipase in fish.
53
There is only one species of salmon farmed commercially in New Zealand
(Chinook, Oncorhynchus tshawytscha) and the fishery is small on a global
scale, at 7500-8000 tonnes/annum (as gilled and gutted fish). Despite this, the
salmon fishery is the largest volume aquacultured fin-fish species in New
Zealand (Gillard 2008). This fish is ideal as a source of enzymes for study and
potential commercial extraction, as the animals are produced and processed in
a highly regulated and controlled manner, and provide very fresh by-products
of consistent composition (Kurtovic et al. 2006).
Hoki (Macruronus novaezelandiae) is the most important wild-caught
commercial fish in New Zealand with allowable catch set at 90,010
tonnes/annum for the 2008-2009 fishing year (Anonymous 2008). Discards
from the hoki and salmon fisheries are currently used to produce crude plant
fertilisers, fish meals and oils, as well as some higher value omega 3
concentrates (Kurtovic et al. 2006; Shi et al. 2007). There is no commercial
exploitation of the enzyme resource. Before this can occur, a full
understanding is needed both of the characteristics of the enzymes and the
methods needed for economic large-scale extraction.
The aim of this study was to purify and characterize digestive lipases from the
pyloric ceca of New Zealand-sourced hoki and Chinook salmon.
3.3. Materials and methods
3.3.1. Biological materials
Whole viscera from farmed Chinook salmon (each fish weighing
approximately 3 kg) were collected from the gutting line at The New Zealand
King Salmon Co Ltd. (Nelson, NZ) during the winter months (July-August)
and transported on ice directly to the laboratory – a distance of about 5 km.
The fish were anesthetized with AQUI-S™ – an isoeugenol-based food grade
anesthetic (AQUI-S™ New Zealand Ltd., Lower Hutt, NZ) prior to handling
and slaughter. In the laboratory, the pyloric ceca (average weight of 97 g per
cecum) were separated from the rest of the viscera and frozen at -80°C until
required.
54
Hoki pyloric ceca were obtained from a local fish processing plant (Sealord,
Nelson, New Zealand) during the winter months. The ceca were removed from
the rest of the processing discards directly as they came off the filleting line,
stored in ice, and then conveyed to the laboratory located approximately 2 km
from the processing plant. The average weight of a hoki cecum was 17 g. The
ceca were frozen at -80°C until required.
3.3.2. Chemicals
Colipase (porcine), N-Ethyl-N′-(3-dimethylaminopropyl)carbodiimide
hydrochloride (EDAC), p-nitrophenol (p-NP), p-nitrophenyl acetate (p-NPA),
p-nitrophenyl butyrate (p-NPB), p-nitrophenyl caprate (p-NPC) and p-
nitrophenyl palmitate (p-NPP), phenyl methyl sulfonyl fluoride (PMSF),
polyethylene glycol with average molecular weight of 1000 (PEG 1000), and
Sephacryl S-300 HR were purchased from Sigma Chemical Co (St. Louis,
MO, USA). Acrylamide/bis (37.5:1, 30%), Affi-Gel 102, Bio-Lyte pH 3–10
ampholyte, Bio-Safe Coomassie (G-250 stain), IEF pI standards (pI 4.45–9.6),
and SDS-PAGE broad range molecular weight standards were obtained from
Bio-Rad (Hercules, CA, USA). 4-methylumbelliferyl butyrate (MUF-
butyrate), 4-methylumbelliferyl oleate (MUF-oleate), and diethyl p-
nitrophenyl phosphate (paraoxon) were from Fluka Chemical Corp
(Milwaukee, WI, USA). All other reagents were of analytical grade.
3.3.3. Sample preparation
The previously frozen and partially defrosted pyloric ceca (n = 23 for salmon
and n = 40 for hoki) were chopped into small pieces, rapidly re-frozen in
liquid nitrogen, and then powdered in liquid nitrogen using a Waring blender
(Waring Laboratory & Science, Torrington, CT, USA) on high setting for 2
min. The fish tissue powder was first lyophilised and then delipidated
sequentially with several organic solvents, as outlined by Iijima et al. (1998),
using a dry powder to solvent ratio of 1:3. The delipidated tissue (pyloric ceca
powder) was air-dried at 18°C for 15 h then stored at -80°C until needed.
55
3.3.4. Purification protocol
An extraction protocol was developed to optimise fish digestive lipase
recovery and purity. The optimised purification procedure is summarised in
Fig. 3.1.
Fig. 3.1. Purification steps for salmon and hoki digestive lipases.
PBE20: 100 mM phosphate, 2 mM benzamidine, 1 mM EDTA and 20% (w/v)
glycerol, pH 7.8
Purification up to and including the dialysis step was carried out at 4°C, and
affinity chromatography and gel filtration were at 8°C.
Extraction buffer (PBE20) consisted of 100 mM phosphate (pH 7.8), 2 mM
benzamidine, 1 mM EDTA and 20% (w/v) glycerol. Three replicate
purifications were done for each fish, starting with 1 g of pyloric ceca powder,
to which 30 mL of PBE20 were added. The powder was stirred in buffer for
90 min, after which the supernatant was recovered by centrifugation at 12,000
Extraction with PBE20 and recovery of supernatant
Addition of PEG 1000 and recovery of supernatant
Dialysis in PBE20
Affinity chromatography using cholate-Affi-Gel 102
Gel filtration using Sephacryl S-300 HR
Dialysate
PEG 1000 fraction
Pyloric ceca powder
Crude extract
Gel filtration (GF) fraction
Concentration by lyophilisation
Affinity fraction conc.
Affinity fraction
56
g for 15 min using a Beckman Avanti® J-25 I centrifuge (Beckman Coulter,
Inc., Fullerton, CA, USA).
PEG 1000 was used as a gentle method to precipitate non-lipase proteins
(Ingham 1990). It was prepared as a 50% (w/v) solution in PBE20 and then
added to the crude extract to a final concentration of 12%. After 30 min, the
supernatant was recovered by centrifugation at 14,000 g for 12 min. The pellet
was discarded.
Dialysis lasted 21–24 h and used tubing with a 12-14 kDa molecular weight
cut off (MWCO) (Medicell International Ltd., London, UK). Three changes,
each of 4.5 L PBE20, were used. Eight mL of dialysate were applied directly
onto a cholate-Affi-Gel 102 affinity chromatography column and the rest was
divided into 8 mL aliquots and stored at -80°C.
Cholate-Affi-Gel 102 resin was prepared using a carbodiimide coupling
method, based on that of Wang (1980) and the manufacturer‘s (Bio-Rad)
instructions: 45 mL of Affi-Gel 102 were used undiluted; Na cholate (25
µmol/mL gel) was dissolved in 30 mL of 50% dimethylformamide and the pH
was adjusted to 4.5 with 1 M HCl; this was added in portions to the gel with
intermittent end-to-end mixing; EDAC was dissolved in 10 mL of 50%
dimethylformamide to a concentration of 0.85 M and its pH was adjusted to
5.0 with 1 M HCl; EDAC solution was then added drop-wise to the gel
suspension over a period of 5 min, with thorough mixing between additions
(final EDAC concentration in the gel suspension was 0.1 M); the pH of the gel
suspension was adjusted to 5.0 with 1 M HCl and the coupling reaction was
allowed to proceed for 16 h at 20°C; the suspension was then adjusted to pH
8.5 with 1 M NaOH, washed exhaustively in a sintered glass funnel with 1 L
of 50% dimethylformamide, followed by 1 L of 0.5 M NaCl, then 1 L of Milli-
Q water, and finally resuspended in PBE20.
The resuspended resin was degassed, packed into a column (10 cm x 1 cm Ø,
7.8 mL bed volume) and equilibrated with PBE20 at 8°C. All solutions were
degassed prior to application onto the column. The dialysate was applied onto
57
the column at a low flow rate (5–10 mL/h). The column was then washed with
PBE20 to elute unbound proteins, and non-specifically bound proteins were
eluted with 0.4% Na cholate in PBE20 at a flow rate of 30–40 mL/h. Bound
lipase was eluted with 1.2% Na cholate in PBE20 at a flow rate of 30 mL/h, by
collecting 3 mL fractions per tube. Fractions containing lipase activity were
pooled (affinity fraction). Any remaining protein was removed with 2% Na
cholate in PBE20, and the column was re-equilibrated with PBE20, ready for
the next sample application.
The affinity fraction was concentrated by lyophilisation in two 6.5 h stages at
15–20°C with refreezing at -80°C in between the stages to compensate for
sample thawing caused by glycerol. The resulting concentrate was a viscous
liquid.
Sephacryl S-300 HR gel was degassed, packed into a column (100 cm x 1 cm
Ø, 76 mL bed volume) and equilibrated with PBE20 at 8°C. All solutions were
degassed prior to application onto the column. The concentrated affinity
fraction (2.8 mL) was applied onto the column very slowly using a syringe.
Elution was with PBE20 at a flow rate of 20 mL/h. Fractions of 3 mL were
collected in individual tubes and fractions with enzymatic activity were pooled
(GF fraction).
3.3.5. Final preparation of lipase samples
The three GF fractions were pooled, dialysed with three changes, each of 4.2 L
of TBE20 buffer (100 mM Tris-HCl pH 8.0, 2 mM benzamidine, 1 mM EDTA
and 20% (w/v) glycerol) for 24 h using tubing with a 12–14 kDa MWCO, and
then lyophilised using a two stage procedure (7 h for each stage, refrozen at -
80°C in between stages). These final preparations are denoted as SDL (salmon
digestive lipase) and HDL (hoki digestive lipase) in the rest of this document.
The samples were kept at -80°C. SDL and HDL samples were used in all of
the characterization studies to elucidate the properties of the lipases.
58
3.3.6. Protein determination
Protein concentrations were measured using a method based on that of Lowry
et al. (1951) with modifications to the concentration of reagents and
incubation times. Bovine serum albumin (BSA) was used as the standard.
3.3.7. Lipase assays
Lipase activity during purification was routinely measured
spectrophotometrically by hydrolysis of p-NPP, using a modified method
based on that of Winkler and Stuckmann (1979). p-NPP was dissolved in
isopropanol at 30°C to obtain a 15 mM stock solution. A 0.25 mM p-NPP
solution was prepared by adding the stock to pre-warmed (30°C) 20 mM Tris-
HCl buffer (pH 8.0, containing 20 mM CaCl2, 5 mM Na cholate and 0.01%
gum arabic). The substrate-buffer solution was allowed to stand at 30°C for at
least 20 min prior to assay. The reaction mixture comprised an aliquot of the
enzyme sample (typically 20 µL) and the 0.25 mM p-NPP substrate-buffer
solution to make a final reaction volume of 3 mL. The enzyme sample was
replaced with PBE20 buffer to establish blank-rate activity. The release of p-
NP was measured at 410 nm and 30°C in a Unicam (model UV4) UV-visible
spectrophotometer (Thermo Electron Corporation, Waltham, MA, USA) fitted
with a peltier temperature controller. One unit (U) of activity was defined as 1
µmol p-NP released/min under the assay conditions. Using a 0-60 µM
standard curve, the extinction coefficient of p-NP under these conditions was
determined as 16,600 M-1
cm-1
.
For the characterization studies, p-NPC was used in place of p-NPP (see
‗Acyl-chain specificity‘ subsection under ‗Results and Discussion‘),
employing the above procedure and the assay temperature of 35°C. TBE20
buffer was used to establish blank-rate activity.
3.3.8. Electrophoresis and zymographic analysis
Native PAGE of SDL and HDL was carried out using the method of Bollag et
al. (1996), with 7.5% acrylamide gels. Running buffer (25 mM Tris, 192 mM
glycine, pH 8.3) contained 2 mM CaCl2 and 2 mM Na cholate. After protein
separation, zymographic analysis was performed, using a method modified
from that of Diaz et al. (1999): the gels were rinsed in water, then briefly in 20
59
mM Tris-HCl buffer (pH 8.0, containing 20 mM CaCl2, 5 mM Na cholate and
0.01% gum arabic), and then covered by 150 µM MUF-butyrate in the same
buffer. Fluorescent activity bands were revealed by UV illumination at 365
nm. After activity detection, the gels were rinsed well in water and stained
with Bio-Safe Coomassie according to the manufacturer‘s instructions.
SDS-PAGE under reducing conditions was carried out based on the method of
Laemmli (1970), using 12% acrylamide gels. Salmon and hoki GF fractions
were concentrated with 10 kDa MWCO Vivaspin concentrators (Vivascience,
Hannover, Germany) prior to loading. The gels were stained for 2.5 h with
0.25% Coomassie brilliant blue R-250 in 40% methanol, 10% acetic acid
aqueous solution, then destained in two steps: 40% methanol, 10% acetic acid
for 2 h, followed by 7% methanol, 10% acetic acid until the background was
removed satisfactorily.
Isoelectric focusing electrophoresis (IEF-PAGE) was performed based on the
method of Robertson et al. (1987), using 5% acrylamide gels and Bio-Lyte pH
3-10 ampholyte. Salmon and hoki GF fractions were concentrated with 10 kDa
MWCO Vivaspin concentrators prior to loading. Electrophoresis was carried
out at 8°C for 1.5 h at 200 V constant voltage, then increased to 400 V
constant voltage for an additional 1.5 h. After electrophoresis, the gels were
fixed in 0.6 M perchloric acid for 30 min at 20°C, and then rinsed in 10%
acetic acid for 5 min. Staining and destaining were carried out as for SDS-
PAGE.
3.3.9. Acyl-chain specificity
To study the effect of fatty acid chain length on the lipase activity, p-NP
palmitate was replaced with acetate (C2), butyrate (C4) or caprate (C10) esters
of p-NP in the standard assay. The basal activity against these substrates was
tested by omitting Na cholate from the standard assay. In this and all other
characterization studies, the assays were done in triplicate.
60
3.3.10. Effect of pH on the activity and stability
The extinction coefficient (εM) of p-NP as a function of pH was determined
prior to activity analyses. The following buffers (50mM) were used: glycine-
NaOH for pH 11, 10 and 9.5; Tris-HCl for pH 9, 8.5, 8, 7.5 and 7; citric acid /
Na citrate for pH 6. 20 mM CaCl2, 5 mM Na cholate, 0.01% gum arabic, and
1.67% isopropanol (final concentration in standard assay) were included in
each buffer. pNP stock (10 mM) was added to each buffer (pH 7-11) to a final
pNP concentration between 0 and 60 µM (in 10 µM increments). For pH 6
buffer, the final concentration of pNP was between 0 and 350 µM (in 50 µM
increments). These final pNP concentration ranges gave absorbance readings
below 1.0.
For the measurement of activity at different pH, buffers were the same as
above. The substrate was prepared in each buffer to a final conc. of 0.25 mM
and the activity measured as per the standard assay.
For pH stability determination, the following buffers (0.6 M) were used
(Gomori 1955): pH 2.0 (potassium chloride-HCl); pH 4.0 and pH 6.0 (citric
acid / Na citrate); pH 7.0, 8.0 and 9.0 (Tris-HCl); pH 10.0 (glycine-NaOH).
The high molarity of the buffers was necessary to achieve the desired pH when
combined with the sample. The ratios of buffer to sample needed to achieve
the required pH during incubation were: pH 2 – 10:1; pH 4 – 4:1; pH 6 – 5:1;
pH 7 – 4:1; pH 8 – 3:1 (sample pH; chosen since minimum ratio for others);
pH 9 – 3:1; pH 10 – 4:1. The sample was incubated with the buffer solutions
for 30 min at 20°C, prior to standard assay.
3.3.11. Effect of temperature on the activity and stability
The lipase activity at different temperatures was measured by equilibrating the
substrate solution at various temperatures (20 to 65°C, in 5°C increments) for
10 min prior to sample addition and standard assay. The lower limit was 20°C
due to substrate precipitation below this temperature. The initial rate of
reaction within the first 1 min was used to calculate the activity.
The temperature coefficients (Q10, Whitaker 1994) for Chinook salmon and
hoki lipase-catalyzed hydrolysis of p-NPC were obtained from the initial rates
of reaction at 20 and 35°C.
61
To investigate the effects of temperature on lipase stability, the sample was
incubated at various temperatures (0 to 60°C, in 10°C increments) for 30 min
using a digital block heater, then rapidly cooled in ice to 0°C, prior to standard
assay.
3.3.12. Enzyme kinetics
The kinetic parameters (Vmax and Km) were determined using a Hanes plot
([S]
/vo against [S]). The turnover number (k2, in units of s-1
) was calculated
from Vmax and enzyme molecular weight (54.9 kDa for the proposed final form
of salmon lipase and 44.6 kDa for hoki lipase). The substrate concentration
range for the kinetic experiment was 0-250 µM p-NPC and reaction
temperature was 35°C. As in the standard assay, TBE20 buffer was used
instead of sample for determination of blank activity at each concentration.
3.3.12.1. Inhibition with paraoxon
The hydrolysis of p-NPC at a sub-Km concentration (50 µM) was followed in
the presence of 0-5 µM paraoxon. Paraoxon stock solution (1 mM) was
prepared in isopropanol and added to the assay at 35°C. For each inhibitor
concentration, the time-course of the reaction was followed for 8 min. First-
order rate constants (kapp values) for each paraoxon concentration were
determined by nonlinear least-squares fitting to equation 1 (Hosie et al. 1987):
[1] %A = (Δ%A)e-kappt
+ %A∞
Here, %A is the % activity remaining at various times, Δ%A is the difference
between the % activity at time 0 and at infinite reaction time, and %A∞ is the
% activity remaining at infinite reaction time. Once kapp values were known, Ki
and k2 values for the inhibition were determined by nonlinear least-squares
fitting to equation 2 (Hart and O'Brien 1973):
k2[I]
[2] kapp = _______________________
Ki(1 + [S]/Km) + [I]
Here, [I] is the inhibitor concentration, and [S] is the substrate concentration.
62
3.3.13. Thermodynamic parameters
The activation energy (Ea), in units of calories per mole (1 cal = 4.1868 J), was
calculated from the Arrhenius equation by plotting ln activity against 1/T (in
units of Kelvin). Slope = -Ea/R, where R is the gas constant (1.986 cal K-1
mol-
1). The temperature range was 20-30°C for salmon and 20-35°C for hoki. The
enthalpy of activation (ΔH*), entropy of activation (ΔS*), and free energy of
activation (ΔG*), all at 35°C, were calculated using the following equations:
[3] ΔH* = Ea – RT
[4] ΔS* = 4.576 (log k2 – 10.753 – log T + Ea/4.576T)
Here, k2 is the turnover number (s-1
), defined previously.
[5] ΔG* = ΔH* – TΔS*
(Lehrer and Barker 1970; Simpson and Haard 1984)
3.3.14. Effect of different bile salts on activity
Na cholate was replaced in the standard assay with Na taurocholate or Na
deoxycholate. Five and 10 mM assay concentrations were tested for cholate
and taurocholate, and 1 mM assay concentration was used for deoxycholate.
No incubation with a bile salt prior to activity assay was carried out, as was
performed in some earlier studies (e.g. Aryee et al. 2007).
3.3.15. Effect of selected chemicals on activity/stability
The effect of these chemicals on the fish lipases was examined: PMSF, KI, the
chlorides of Fe, Hg, Zn, Mg and Mn, and NaN3. Stock solutions (10 and 2 mM
for PMSF; 20 and 2 mM for the others) were prepared in Milli-Q water, except
for PMSF which was prepared in 10% isopropanol. The sample was incubated
with the chemical in 1:1 ratio for 30 min at 20°C, prior to standard assay.
Incubation without an inhibitor (i.e. just water or 10% isopropanol) provided
the control activity.
Similarly, the effect of detergents (Triton X-100 and Tween 80) was tested by
direct incubation with the sample, prior to standard assay. Stock solutions (20
and 2%) were prepared in TBE20 and incubation with this buffer served as
control.
63
The effect of colipase was tested by incubating the sample with colipase in 1:1
ratio for 10 min at 20°C prior to standard assay. Porcine colipase solutions (20
and 2 µM) were prepared in TBE20 and incubation with this buffer served as
control.
3.3.16. Stability in organic solvents
The effect of water-miscible solvents (acetone, methanol and isopropanol) was
tested by direct incubations with the sample for 30 min at 20°C, prior to
standard assay. Pure solvent and a 20% solution prepared in TBE20 were
used. Incubation with the TBE20 buffer served as control.
For water-immiscible solvents (hexane, heptane and isooctane), a modified
method based on that of Sztajer et al. (1992) was followed. The sample was
adsorbed onto a square (1cm2) of filter paper (Whatman No.4) fitted inside
cuvette and incubated with excess solvent (300 µL) for 30 min at 20°C. The
solvent was evaporated under a stream of nitrogen, and the substrate solution
added straight to cuvette. No extraction of sample from the filter paper was
carried out. Instead of rate measurement, the fixed end point method was used
and the absorbance was read after thorough mixing at intervals over a 10 min
period. The temperature of the cuvette was maintained at 35°C during this
period. For the control experiment, the adsorbed sample was incubated in air.
TBE20 buffer adsorbed onto the paper instead of sample served as the blank
activity for all incubations.
3.3.17. Amino acid composition and N-terminal sequencing
SDL and HDL samples were prepared by using centrifugal filter devices to
replace the buffer with Milli-Q water. The samples then underwent 24 h gas
phase hydrolysis with 6 M HCl at 110°C. High sensitivity amino acid analysis
was done by reversed phase HPLC of derivatized amino acids.
For N-terminal sequencing, the samples were concentrated under vacuum and
automated Edman degradation using an Applied Biosystems 494 Procise
Protein Sequencing System and the ‗Pulsed Liquid‘ sequencing method was
carried out (studies conducted by Australian Proteome Analysis Facility,
Macquarie University).
64
3.4. Results and discussion
3.4.1. Sample preparation
In order to maintain lipase activity, it was crucial to remove free water prior to
application of solvents to the crushed pyloric ceca. When the ceca were
delipidated without first lyophilising them, most of the lipase activity was lost.
This activity loss is attributed to the removal of a water monolayer around the
lipase, together with the free water, by the hydrophilic solvents (e.g. acetone)
used during delipidation. It appears that after lyophilisation the water
monolayer around the lipase is held tightly, shielding the enzyme from direct
interaction with the bulk solvent and allowing it to maintain its native state
(Zaks and Klibanov 1988; Klibanov 1989).
3.4.2. Lipase stability during purification and choice of buffer
The hoki and salmon lipases were both found to be very labile and buffers
needed to be developed to offer some assistance with maintaining
conformational stability and activity during purification procedures.
Preliminary extracts of pyloric ceca powder using the buffer of Gjellesvik et
al. (1992) - TBE (25 mM Tris-HCl, 2 mM benzamidine and 1 mM EDTA) in
the pH range 8 to 9, were very unstable, in that all lipase activity was lost
within 24 h at 4°C. Addition of 20 mM Na cholate to this extraction buffer
(pH 8.5) improved the stability considerably, with all of the activity remaining
up to 24 h under the same storage conditions. Besides contributing to higher
stability of the crude extract (Blackberg and Hernell 1993; Loomes 1995),
inclusion of Na cholate allowed for more complete extraction and full
activation of lipase. Despite this, Na cholate was not included in the final form
of the extraction buffer (as used in the protocol in Fig. 3.1) as it interfered with
lipase binding to the cholate affinity resin. Dialysis using cholate-free TBE
buffer prior to the affinity step did not improve binding. It is thought that some
cholate remained bound to the lipase, affecting the degree of binding and
leading to inconsistent recoveries and degrees of purification.
Addition of glycerol to TBE buffer resulted in improved extraction activity
and higher stability of the lipases. This effect was proportional to the amount
65
of glycerol added, up to 20% (w/v). Concentrations higher than 20% produced
fairly viscous solutions, which made subsequent purification steps more
difficult. Subsequent trials found that the stability of a crude extract over a 4-
day period was higher in phosphate buffer. PBE20 (100 mM phosphate, 2 mM
benzamidine, 1 mM EDTA and 20% (w/v) glycerol, pH 7.8) was therefore
selected as the extraction/purification buffer in the ultimate protocol.
3.4.3. Lipase assay
Highest lipase activity was obtained with 0.25 mM of the substrate (p-NPP or
p-NPC). Several-fold higher p-NPP concentrations were used in some earlier
studies (e.g. Winkler and Stuckmann 1979; Pencreac'h and Baratti 1996), but
in our study, increasing the substrate concentration above 0.25 mM resulted in
slightly lower activity. At this time, it is unknown whether this effect is due to
differences in the composition of substrate-buffer solution affecting substrate
accessibility, or some form of substrate inhibition.
Very low activities were measured in the absence of calcium ions and Na
cholate, such that their inclusion in the assay improved the lipase activity
approximately 100-fold. Calcium ions prevent product inhibition of the active
site by precipitating fatty acids following substrate hydrolysis, and may act as
a cofactor by binding to a specific site on the enzyme (Bier 1955; Leger et al.
1977; Winkler and Gubernator 1994; Anthonsen et al. 1995). The bile salt, Na
cholate, acts a surfactant, facilitating improved lipid binding and substrate
access by lipases. More importantly, Na cholate specifically activates CELs
via direct binding to the enzyme (Blackberg and Hernell 1993; Wang et al.
1997; Hui and Howles 2002).
3.4.4. Purification of salmon and hoki lipases
The purification steps for salmon lipase are presented in Table 3.1. The results
for hoki lipase have been omitted for brevity but follow a similar pattern. The
crude extract obtained from salmon pyloric ceca had approximately three
times higher specific activity than the hoki equivalent. This is likely to be a
reflection of the Chinook salmon diet, warmer habitat and less traumatic
66
harvest method (Kurtovic et al. 2006). Whereas farmed salmon are fed a
uniform high-lipid diet, the diet of predatory hoki is highly variable.
Table 3.1
Purification scheme for Chinook salmon digestive lipase
Purification step Total
volume (mL)
Protein conc.
(mg/mL)
Total protein
(mg)
Total activity
(U)
Specific
activity (U/mg)
Activity
recovery (%)
Purification
(-fold)
Crude extract 27 ± 0 11.5 ± 0.6 307 ± 13 109 ± 13 0.36 ± 0.06 100 1
PEG 1000
fraction
33 ± 0 8.3 ± 0.3 271 ± 7 82 ± 3 0.30 ± 0.02 75 ± 8 0.9 ± 0.1
Dialysate 65 ± 1 3.0 ± 0.2 193 ± 9 72 ± 32 0.37 ± 0.14 65 ± 20 1.0 ± 0.2
Affinity fraction 36 ± 3 0.03 ± 0 1.0 ± 0.2 3.2 ± 0.6 3.1 ± 0.2 24 ± 1* 8.9 ± 1.3
Affinity fraction
conc.
9 ± 1 0.11 ± 0.02 0.9 ± 0.1 3.2 ± 0.6 3.4 ± 0.4 25 ± 1* 9.6 ± 0.6
GF fraction 19 ± 2 0.010 ± 0.004 0.20 ± 0.09 0.9 ± 0.2 4.8 ± 1.3 25 ± 5* 14 ± 5
These results are compiled from three separate purifications. Values are means
± SD. *Recovery back-calculated as if all material from previous step was
used for further purification.
Dialysis was used to remove lower molecular weight contaminants, including
PEG 1000, and proteins less than ~10 kDa that could interfere with binding,
before sample application onto the affinity column. Based on the specific
activity, cholate-Affi-Gel 102 was more selective for the salmon enzyme,
where an almost two-fold higher purification was obtained for salmon lipase in
comparison to hoki lipase.
The overall recovery of activity from the affinity resin was variable. This was
especially apparent for the salmon sample, which ranged from 70 to 90%
recovery of the applied activity. Recovery of 90–100% was obtained for the
hoki samples. For the removal of low affinity bound proteins, 0.4% Na cholate
was chosen as a compromise between satisfactory removal of non-lipase
proteins and minimal removal of lipase. Lipase was removed completely with
1.2% Na cholate, while some of the more tightly bound proteins were left
behind. Lipase was eluted in a single protein and activity peak. The recovery
of lipase eluted with 1.2% cholate was typically 35–40% for salmon (relative
to the applied dialysate activity), and 60–70% for hoki. Gjellesvik et al. (1992)
67
reported lower recovery values from a cholate-Sepharose resin in the course of
cod lipase purification.
Elution of lipase from cholate-Affi-Gel 102 with 2 or 2.5% Na taurocholate
(Gjellesvik et al. 1992) was unsuccessful, mainly because it caused the gel to
compact until no flow was possible. For the same reason, 0.5–1% Na
deoxycholate (Wang 1980; Wang and Johnson 1983) was found unsuitable for
the removal of non-specifically bound proteins. These researchers used
cholate-Sepharose rather than cholate-Affi-Gel 102. We consider it likely that
the compacting was due to the dissimilar solubility and hydrophilic properties
of cholate, taurocholate, and deoxycholate, and their interaction with the resin.
It was necessary to concentrate the affinity fraction to ensure sufficient activity
was applied to the gel filtration column. Initial attempts to concentrate the
affinity fraction were carried out with 10 kDa MWCO Vivaspin concentrators
or 10 kDa MWCO Amicon Ultra centrifugal filter devices (Millipore
Corporation, Billerica, MA, USA), but more than half of the activity was lost
with this approach. A similar outcome was reported by Aryee et al. (2007),
and Gjellesvik et al. (1992) reported destabilization of cod affinity fraction
upon concentration. In our study, lyophilisation in the presence of glycerol and
cholate was used as a successful alternative approach for sample
concentration, resulting in a viscous solution rather than a powder. The lipase
was very stable in this solution, with high glycerol (75-80%) and cholate
(4.6%) concentrations.
A single lipase activity peak was eluted from the gel filtration step, for both
salmon and hoki samples. The salmon lipase activity peak comprised of two
partially resolved protein peaks (Fig. 3.2), whereas the hoki lipase activity
peak corresponded to a single protein peak. An equivalent increase in
purification was obtained for both salmon and hoki lipases following this step,
with no loss of lipase activity between the affinity concentrate and GF
fraction.
68
0
0.01
0.02
0.03
0.04
0.05
0.06
5 10 15 20 25 30
A280
(
)
or
Lip
ase
act
ivit
y (
U/m
l) (
)
tube number
Fig. 3.2. A representative elution profile of salmon lipase on Sephacryl S-300
HR. Tubes 15-20 (indicated by dashed line) were pooled as the GF fraction
Markedly varying recoveries and purifications have been reported for purified
fish digestive lipases (Mukundan et al. 1985; Gjellesvik et al. 1992; Iijima et
al. 1998). In general, relatively lower recoveries were obtained for highly
purified preparations (e.g. Iijima et al. 1998). Although lower final purification
values (fold) were achieved in the present study (14 ± 5 for salmon and 7.4 ±
1.3 for hoki lipase), the enzyme activity recoveries in the final fractions (25 ±
5% for salmon and 48 ± 3% for hoki lipase) were higher than in most of the
previously published studies. This difference may be due to the relatively high
specific activities in the starting material seen in the present study. Using TBE
buffer containing 20 mM Na cholate, the specific activity in the crude extract
was <0.2 U/mg and <0.1 U/mg for salmon and hoki samples respectively, but
with our PBE20 buffer, the activity was 0.36 and 0.13 U/mg.
3.4.5. Preparation of SDL and HDL
The pooled GF fractions were dialysed to remove phosphate from the samples,
making zymographic analysis possible. Lyophilisation facilitated in-gel
activity detection by providing a 4-fold concentrated sample and allowed for
easier storage of the final preparations. As was the case with the affinity
fractions, very little activity was lost after lyophilising the GF dialysates. The
69
lipase was again very stable (> 3 days at 4°C) in this viscous solution
containing 80% glycerol.
3.4.6. Electrophoresis and zymographic analysis
For the first time, in-gel activity of purified fish lipases has been demonstrated
(Fig. 3.3).
a b
Fig. 3.3. Zymograms, (a) purified hoki and salmon digestive lipases (3.6 µg
HDL and 1.7 µg SDL), (b) 4.2 µg SDL (left) and 300 µg salmon crude extract
(right)
After UV illumination, fluorescent activity bands became visible in 2–10 min.
A single activity band was present in all lanes.
For successful in-gel activity detection, low % acrylamide gels (7.5%) were
used. The absence of activity in more rigid gels could be due to the impaired
flexibility in the enzyme structure needed for catalysis. Both SDS and Triton
X-100 (as used by Diaz et al. 1999) were found inhibitory to fish lipases and
were omitted. Calcium ions and Na cholate were absolutely necessary for
stability during electrophoresis and were included in the running buffer.
When MUF-oleate was used instead of MUF-butyrate, it took much longer (3-
4 h) for any fluorescence to appear. It seems likely that this is due to slower
permeation of the bulkier substrate into the gel.
70
The removal of phosphate from the samples (by dialysis in TBE20) was
necessary in order to avoid the formation of calcium phosphate precipitate in
the gel. When the native gels were run with the samples in phosphate buffer
(PBE20), Ca phosphate precipitate interfered with lipase stability in the gel
and no activity was detected.
When the native gels were subsequently stained with Bio-Safe Coomassie,
images as shown in Fig. 3.4 were obtained.
Fig. 3.4. 7.5% acrylamide native-PAGE of purified hoki and salmon digestive
lipases. Lane 1, 3.6 µg HDL; lane 2, 1.7 µg SDL; lane 3, 4.2 µg SDL; lane 4,
300 µg salmon crude extract
A single band, responsible for the lipase activity, was present in HDL and
SDL lanes.
Fig. 3.5 shows the SDS-PAGE analysis of the purified lipases.
2 1 3 4
71
Fig. 3.5. 12% acrylamide SDS-PAGE, (a) 4.5 µg purified salmon digestive
lipase, (b) 10 µg purified hoki digestive lipase. A concentrated gel filtration
fraction was loaded in duplicate. Molecular weight standards (kDa) are in the
outside lanes
In 12% acrylamide SDS-PAGE, salmon lipase lanes displayed two bands with
relative mobility corresponding to an average molecular weight of 79.6 kDa
and 54.9 kDa (2 trials). Fish digestive lipases studied to date have molecular
weights ranging from 46 to 64 kDa. CELs isolated from mammals have
molecular weights ranging from 60 to 100 kDa, depending on the size of the
proline-rich region near the C-terminus (reviewed by Kurtovic et al. 2009a).
The two salmon proteins seen in Fig. 3.5 (79.6 kDa and 54.9 kDa) may be
lipase isozymes. However, it is also possible that they are the same enzyme
but one has been subjected to partial proteolytic cleavage during purification,
as was proposed for the porcine pancreatic CEL (Rudd et al. 1987; Wang and
Hartsuck 1993). The possible presence of an uncleaved and a final form is
supported by the following observations: a single activity band was seen in the
native gels and this was the only protein band revealed after Coomassie
staining (Figs. 3.3 and 3.4); the lipase activity peak eluted from the gel
filtration column comprised of two partially resolved protein peaks (Fig. 3.2);
200
116
97.4
66.2
45
31 a b
72
and two proteins of very similar pI values were revealed by IEF-PAGE
analysis (see below).
Purified hoki lipase was seen as a homogenous band corresponding to a
molecular weight of 44.6 kDa (3 trials). This molecular weight is at the lower
end of fish digestive lipases studied to date (as indicated in the review by
Kurtovic et al. 2009a).
IEF-PAGE analysis (5 trials) revealed one protein band in the hoki sample,
whereas two bands were seen for salmon. pI values of 5.8 ± 0.1 for hoki
lipase, and 5.8 ± 0.1 and 5.7 ± 0.1 for the two salmon lipase forms were thus
obtained. Mammalian bile salt activated lipases and fish lipases (very limited
data available) have pI values in the range of 4 to 6 (reviewed by Kurtovic et
al. 2009a).
3.4.7. Acyl-chain specificity
The activity profile against different p-NP esters was very similar for both
lipases, and they exhibited the highest activity against a medium chain (C10)
ester of p-NP. This activity was approximately 50% higher compared to that
against p-NPP (C16) for the salmon enzyme (Fig. 3.6) and up to 43% higher
than that against p-NPP for hoki. p-NPC was therefore used in the present
study for all the characterization experiments.
Previous studies have reported a range of chain length specificities for
different lipases using the fatty acid esters of p-NP as substrate. Aryee et al.
(2007) found the highest activity against p-NPP for grey mullet lipase,
whereas Gjellesvik et al. (1992) reported the highest activity towards the
caproyl (C6) ester of p-NP with cod lipase. Lipolytic crude extract from
Penicillium aurantiogriseum was most active against p-NPC (Lima et al.
2004), while Rhizopus chinensis lipase exhibited highest activity with p-NP
caprylate (C8) as the substrate (Sun et al. 2009).
In our study, p-NPA and p-NPB exhibited very high rates of spontaneous
hydrolysis and this may have been partially responsible for the low activities
with these substrates in the above, as well as in our study. The high blank rates
73
are likely due to the more hydrophilic nature of short chain, acetate and
butyrate fatty acids.
In the absence of any bile salts, both lipases had basal activity against short
chain fatty acid substrates, but very little against the medium chain and no
activity against the long chain. This is a feature of CELs (Wang and Hartsuck
1993) and the two fish lipases likely belong to this group.
Fig. 3.6. Acyl-chain specificity of Chinook salmon lipase with p-nitrophenyl
esters. Activities are relative to p-NPC. b = basal activity in the absence of Na
cholate. Values are means ± SD (n = 3)
3.4.8. pH optimum and pH stability
The extinction coefficient of p-NP was determined at different pH values.
These are shown in Table 3.2.
Table 3.2
Extinction coefficient of p-NP as a function of pH
pH 6 7 7.5 8 8.5 9 9.5 10 11
εM (M-1
cm-1
) 2300 11800 15000 16600 17100 16900 17000 17200 17200
Values are averages of duplicate determinations.
The highest εM values were in the pH range of 8-11. This is consistent with
findings from previous studies and the actual values are very close to the ones
reported by other researchers (e.g. 12750 at pH 8 (Pencreac'h and Baratti
0
20
40
60
80
100
2 2b 4 4b 10 10b 16 16b
Rel
ati
ve
act
ivit
y (
%)
Fatty acid carbon number
74
1996); 15000 at pH 8 (Kordel et al. 1991); 14000-19000 in the pH range 8-10
(Lima et al. 2004).
Chinook salmon and hoki lipases both showed very similar pH optima (8 and
8.5, respectively) for the hydrolysis of p-NPC (Fig. 3.7). The salmon enzyme
exhibited a high activity over a pH range (8-9.5). pH 6 was the lowest pH
tested due to lack of absorbance of p-NP at acidic pH and limited solubility of
Na cholate below this pH, however it is clear that there would be no
significant activity below this pH (Fig. 3.7). Spontaneous substrate hydrolysis
occurred at pH 9.5-11, being very high at pH 11, and this was accounted for in
the final results. Owing to its lower specific activity, measurements of the hoki
enzyme activity included greater errors at most pH values. The results for the
salmon and hoki lipases are typical of mammalian and fish lipases, where the
highest activity is at neutral to moderately alkaline pH (reviewed by Kurtovic
et al. 2009a).
Fig. 3.7. Activity of Chinook salmon (—) and hoki (– – –) lipase against p-
NPC as a function of pH. The buffers used are listed in section 3.3.10. Values
are means ± SD (n = 3)
Almost identical patterns are seen regarding the effect of pH on lipase
stability, with both enzymes retaining more than 60% of activity between pH
4-8 and being most stable at pH 7 (Fig. 3.8). Crab digestive lipase (Cherif et
al. 2007) was also stable in this pH range, but in addition had significantly
0
20
40
60
80
100
5 6 7 8 9 10 11 12
Rel
ati
ve
act
ivit
y (
%)
pH
75
higher residual activities at pH 8-10 than the two lipases in our study. A lipase
from the hepatopancreas of oil sardine was stable between pH 5 and 9.5
(Mukundan et al. 1985) and intestinal lipase from Nile tilapia was stable
between pH 6.5 and 8.5 (Taniguchi et al. 2001). Thus, the Chinook salmon
and hoki enzymes appear more acid stable compared to other fish lipases.
Fig. 3.8. Effect of pH on the stability of Chinook salmon (—) and hoki (– – –)
lipase. Residual activities are relative to 20°C data from the temperature
stability experiment in order to remove the temperature effect during
incubation. The buffers used are listed in section 3.3.10. Values are means ±
SD (n = 3)
3.4.9. Temperature optimum and thermal stability
The salmon lipase displayed two temperature optima for the hydrolysis of p-
NPC, at 35-40°C and 50°C (Fig. 3.9). This was verified with a two-sided,
unpaired Student‘s t-test (p < 0.05), applied to both 40/45°C and 40/50°C pairs
of data. The two optima of salmon lipase may correspond to each of the lipase
forms visualized on SDS and IEF-PAGE. The hoki lipase was most active at
35°C. The temperature optima of fish lipases vary with the habitat of the
organism and are typically close to 35°C (Mukundan et al. 1985; Gjellesvik et
al. 1992; Taniguchi et al. 2001).
0
20
40
60
80
100
120
140
160
1 2 3 4 5 6 7 8 9 10 11
Res
idu
al
act
ivit
y (
%)
pH
76
Fig. 3.9. Activity of Chinook salmon (—) and hoki (– – –) lipase against p-
NPC as a function of temperature. Values are means ± SD (n = 3)
Based on the temperature-activity data, the temperature coefficients (Q10) for
the hydrolysis of p-NPC by chinook salmon and hoki lipases were calculated
as 1.22 ± 0.10 and 1.54 ± 0.14, respectively, for 20 to 35°C temperature range.
Aryee et al. (2007) reported an average Q10 of 1.3 for grey mullet lipase with
pNPP as substrate (temperature range of 20 to 50°C). Our results suggest that
Chinook salmon and hoki lipases have relatively high hydrolytic activity at
lower temperatures, and are not very sensitive to temperature changes up to
the optimum temperature for hydrolysis.
Both lipases were thermally labile (Fig. 3.10) and lost significant amount of
activity when incubated at above 30°C. The salmon lipase was more stable
than the hoki up to 30°C. This is likely a to be a reflection of the habitat
temperatures of these teleosts with hoki being a mid-depth fish and penned
salmon living close to the surface, but neither experiencing water temperatures
above 18°C. There was a small thermal activation effect observed for the
salmon enzyme exposed to 20°C for 30 min. This is an interesting property
that could be exploited in applications of this enzyme. The thermal
vulnerability of salmon and hoki lipases is a feature shared with other fish
lipases (as indicated in the review by Kurtovic et al. 2009a) and could be
0
20
40
60
80
100
15 25 35 45 55 65
Rel
ati
ve
act
ivit
y (
%)
temperature ( C)
77
useful in low temperature applications of these enzymes where relatively low
heat treatments could be applied to terminate the reaction.
Fig. 3.10. Effect of temperature on the stability of Chinook salmon (—) and
hoki (– – –) lipase. Residual activities are relative to incubation at 0°C. Values
are means ± SD (n = 3)
3.4.10. Kinetic parameters
The Lineweaver-Burke plot has previously been used in several fish enzyme
kinetic studies (e.g. Mukundan et al. 1985). However, the Hanes modification
of the Michaelis-Menten equation was chosen for the determination of kinetic
values in this study since the experimental errors retain a constant contribution
over the whole range of substrate concentrations.
The kinetic parameters for salmon lipase with p-NPC as substrate were Km of
78 ± 8 µM and a k2 of 3.7 ± 0.3 s-1
. The Km for the hoki enzyme was 68 ± 13
µM with a k2 of 0.71 ± 0.05 s-1
. The two lipases have similar affinities for p-
NPC, based on the Km values. These values are relatively low, indicating high
substrate affinity, when compared to the corresponding values for several
other fish lipases: 140 ± 40 µM for cod CEL with p-NP caproate as substrate
(Gjellesvik et al. 1992); 0.3 and 0.04 M for sardine lipase with triacetin and
tributyrin, respectively (Mukundan et al. 1985); and 58-157 µM for tuna
esterase with various short chain fatty acid esters of p-NP and N-methyl
indoxyl (Maurin and Gal 1996). It is evident that the diversity of substrates
used to estimate Km values makes direct comparisons between studies difficult.
0
20
40
60
80
100
120
0 10 20 30 40 50 60
Res
idu
al
act
ivit
y (
%)
temperature ( C)
78
Salmon lipase has approximately five-fold higher turnover number (k2) than
the hoki lipase, in agreement with the higher specific activity of the salmon
enzyme. A k2 value of 193 ± 21 s-1
(at 25°C) was reported for the cod lipase. In
the same study, the corresponding value for the human CEL was 1670 ± 48 s-1
.
Reports of k2 values for other fish lipases are scarce as few enzymes have been
purified sufficiently. The catalytic efficiency (k2/Km) of the three fish lipases is
much lower than that of the human enzyme (Km of 44 ± 6 µM). One
explanation for this could be related to the slower rates of digestion in fish
than in mammals, i.e. a food mass spends more time in the pyloric ceca and
intestine of fish than it does in the small intestine of mammals. The pyloric
cecum in particular provides an extensive surface area for substrate-enzyme
contact and nutrient absorption (Lopez-Amaya and Marangoni 2000b). Thus,
cod and other teleosts containing pyloric ceca may be able to ‗afford‘ to
secrete digestive lipases with relatively lower catalytic efficiencies.
3.4.10.1. Inhibition by an organophosphate
Lipases are irreversibly inactivated by organophosphates. We studied the
mechanisms of this type of inhibition with paraoxon as a representative
organophosphate. The requirement for pseudo first-order kinetics was satisfied
by ensuring that both the inhibitor (1-5 µM) and substrate (50 µM)
concentrations were much greater than lipase active sites (4 nM for salmon, 11
nM for hoki). Thus, the inhibition rate of the lipase was directly proportional
to the decrease in substrate hydrolysis rate (Hart and O'Brien 1973; Hosie et
al. 1987). The inhibition showed saturation kinetics with increasing paraoxon
concentration.
Fig. 3.11 shows the obtained kapp values against the inhibitor concentration for
each enzyme. The stoichiometry of the inhibition was determined as 0.61 ±
0.07 for the salmon and 0.79 ± 0.05 for the hoki lipase, from the slope of the
log kapp vs. log [paraoxon] plot (Abouakil et al. 1989). It is therefore likely that
the inhibitor and each lipase active site react in a 1:1 ratio.
Ki (inhibitor concentration for 50% inhibition) was calculated as 2.28 ± 0.43
µM and 5.17 ± 1.89 µM for the salmon and hoki enzymes, and k2 for the
inhibition was 0.052 ± 0.006 s-1
and 0.064 ± 0.016 s-1
for salmon and hoki
79
lipases, respectively. These Ki values are several fold higher compared to that
reported for the inhibition of porcine CEL by p-nitrophenyl-N-butyl carbamate
(Hosie et al. 1987), and lower than the value reported for the inhibition of cod
CEL by di-isopropyl fluorophosphate (Gjellesvik et al. 1992).
Fig. 3.11. Michaelis-Menten plot for paraoxon inhibition of Chinook salmon
(—) and hoki (– – –) lipases. The markers indicate kapp values obtained
experimentally and the lines show the theoretical curves calculated from
equation 2 using the estimated values of Ki and k2 for the inhibition. Values are
means ± SD (n = 3)
3.4.11. Thermodynamic parameters
The Arrhenius plots used to obtain the activation energies for the hydrolysis of
p-NPC by Chinook salmon and hoki lipase are shown in Figs. 3.12 and 3.13,
respectively. The Ea and other thermodynamic values are presented in Table
3.3. Chinook salmon lipase achieved a very low Ea value, similar to the one
reported for grey mullet lipase / p-NPP hydrolase reaction (1.94 kcal/mol,
Aryee et al. 2007). The hoki enzyme, however, achieved a value more
comparable to beef fat hydrolysis by mammalian pancreatic lipase (4.6
kcal/mol, German et al. 2002) and Greenland cod trypsin amidase reaction
(6.6 kcal/mol, Simpson and Haard 1984). The lower activation energy attained
with the salmon lipase is consistent with the higher value for the enzyme‘s
turnover number, in that both indicate that the salmon enzyme is a more
efficient catalyst than the hoki enzyme.
0
0.005
0.01
0.015
0.02
0.025
0.03
0.035
0 1 2 3 4 5
ka
pp
(s-1
)
[Paraoxon], (µM)
80
The very low ΔH* value for the reaction catalyzed by salmon lipase is a
reflection of the relative insensitivity of this enzyme to temperature change,
and is consistent with the low Q10 value presented above. Hoki lipase-
catalyzed hydrolysis of p-NPC has an enthalpy of activation value very similar
to that for poikilothermic Greenland cod trypsin amidase reaction at 35°C
(Simpson and Haard 1984), whereas muscle aldolase from the endothermic
rabbit displayed an almost two-fold higher value for fructose 1,6-diphosphate
cleavage reaction at 31.9°C (Lehrer and Barker 1970). Chinook salmon and
hoki digestive lipases thus appear well adapted to operating in colder, variable
environments.
The salmon enzyme lowered slightly the free energy of activation (ΔG*) for
the hydrolysis reaction compared to the hoki. Since the contributions of ΔH*
and ΔS* to ΔG* are significantly different for the two lipases, it is very likely
that the internal structures and 3-D conformations of the enzymes are not
alike, and these results substantiate the findings from electrophoretic analyses,
reported above.
Fig. 3.12. Arrhenius plot for the hydrolysis of p-NPC by Chinook salmon
lipase. The temperature range was 20-30°C. Values are means ± SD (n = 3)
y = -1.021x + 0.2216
R² = 1
y = -1.2332x + 0.9894
y = -0.8088x - 0.5461
-3.35
-3.3
-3.25
-3.2
-3.15
-3.1
-3.05
3.28 3.3 3.32 3.34 3.36 3.38 3.4 3.42
ln a
ctiv
ity
1/T x 103 (K-1)
81
Fig. 3.13. Arrhenius plot for the hydrolysis of p-NPC by hoki lipase. The
temperature range was 20-35°C. Values are means ± SD (n = 3)
Table 3.3
Thermodynamic parameters for lipase-catalyzed hydrolysis of p-NPC at 35°C
Chinook salmon Hoki
Ea (kcal/mol) 2.0 ± 0.4 7.6 ± 0.8
ΔG* (kcal/mol) 17.3 ± 2.4 18.3 ± 1.5
ΔH* (kcal/mol) 1.4 ± 0.3 7.0 ± 0.8
ΔS* (eu, cal mol-1
K-1
) -51.4 ± 7.7 -36.7 ± 4.1
Values are means ± SD (n = 3); R2 value for the fit of data to Arrhenius
equation was 1 for salmon and 0.98 for hoki; eu, entropy unit.
3.4.12. Lipase activity with different bile salts
While CELs are activated by bile salts, the tri-hydroxylated are much more
potent activators than the di-hydroxylated types (Wang and Hartsuck 1993;
Hui and Howles 2002). For the salmon and hoki lipases, the highest activity
was seen with the two tri-hydroxylated bile salts cholate and taurocholate,
with taurocholate activating the salmon lipase slightly more than cholate (Fig.
3.14). No significant difference in activity was seen between cholate and
taurocholate (at both 5 and 10 mM) for the hoki lipase. The activity was not
increased in the presence of 1mM deoxycholate. This was the highest
concentration that could be used due to precipitation with Ca. Although the
level of deoxycholate was relatively low, it is unlikely that any activation
y = -3.8202x + 9.1225
R² = 0.9792
y = -3.407x + 8.0289
y = -4.2334x + 10.216
-4.4
-4.2
-4
-3.8
-3.6
-3.4
-3.2
-3
3.2 3.25 3.3 3.35 3.4 3.45
ln a
ctiv
ity
1/T x 103 (K-1)
82
would be achieved with higher concentrations. The effect of bile salts on the
activity of the two fish enzymes strongly suggests that they are CELs.
Fig. 3.14. Effect of bile salts on salmon lipase activity against p-NPC.
Activities are relative to 5mM cholate (standard assay). Values are means ±
SD (n = 3)
3.4.13. Activity/stability with selected chemicals
Metal ions, halide ions, and detergents generally have similar effects on all
lipases. The effects of several of these compounds, in addition to sodium azide
and colipase, on the salmon and hoki lipases are presented in Table 3.4.
Table 3.4
Effect of selected chemicals on the stability of Chinook salmon and hoki
digestive lipases
Chemical Incubation concentration Residual activity (%)
Chinook salmon Hoki
FeCl2 1 mM 52 ± 11 18 ± 2
10 mM 14 ± 17 7 ± 16
HgCl2 1 mM 4 ± 2 5 ± 6
10 mM 0 0
KI 1 mM 94 ± 2 96 ± 12
10 mM 98 ± 6 90 ± 14
0
20
40
60
80
100
120
140
Rel
ati
ve
act
ivit
y (
%)
Bile salt
83
NaN3 1 mM 75 ± 6 85 ± 15
10 mM 101 ± 14 78 ± 10
ZnCl2 1 mM 101 ± 3 85 ± 11
10 mM 51 ± 10 49 ± 5
MgCl2 1 mM 80 ± 7 86 ± 9
10 mM 96 ± 10 94 ± 17
MnCl2 1 mM 103 ± 5 100 ± 13
10 mM 143 ± 11 113 ± 11
PMSF 1 mM 66 ± 8 81 ± 9
5 mM 36 ± 6 54 ± 4
Tween 80 1% 67 ± 5 51 ± 8
10% 33 ± 4 20 ± 6
Triton X-100 1% 17 ± 2 27 ± 9
10% 0 0
Colipase 1 µM 93 ± 9 106 ± 11
10 µM 96 ± 8 113 ± 14
Values are means ± SD (n = 3); the residual activity (%) is relative to the
appropriate control (100%), as described in section 3.3.15.
Mercury ion was the strongest inhibitor, followed by Fe2+
. This outcome is
typical for these ions (Patkar and Bjorkling 1994; Anthonsen et al. 1995). Hg2+
is a thiol group inhibitor, suggesting the presence of cysteine residues and
disulfide bridges in the Chinook salmon and hoki digestive lipases. Hg2+
(1
mM) also inhibited completely grey mullet and P. aurantiogriseum lipases
(Lima et al. 2004; Aryee et al. 2007), whereas ferric and ferrous ions (1 mM)
inhibited oil sardine lipase (Mukundan et al. 1985).
Of all the metal chlorides, Mn2+
enhanced the activity of the fish lipases the
most. This was particularly evident with the salmon enzyme and 10 mM
concentration of the activator. The same was reported by Aryee et al. (2007)
and Lima et al. (2004). This result is important, as it may offer an
improvement over Ca2+
ions in an optimized lipase assay.
Mg2+
, at both 1 and 10 mM, and Zn2+
at 1 mM did not influence the lipases to
a great extent, as was the case in some previous studies (Mukundan et al.
1985; Lima et al. 2004; Aryee et al. 2007). However, at a level of 10 mM,
84
ZnCl2 caused an activity loss of ~ 50% for both salmon and hoki enzymes, and
the same was reported for R. chinensis lipase (Sun et al. 2009).
Iodide ion did not inhibit either lipase. Mukundan et al. (1985) observed a
small inactivation (11%) with this ion on sardine lipase activity.
PMSF, a commonly used serine protease inhibitor, caused a decrease in the
residual activity of both lipases, with the higher concentration of the inhibitor
having a greater impact. This result is not surprising, considering the presence
of a common catalytic triad in both serine proteases and lipases (Winkler and
Gubernator 1994). Correspondingly, PMSF (10 mM) completely inhibited
grey mullet lipase (Aryee et al. 2007), and the fluoride ion (in the form of 1
mM NaF) completely inhibited sardine lipase (Mukundan et al. 1985). Thus,
PMSF cannot be used to protect these lipases from protease hydrolysis during
purification. Benzamidine (2 mM) was used in the present study at all stages
of purification. Preliminary investigations with crude pyloric ceca extracts
showed that inclusion of ≤ 5 mM benzamidine did not have any adverse
effects on Chinook salmon and hoki lipase activity.
A commonly used antimicrobial preservative, NaN3 appeared to have a small
inhibitory effect on the activity of the two lipases. In contrast, grey mullet
lipase, P. aurantiogriseum lipase, and three other fungal lipases all had
slightly enhanced activities after exposure to this compound (Lima et al. 2004;
Aryee et al. 2007).
Triton X-100 and Tween 80 have similar hydrophilic/lipophilic equilibrium
values (13.5 and 15, Lima et al. (2004)), and they both inhibited the lipases to
varying degrees. Triton X-100 had greater incompatibility with the enzymes,
and the same was reported by Aryee et al. (2007) and Lima et al (2004). On
the other hand, these researchers did not find inhibitory effects with ≤ 1%
Tween 80 on grey mullet and P. aurantiogriseum lipases.
During incubation, colipase was present in molar excess relative to both
lipases (0.32 µM for salmon and 0.84 µM for hoki), even at the lower
concentration of 1 µM. Under such conditions, it did not significantly activate
85
either fish lipase indicating that Chinook salmon and hoki lipases are most
likely not pancreatic lipase-type enzymes.
3.4.14. Stability in organic solvents
The lipases were unstable in hydrophilic solvents methanol, isopropanol and
acetone at 50% concentration (Table 3.5). This is consistent with the findings
of several previous studies on fungal and fish lipases (Lima et al. 2004; Aryee
et al. 2007; Sun et al. 2009). The water-miscible solvents tend to strip off the
essential water molecules from the enzyme surface, resulting in unfolding and
denaturation of the enzyme (Zaks and Klibanov 1988). Even at 10%
incubation concentration, these solvents caused some loss in residual activity.
In contrast, the water-immiscible solvents did not have an apparent adverse
effect on the lipase stability, except for isooctane, where a reduction in activity
was observed after incubation. The apparent stability of the fish lipases in
lipophilic solvents could indicate potential for their use in applications where
low water content is desirable, such as synthesis reactions and lipid
modifications.
Table 3.5
Effect of organic solvents on the stability of Chinook salmon and hoki
digestive lipases
Solvent Incubation concentration
(%)
Residual activity (%)
Chinook salmon Hoki
Methanol 10 67 ± 8 58 ± 10
50 0 0
Isopropanol 10 64 ± 5 77 ± 8
50 23 ± 4 36 ± 9
Acetone 10 69 ± 5 66 ± 5
50 0 0
Hexane 100 82 ± 19 90 ± 16
n-Heptane 100 95 ± 21 103 ± 17
Isooctane 100 58 ± 21 71 ± 16
86
Values are means ± SD (n = 3); the residual activity (%) is relative to the
appropriate control (100%), as described in section 3.3.16.
3.4.15. Amino acid analysis
Table 3.6 compares the amino acid composition of Chinook salmon and hoki
lipases. There are no significant differences in composition between the
lipases in this study, mammalian CELs (van den Bosch et al. 1973; Lombardo
et al. 1978; Albro et al. 1985; Rudd et al. 1987; Abouakil et al. 1988) and
sardine lipase (Mukundan et al. 1985), except for the relatively low proline
content seen in the salmon and hoki lipases. This difference is related to the
existence of proline-rich repeats near the C-terminus, which are present in
many mammalian CELs, most notably human CEL (Lombardo et al. 1978;
Wang and Hartsuck 1993). The proline content of Chinook salmon and hoki
proteins suggests that these enzymes have very few or no proline-rich repeats.
The repeats are absent in the 60 kDa cod CEL (Gjellesvik et al. 1992), as well
as in < 60 kDa Atlantic salmon CEL (Gjellesvik et al. 1994). The absence of
C-terminal repeats is in agreement with the low molecular weights of salmon
and hoki lipases. The similarity in amino acid composition with the
mammalian suggests that Chinook salmon and hoki enzymes are of CEL type.
Table 3.6
Amino acid composition of Chinook salmon and hoki digestive lipases
Amino
acid
Chinook
salmon
Hoki
Lys
His
Arg
Asx
Thr
Ser
Glx
Pro
Gly
6.0
2.1
4.8
10.0
5.8
6.7
9.7
4.3
9.5
5.9
1.8
4.1
10.8
6.1
7.5
12.1
5.1
10.1
87
Ala
Cys
Val
Met
Ile
Leu
Tyr
Phe
Trp
8.0
nd
7.5
2.4
5.4
9.8
3.3
4.8
nd
8.7
nd
6.7
1.7
4.5
8.5
2.8
3.5
nd
The values represent molar percentages; nd = not determined.
N-terminal sequences of both lipases could not be determined in the present
study after repeated attempts, suggesting a blocked N-terminal in both
enzymes. Gjellesvik et al (1992) concluded the same after their attempts to
determine the N-terminal sequence of cod lipase failed.
3.5. Conclusion
Through an understanding of the properties of Chinook salmon and hoki
digestive lipases, there appears to be potential to apply these enzymes in real
systems. Considering the molecular weight, amino acid composition, lack of
any activation by colipase, lack of basal activity above C4 ester of p-NP, and
that the highest activity was achieved with tri-hydroxylated bile salts, it is
concluded that the Chinook salmon enzyme is highly homologous to
mammalian CELs. The same is most likely true for the hoki enzyme.
The apparent stability of the fish lipases in lipophilic solvents may offer
application advantages in anhydrous or low water content systems, such as
organic synthesis. The salmon enzyme significantly lowered the activation
energy for the hydrolysis of the synthetic substrate, displayed a relative
insensitivity to temperature change and relatively high activity at lower
temperatures, as evidenced by Q10 and ΔH* values. These results suggest that
the Chinook salmon lipase could find uses in low temperature applications.
Both lipases exhibited the highest activity against a medium chain (C10) ester
of p-NP. This property could be exploited for applications such as the
88
hydrolysis of milk lipids, and release of flavour volatiles desired in specialty
cheeses and other ‗ripened‘ dairy products.
In order to assess further potential applications, it is essential to determine the
fatty acid specificity and regiospecificity of these lipases by using natural
substrates (e.g. plant and animal oils). These investigations are the focus of our
current research.
3.6. Acknowledgements
This study was supported by funds from the New Zealand Foundation for
Research, Science & Technology (FRST) under contracts C02X0301 and
C02X0806. We thank The New Zealand King Salmon Co Ltd. for providing
salmon tissue samples and Sealord Ltd. for providing hoki tissue samples. This
research was facilitated using infrastructure provided by the Australian
Government through the National Collaborative Research Infrastructure
Strategy (NCRIS) and we thank Australian Proteome Analysis Facility,
Macquarie University for amino acid composition and N-terminal analyses.
89
CONNECTING STATEMENT 2
The chemical and catalytic properties of Chinook salmon and hoki digestive
lipases are described in the previous chapter. The lipases were classified as
carboxyl ester lipases (also known as bile salt activated lipases). Maximal
activity against a medium chain ester of p-NP indicated a potential for the
hydrolysis of lipids containing short and medium chain FAs. This led to the
design of the next study. In chapter IV, Chinook salmon and hoki bile salt
activated lipases were evaluated as flavour modifying agents in dairy cream.
The performance of the fish lipases was compared to that of two commercially
available lipases used in dairy product flavour development.
90
CHAPTER IV. FLAVOUR DEVELOPMENT IN DAIRY CREAM
USING FISH DIGESTIVE LIPASES FROM CHINOOK SALMON
(ONCORHYNCHUS TSHAWYTSCHA) AND NEW ZEALAND HOKI
(MACRURONUS NOVAEZELANDIAE) (MANUSCRIPT 3)
Published in Food Chemistry, 2011, 127:1562-1568
Authors: Ivan Kurtovic1,2
, Susan N. Marshall2*, Matthew R. Miller
2 and Xin
Zhao1
Affiliations:
1Department of Animal Science, McGill University (Macdonald Campus)
21,111 Lakeshore Road, Ste. Anne de Bellevue (QC) Canada H9X 3V9
2The New Zealand Institute for Plant and Food Research Limited, PO Box
5114, Port Nelson, Nelson, New Zealand
* Corresponding Author: Dr. Susan N. Marshall
The New Zealand Institute for Plant and Food Research Limited
PO Box 5114, Port Nelson, Nelson, New Zealand
Tel.: + 643 9897611, Fax: + 643 5467049,
E-mail address: [email protected]
4.1. Abstract
Digestive lipases from Chinook salmon and New Zealand hoki were evaluated
as flavour modifying agents in dairy products. Cream was either incubated
with fish lipase or commercially available lipases used in dairy flavour
development – calf pregastric esterase (Renco™ PGE) and microbial lipase
(Palatase® 20000 L). The fish enzymes were more similar to calf PGE in terms
of the total amount and types of fatty acids released over the course of the
reaction. Like the pregastric esterase, the fish enzymes released mainly short
chain fatty acids from cream triglycerides. The highest specificity was towards
the key dairy product flavour and odour compounds, butanoic and hexanoic
acids. The odour intensity of hexanoic acid produced by the salmon lipase, as
measured by SPME-GC-MS, was similar to that produced by both Palatase®
and PGE. Free fatty acid composition, together with sensory characteristics of
91
lipase-treated creams, demonstrated the potential for flavour enhancement in
dairy products using fish lipases.
Keywords: Chinook salmon; cream; hoki; lipase; short chain fatty acid
4.2. Introduction
Lipases are commonly used to generate desirable flavours in dairy products
because of their unique specificities towards fatty acids and controlled
hydrolysis of milkfat triglycerides. The volatile free fatty acids (FFAs)
generated by lipases provide characteristic flavours in dairy products such as
cheese and dairy flavour concentrates, and in addition impart organoleptic
properties of richness and creaminess (Balcão and Malcata 1998). Fatty acids
(FAs) like butanoic, hexanoic and octanoic acid are important contributors to
the desired ―buttery‖ flavours (Kinsella 1975), and thus the focus is on
enzymes which have high specificities towards these short chain FAs.
Pregastric esterase (PGE), also called pregastric lipase or lingual lipase, is
sourced from ruminants (calf, kid and lamb) and used for accelerated ripening
and development of characteristic flavours in several cheeses and other dairy
products (Birschbach 1994; Lai et al. 1998; Hernández et al. 2009). This is due
to the specificity for short chain FA of all the ruminant PGE enzymes
(O'Connor et al. 2001). In addition, each type of PGE gives rise to its own
characteristic flavour due to slight differences in FA specificity. Pregastric
esterases are sn-1,3-specific (Ha and Lindsay 1993; Birschbach 1994). Ha and
Lindsay (1993) showed that all three pregastric lipases preferentially
hydrolyzed volatile branched-chain and short n-chain FAs from ruminant
milkfat. Commercially, the enzyme has been used in the manufacture of
piquant Italian cheeses including Provolone, Romano and Parmesan (Harper
1957; Woo and Lindsay 1984), and other Mediterranean cheeses, like
Idiazabal (Hernandez et al. 2001).
Microbial lipases have also been used commercially for the development of
desired flavours in cheese (Birschbach 1994). Palatase® 20000 L is a purified,
sn-1,3-specific lipase, derived from the fungus Mucor miehei, and developed
92
for the production of cheese flavours or enzyme-modified cheeses. It is
recommended for use in Italian cheese-making (Novozymes 2010). The
enzyme has also been evaluated for suitability in the manufacture of several
types of Spanish cheeses, including Manchego (Fernandez-Garcia et al. 1994)
and Idiazabal (Hernandez et al. 2001). However, it failed to produce the
characteristic flavours of these cheeses due to excessive release of long chain
FAs.
Lipases from the marine environment have the potential to expand the
available activities and specificities. Fish are a prospective source of lipases,
with the digestive organs of these animals being an abundant and underutilized
by-product of fish processing. Because of the environment in which different
fish live (e.g. at low temperatures) and in response to their diets, the lipases
from some fish have higher catalytic efficiencies at lower temperatures than
either their microbial or mammalian counterparts (Kurtovic et al. 2009a).
There may be an opportunity to use fish lipases in commercial dairy
applications, especially if they can deliver particular and desirable flavours not
provided by the enzymes in use now.
We have previously purified and characterized the digestive lipases from
Chinook salmon, Oncorhynchus tshawytscha, and New Zealand hoki,
Macruronus novaezealandiae (Kurtovic et al. 2010). These enzymes are
carboxyl ester lipases (often called bile salt activated lipases) and require
calcium ions and sodium cholate for optimum activity. Nevertheless, both
lipases can operate at a lower rate without bile salts, so these could be omitted
from a food product application, if necessary. Both lipases exhibit the highest
activity against a medium chain (C10:0) ester of p-nitrophenol. The Chinook
salmon lipase, in particular, exhibits relatively high activity at lower
temperatures compared to other fish and mammalian carboxyl ester lipases
(Kurtovic et al. 2010). These properties could make the enzymes suitable for
applications such as the hydrolysis of milkfat, i.e. release of FAs and flavour
volatiles desired in specialty cheeses and other ‗ripened‘ dairy products.
93
This manuscript describes the release of short chain FAs and development of
other flavour compounds through the hydrolysis of cream lipids by Chinook
salmon and hoki digestive lipases. For comparison, calf PGE and Palatase®
20000 L are included in the study.
4.3. Materials and methods
4.3.1. Substrate and other chemicals
Fresh cream (pasteurised and non-homogenised, 35% fat; Meadow Fresh,
Manukau, New Zealand) was purchased from a local supermarket. All samples
used in the study were from the same production batch. Authentic Parmesan
cheese (Parmigiano Reggiano from Virgilio, Montova, Italy) was also
purchased from a local supermarket. All solvents and other reagents were of
chromatography or analytical grade.
4.3.2. Enzymes
Partially purified Chinook salmon and hoki digestive lipases were prepared
from lyophilised and delipidated pyloric ceca powder by extraction with 100
mM phosphate buffer (pH 7.8, containing 2 mM benzamidine, 1 mM EDTA
and 20% (w/v) glycerol), precipitation with polyethylene glycol 1000, and
dialysis of the resulting supernatant in the named buffer with 12–14 kDa
molecular weight cut off tubing (Medicell International Ltd., London, UK).
The salmon and hoki lipases had activities against p-nitrophenyl palmitate of
1.11 and 0.24 U/mL (0.37 and 0.11 U/mg protein), respectively (Kurtovic et
al. 2010).
PGE from calf epiglottis (80 lipase forestomach units/g, Committee on Codex
Specifications 1981) was supplied by Renco New Zealand (Eltham, New
Zealand). The preparation contained both esterase and lipase activity. The
powder (0.5 g) was prepared in 5 mL Milli-Q water for use in the experiments.
Palatase® 20000 L liquid preparation (20000 lipase units/g) was purchased
from Novozymes A/S (Bagsvaerd, Denmark) and used in its original form.
94
4.3.3. Standardized lipase assay using cream
Lipase activity against cream was measured titrimetrically for all of the
enzymes with a modified method based on that for measuring lipase activity
against tributyrin (Committee on Codex Specifications 1981). Reactions were
carried out at 25 °C with 75 mL cream and sample volumes ranging from 10
µL (Palatase®
) to 1 mL (the fish lipases), in a stirred pH stat vessel (718 STAT
Titrino, Metrohm, Herisau, Switzerland). Calcium chloride (2 mM) and
sodium cholate (5 mM) were included in assays with the fish lipases. Each
reaction was monitored for 30 min. The released FFAs were titrated with 0.01
M NaOH with the pH stat set at 6.75 (pH of the fresh cream was 6.71–6.73).
The slope (mL 0.01 M NaOH/min) in the linear region of the titration curve
(typically the initial rate of reaction) was used to calculate the activity. Blank-
rate activity was determined in the absence of any enzyme sample. One unit
(U) of activity was defined as 1 µmol FFA released/min under the assay
conditions. All the assays were carried out in duplicate.
4.3.4. Hydrolysis of cream lipids
Hydrolysis reactions were carried out by adding the enzyme to pre-warmed
cream (100 mL) and maintaining the mixture at 25 °C for 24 h using a water
bath. Calcium chloride (2 mM) and sodium cholate (5 mM) were included in
the reaction mixture for fish lipases. The mixture was mixed manually with a
metal spatula every 30 min for the first 3 h, then every 60 min for the next 11
h before leaving it unstirred for the remainder of the time. The control reaction
omitted any enzyme. All reactions were carried out in triplicate. The values of
pH and degree of lipid hydrolysis (FFA% – g oleic acid/100 g lipid) were
measured at 0, 1, 2, 5, 11 and 24 h. FFA% was determined titrimetrically with
0.1 M NaOH (AOCS 1993).
4.3.5. Sensory analysis
Sensory analysis of the lipase-treated creams was carried out by three
laboratory staff who are used to sensory analysis of lipid products but who are
not a trained panel.
95
4.3.6. SPME-GC-MS analysis
Volatile organic compounds in the lipase-treated creams were analysed by
solid phase micro extraction (SPME)-GC-MS, using a modified method based
on that of van Aardt et al. (2005). Each cream sample was analysed in
triplicate, thus providing up to 9 sets of data for each lipase treatment. A
sample of cream (2.0 g) was taken at the end of the hydrolysis (24 h), and
placed in a 20 mL glass vial fitted with a self-sealing septum. Finely
comminuted Parmesan cheese (2.0 g) was also analysed for comparison. The
vial was incubated at 45 °C for 22 min with agitation, prior to volatiles
adsorption for 5 min using a 85 µm carboxen poly(dimethyl siloxane)-coated
SPME fibre (Supelco, Bellefonte, PA, USA). The adsorbed volatiles were then
analysed by GC-MS (GC-2010 gas chromatograph combined with GCMS-
QP2010 mass-spectroscopy unit and equipped with AOC-5000 automatic
injector, Shimadzu, Kyoto, Japan) under the following conditions: injection
temperature 280 °C; splitless mode; 1 min desorption time; Rtx-5Sil MS
capillary column – 30 m x 0.25 mm i.d. x 0.25 µm film thickness (Restek,
Bellefonte, PA, USA); oven temperature programme: 35 °C start, 15 °C/min to
180 °C, 20 °C/min to 260 °C, hold at 260 °C for 0.5 min; He carrier gas flow
rate 1.0 mL/min; detector temperature 260 °C. The peaks were identified by
comparison with the mass spectral database (Wiley7.0).
4.3.7. Lipid extraction and separation of free fatty acids
Lipid extraction and separation of FFAs were carried out according to the
method of de Jong and Badings (1990), with modifications to the volumes
employed during the procedures. A sample of cream (1.5 g) was taken at the
end of the hydrolysis (24 h) and combined with 1.5 mL ethanol, 0.15 mL 2.5
M sulphuric acid and 0.5 mL of FA recovery markers solution in a 15 mL
glass centrifuge tube. The FA recovery markers were prepared in ethanol (0.5
mg/mL each) and consisted of C5:0, C7:0, C9:0, C11:0, C13:0 and C15:0 (Nu-
Chek Prep, Elysian, MN, USA). Lipid was extracted with 2.25 mL
ether/heptane (1:1, v/v). The mixture was then centrifuged at 1500g for 5 min
at 20 °C and the upper solvent layer was transferred to a 10 mL test tube
containing 0.25 g anhydrous sodium sulphate to remove residual water. The
96
extraction was repeated twice with 1 mL ether/heptane and all the extracts
were combined.
Solid phase extraction with 500 mg/3 mL GracePure aminopropyl columns
(Grace, Columbia, MD, USA) was used to separate the FFAs. Each column
was first conditioned with 10 mL heptane and the entire lipid extract (~4.3
mL) was then applied onto the column. The neutral lipids were eluted first
with 10 mL chloroform/2-propanol (2:1, v/v), followed by elution of the FFAs
with 5 mL diethyl ether containing 2% formic acid.
4.3.8. Gas chromatography
A sample (2 µL) of the FFA extract was injected onto an Econo-Cap EC-1000
capillary column, 30 m x 0.32 mm i.d. x 0.25 µm acid modified polyethylene
glycol film (Grace, Columbia, MD, USA) and analysed by the previously
mentioned GC-MS in splitless mode. The injection temperature was 230 °C
and He carrier gas flow rate was 2.0 mL/min. During analysis the oven
temperature was raised from 40 to 225 °C at a rate of 10 °C/min, then held at
225 °C for 12 min. The detector temperature was 230 °C. A mass selective
detection method was used. The m/z 60 ion was used in the selected ion
monitoring (SIM) mode as quantifier for all fatty acids.
For the external calibration standard, nonanoic acid (C9:0) was prepared in
diethyl ether containing 2% formic acid. A total of 9 points, each measured in
duplicate, was used for the calibration curve (0.0015–1.5 mg/mL).
4.3.9. Statistical analysis
SPSS statistical software (v. 17.0, SPSS, Chicago, IL, USA) was used. One-
way analysis of variance (ANOVA) was conducted to establish the presence or
absence of significant differences in (i) volatile organic compound, and (ii)
FFA compositions of the creams treated with different lipases.
4.4. Results and discussion
4.4.1. Lipase assay and properties of the enzymes
The activity against cream for each lipase was determined in order to
standardize the activity parameter as different methods and substrates had
97
been used for routine activity measurements and to determine the
characteristics of the enzymes (titrimetric assay against tributyrin and olive oil
for the calf PGE and Palatase® respectively; spectrophotometric assay against
p-nitrophenyl palmitate for salmon and hoki lipases). The activities against
cream and several general characteristics of the lipases are presented in Table
4.1.
Table 4.1
Characteristics of the four lipase preparations
Enzyme Activity a
against cream
Total units used
in experiment
pH
optimum
pH working
range
Temperature
optimum (°C)
Chinook salmon lipase 3.2 U/mL 16.0 8.0–9.5 b 6.0–10.0 b 35–40 b
Hoki lipase 2.0 U/mL 10.0 8.0–9.0 b 6.0–10.0 b 35 b
Palatase® 20000 L 250 U/mL 17.5 6.5–7.5 c 5.0–9.5 c 45–65 c
Calf PGE 31 U/g 15.5 6.5–7.0 d 4.0–8.0 d 35–40 d
a 1 U = 1 µmol FA released/min.
b (Kurtovic et al. 2010).
c (Moskowitz et al. 1977; Uvarani et al. 1998).
d (O'Connor et al. 1993).
An assay temperature of 25 °C was chosen for the actual hydrolysis reactions
since the salmon and hoki lipases are unstable above 30 °C (for 30 min or
more), although their temperature optimum is around 35 °C (Kurtovic et al.
2010). Thus, 25 °C represents a compromise between a relatively high rate of
activity and stability. For comparative purposes, PGE and Palatase® were also
tested at 25 °C.
4.4.2. Reaction progress curves for cream lipid hydrolysis
Of the four lipases, Palatase® took the longest time to reach the stationery
phase of hydrolysis (15–24 h vs. 10–15 h for PGE and salmon lipase and ~10
h for hoki lipase). Palatase® also produced the most FFAs (23.5 ± 1.0 g oleic
acid/100 g lipid), and had the largest drop in pH (Fig. 4.1). The results for
Palatase®
-treated cream suggest that this enzyme has activity against many
different FAs, i.e. that it lacks FA specificity. In contrast, calf PGE is reported
to have high specificity towards short chain FAs (Ha and Lindsay 1993;
98
O'Connor et al. 2001) and this appears to be supported by the relatively low
FFA% after 24 h (5.4 ± 0.3 g oleic acid/100 g lipid). The salmon lipase
showed a very similar FFA% curve to that of PGE, although the pH did not
drop as much. The pH at the end of the salmon lipase hydrolysis reaction was
0.57 units higher than the corresponding pH for the PGE hydrolysis reaction,
probably due to the differences in the type and corresponding amount of
released FFAs. The cream lipolysed with hoki lipase showed the smallest
change in pH and FFA% at the end of the reaction.
Several trials were done to establish the required enzyme amounts, such that
the stationary phase of reaction could be reached by 24 h and which provided
similar number of activity units. Hoki lipase-treated cream took the least time
to reach the stationary phase (≤10 h). When more than 10 U of this lipase was
used, the reaction velocity was even higher, although the total amount of FFAs
produced was still the same. Thus, the final enzyme volumes provided
adequate reaction velocity and comparable total activity units.
For the salmon and hoki lipase-treated creams, the starting pH (6.72) was
below the optimum for these fish enzymes. However, the long reaction time
allowed hydrolysis at sub-optimal pH values. The pH after 24 h incubation
(6.01 and 6.30 respectively) was at the lower limit of these enzymes‘ pH
working range (Table 4.1), and may have inhibited the enzymes from further
hydrolysis.
a) b)
c) d)
0
5
10
15
20
25
30
5
5.4
5.8
6.2
6.6
7
0 5 10 15 20 25
g o
leic
acid
/10
0 g
lip
id
pH
Time (h)
0
1
2
3
4
5
6
7
5
5.4
5.8
6.2
6.6
7
0 5 10 15 20 25
g o
leic
acid
/10
0 g
lip
id
pH
Time (h)
99
Fig. 4.1. Changes in pH (—) and FFA% (– – –) for cream when treated with
four different lipase preparations: a) Palatase®; b) calf PGE; c) salmon; and d)
hoki, as described in section 4.3.4. The values are mean ± SD (n = 3). The
values after t = 0 h were adjusted using the control cream data (fresh cream
had a pH of 6.71–6.73 and FFA% of 1.5–2.0, and after 24 h incubation time
the pH had dropped to 6.62–6.67 and FFA% increased to 2.2–2.5).
4.4.3. Analysis of volatile compounds by SPME-GC-MS
The odour compounds identified in the lipolysed creams (those accounting for
more than 0.5% of the total volatiles peak area) are summarized in Table 4.2.
Table 4.2
Volatile organic compounds in lipase-treated creams and Parmesan cheese
Rt Compound Odour
description f
Cream sample Parmesan P
Control Palatase® Calf PGE Salmon Hoki
0.96 Ethanol Sweet 1.2 ± 0.6 a,b – a – a – a – a 2.9 ± 0.1 b **
1.00 Acetone Sweet, perfume 29.2 ± 5.1 c 2.5 ± 0.8 a,b 0.8 ± 0.3 a 3.2 ± 0.6 a,b 16.2 ± 3.0 b,c 3.7 ± 0.1 a,b **
1.11 2-Methylpropanal Dark chocolate – a – a – a – a – a 1.1 ± 0.1 b **
1.20 Butanone Ether, fruity 43.2 ± 3.8 b 2.3 ± 0.9 a 0.6 ± 0.2 a 2.3 ± 0.5 a 10.1 ± 2.7 a 0.8 ± 0.1 a **
1.24 Ethyl acetate Pineapple,
fruity
– a – a – a – a – a 1.1 ± 0.1 b **
1.42 3-Methylbutanal Chocolate – a – a – a – a – a 1.9 ± 0.3 b **
1.55 2-Pentanone Ether, fruity – a – a – a 1.3 ± 0.2 c 0.6 ± 0.2 b – a **
1.57 Acetic acid Vinegar 1.3 ± 0.8 a – a – a – a – a 21.9 ± 0.4 b **
1.62 Pentanal Sour, cut grass 8.9 ± 2.1 b – a – a – a 0.7 ± 0.2 a – a **
1.84 Acetoin Buttery, creamy 0.7 ± 0.3 a – a – a – a – a – a ns
2.08 Toluene Paint 0.6 ± 0.1 b – a – a – a – a 0.9 ± 0.1 c **
2.32 Ethyl butanoate Fruity, sweet – a – a – a – a – a 3.1 ± 0.2 b **
3.19 Butanoic acid
(+ 2-heptanone) g
Sharp, cheesy,
rancid, sweaty
0.6 ± 0.4 a 62.2 ± 2.1 c 75.7 ± 1.0 d 42.2 ± 1.4 b 46.4 ± 3.0 b 48.1 ± 0.7 b **
3.61 Pentanoic acid Sweaty, Swiss
cheese
– a – a 0.5 ± 0.1 b – a – a – a
**
4.28 Ethyl hexanoate Fruity, winy, – a – a – a – a – a 0.8 ± 0.1 b **
0
1
2
3
4
5
6
7
5.8
6
6.2
6.4
6.6
6.8
7
0 5 10 15 20 25
g o
leic
acid
/10
0 g
lip
id
pH
Time (h)
0
1
2
3
4
5
6
7
6
6.2
6.4
6.6
6.8
7
0 5 10 15 20 25
g o
leic
acid
/10
0 g
lip
id
pH
Time (h)
100
apple peel
4.88 Hexanoic acid Pungent,
musty, cheesy,
acrid
– a 29.3 ± 2.0 d 21.3 ± 1.0 c,d 48.0 ± 2.1 e 18.4 ± 3.6 b,c 11.4 ± 0.4 b
**
5.85 2-Ethyl hexanoic
acid
Herbaceous,
sweet, musty
– a 0.7 ± 0.4 a – a 0.6 ± 0.4 a 1.0 ± 0.3 a – a *
6.43 Octanoic acid Sweaty, cheesy – a 1.1 ± 0.4 b – a 0.5 ± 0.1 a,b – a – a **
Others 14.4 ± 3.1 b 1.9 ± 0.4 a 0.9 ± 0.2 a 1.9 ± 0.5 a 6.5 ± 1.5 a,b 2.2 ± 0.5 a **
The data (peak area %) represent mean values ± SE (n = 3 for Parmesan; 6 for
Palatase®, salmon and hoki; 7 for PGE; and 9 for the control cream); ns: not
significant (p>0.05); * p<0.05;
** p<0.01. For each compound, values without a
common letter are significantly different (Tukey‘s test, p<0.05). Compounds
less than 0.5% are grouped as ―others‖ or indicated by –.
f From (Peterson and Reineccius 2003; van Aardt et al. 2005; Lozano et al.
2007; d'Acampora Zellner et al. 2008; Mallia et al. 2008; Acree and Arn
2010).
g Coelution of peaks in Parmesan, Palatase
® and salmon samples.
A coelution of butanoic acid and 2-heptanone was present at a Rt ~3.2 in
Parmesan, Palatase®
and salmon profiles. While this peak could not be
resolved accurately, the mass spectrum data showed that 2-heptanone had a
minor contribution in all three profiles. In contrast, in some previous analyses
of the volatile components of Parmesan, 2-heptanone had a considerable
contribution (≤26% of the total volatiles peak area, Bellesia et al. 2003; Lee et
al. 2003), although its contribution varied greatly among the different samples
analysed. The discrepancy regarding 2-heptanone between our and these
previous studies is likely due to the source of the Parmesan. The 2-heptanone
peak could be clearly resolved from the butanoic acid peak in the hoki profile,
where it contributed slightly less than 0.5% of the total volatiles peak area so
is reported in the ―others‖ category in Table 4.2. In the control cream and PGE
profiles this compound was not detected.
Low molecular weight ketones (acetone and butanone) dominated the control
cream volatiles. These compounds have previously been reported as the major
volatile compounds in fresh cream (Wong 1963), and are responsible in part
for the sweet odour of cream. Small amounts of FFAs (acetic and butanoic
101
acid) were present, most likely due to a combination of the effects of heat and
light exposure and microbial contamination, since these compounds were not
detected at significant levels in the fresh cream (data omitted for brevity).
Pentanal was also present at a much lower level (<2%) in the fresh cream. The
increased levels of pentanal in the control cream are likely due to the light-
induced lipid oxidation during the 24 h incubation period (van Aardt et al.
2005). A large variety of minor compounds (―others‖) was present in the
control cream.
In contrast to the control cream, short chain FAs (butanoic and hexanoic acid)
were the dominant volatiles in all of the lipolysed creams. These FAs are
responsible for the cheesy (sharp, acrid and piquant) odours (d'Acampora
Zellner et al. 2008). Parmesan, due to the aging process, displayed a more
complex odour profile, including several low molecular weight esters and
aldehydes, which contribute fruity odours to this mature product. Another
distinctive characteristic of Parmesan is the relatively high level of acetic acid,
which was a minor volatile compound in the lipolysed creams. Both salmon
and hoki lipases released butanoic and hexanoic acids as the dominant
volatiles, with the salmon enzyme showing a significantly higher specificity
towards hexanoic acid. These two fatty acids provide a significant contribution
to the odour and flavour of many dairy products (Kinsella 1975; d'Acampora
Zellner et al. 2008). The combined contribution of these two fatty acids to the
total volatiles peak area was lower in the hoki than in the salmon lipase-treated
cream. This is in agreement with the sensory characteristics of the lipolysed
creams as the salmon lipase-treated cream was very similar to both Palatase®
and PGE-treated creams, whereas the hoki lipase-treated cream only shared
some of the sensory characteristics of the creams treated with the commercial
lipases (Table 4.3). The SPME-GC-MS analysis results indicate a potential use
of the fish lipases for the generation of desirable flavours in dairy products,
which would be different to the flavour profiles produced by Palatase® and
PGE.
Table 4.3
Sensory characteristics of the creams treated with different lipase preparations
102
Cream sample Odour Consistency
Control Sweet, slightly buttery and fatty Very smooth, fluid
Palatase® Strong sharp, acrid and piquant Very thick
Calf PGE Cheesy, rancid, sharp
(characteristic of Italian cheeses, e.g. Parmesan)
Quite smooth, fluid
Salmon Sharp, cheesy, piquant (resembling Parmesan) Thick
Hoki Sweet, buttery with some acrid, cheesy notes Smooth, fluid
The only volatile branched-chain FA detected in this study, 2-ethyl hexanoic
acid, could have been a contaminant from packaging, since it is used as a
stabilizer in plastics (Elss et al. 2004). The only FA with an odd carbon
number was pentanoic acid which was detected in significantly higher levels
in the PGE sample (~0.5% of the total volatiles peak area). This is most likely
due to this enzyme‘s high specificity towards all short chain FAs.
Since the SPME-GC-MS technique is in principle quantitative, the actual peak
areas for the most dominant FAs (butanoic and hexanoic acid) were compared
amongst the four lipase-treated creams (Fig. 4.2).
Fig. 4.2. Peak area intensity for butanoic (C4:0) and hexanoic (C6:0) acids
after SPME-GC-MS analysis of creams treated with different lipases. The
values are mean ± SE (n = 6 for Palatase®, salmon and hoki; and 7 for PGE).
0
20
40
60
80
100
120
140
Butanoic acid * Hexanoic acid
Pea
k a
rea
in
ten
sity
(%
)
PGE
Palatase
Salmon
Hoki
b
a
a
aa
a,b
bb
103
Overall significance for each FA: p<0.01. For each FA, values without a
common letter are significantly different (Tukey‘s test, p<0.05).
* includes 2-heptanone in Palatase® and salmon samples.
The highest amount of butanoic acid was produced by PGE. On the other
hand, the amount of hexanoic acid produced by PGE, Palatase® and the
salmon lipase was not significantly different, indicating that the salmon lipase
could be used to successfully produce flavour intensities similar to those
produced by the commercial lipases.
4.4.4. Free fatty acid composition
In agreement with the FFA% result from the reaction progress curve (Fig.
4.1a), Palatase® produced the greatest change in FFA composition (Fig. 4.3a).
Specificity towards short chain FAs has been attributed to both calf PGE (Ha
and Lindsay 1993; O'Connor et al. 2001) and Palatase®. According to
Novozymes (2010), the preference of Palatase®
for hydrolyzing small FAs
results in optimal flavour formation. However, our results for Palatase®-
lipolysed cream showed a preference towards primarily medium chain FAs,
followed by short chain FAs. Similarly, Hernandez et al. (2001) found that
Palatase®
20000 L did not produce enough short chain FAs required for the
typical piquant flavour of Idiazabal cheese. In our study, specificity towards
short chain FAs was confirmed for the calf PGE.
The pattern of change in FFA composition of both salmon and hoki lipase-
treated creams is similar to that of PGE-treated cream. Both salmon and hoki
lipases released predominantly short chain FAs. Furthermore, the differences
in FFA composition between PGE and salmon lipase-treated creams were not
significant (Fig. 4.3a). The changes in the individual short chain FAs were also
compared (Fig. 4.3b), since they have the greatest impact on the flavour of a
dairy product.
104
a)
b)
Fig. 4.3. a) Changes in FFA composition of cream after treatment with
different lipases. The results are expressed as changes in peak area % as
compared to control cream (n = 3). Overall significance for each FFA
grouping: p<0.01. For each FFA grouping, values without a common letter are
significantly different (Tukey‘s test, p<0.05).
b) Changes in butanoic, hexanoic and octanoic acid concentrations in
cream after treatment with different lipases. The results are expressed as
-50
-40
-30
-20
-10
0
10
20
30
40
50
Palatase PGE Salmon Hoki
Ch
an
ge
in p
eak
are
a %
C16:0, 18:0, 18:1
C10:0-14:0
C4:0-8:0
a,ba b,cc
aa
ab
a a,b
b,c
c
0
5
10
15
20
25
Palatase PGE Salmon Hoki
Incr
ea
se i
n p
eak
are
a %
C4:0
C6:0
C8:0b
a
a
a
a a
a,b
b
a
b
a,ba,b
105
changes in peak area % as compared to control cream. The values are mean ±
SE (n = 3). Overall significance for each FA: p<0.05. For each FA, values
without a common letter are significantly different (Tukey‘s test, p<0.05).
Both salmon and hoki lipases produced major increases in butanoic and
hexanoic acids, with the hoki lipase profile resembling that of PGE. The
salmon lipase produced hexanoic acid in the highest amounts, followed by
butanoic acid. Thus, a tailored flavour profile could be obtained in cheeses
made with these fish lipases through the selection of one or the other. The FFA
composition with respect to specificity towards short chain FAs is in overall
agreement with the analysis of volatile compounds by SPME-GC-MS (e.g.
butanoic acid is both the dominant volatile and the most abundant FFA in PGE
and hoki lipase-treated cream, and the same is true for hexanoic acid in the
salmon lipase-treated cream).
It has been demonstrated that in milkfat triglycerides the two short chain FAs
(butanoic and hexanoic acids) are esterified almost exclusively on the sn-3
position, and the medium chain FAs (C10:0 to C14:0) are mainly on the sn-2
position (Parodi 1982). Consequently, the salmon and hoki lipases, like the
calf PGE (O'Connor et al. 2001), appear to be sn-1,3-specific lipases against
milkfat triglycerides. Interestingly, since it released both short and medium
chain FAs, Palatase® appears to have no preference towards either external
(sn-1,3) or internal (sn-2) FAs, despite being described as a sn-1,3-specific
lipase (Novozymes 2010).
The high reactivity of both salmon and hoki lipases towards short chain FAs is
attractive from an applications standpoint. If used in commercial dairy
products, unique flavour profiles for salmon and hoki lipase-modified dairy
lipids would be expected due to differences in the short chain FA specificity
between the fish, mammalian and microbial lipases used in this study. To
make commercial applications successful, there are barriers to overcome.
Despite the raw material (pyloric ceca) being both cheap and abundant, there
are significant costs associated with developing large-scale, food-grade
extraction processes. There may also be issues with consumer acceptance.
106
4.5. Conclusion
Based on the sensory characteristics, FFA composition, and the analysis of
volatile organic compounds present in the lipolysed creams, both Chinook
salmon and hoki digestive lipases showed considerable potential for flavour
enhancement in dairy products. The enzymes were able to release the short
chain FAs from cream triglycerides, with the highest specificity towards the
key flavour and odour compounds, butanoic and hexanoic acids. Differences
in the short chain FA specificity of the two fish lipases could be used to
generate distinctive flavour profiles in dairy products.
4.6. Acknowledgements
This study was supported by funds from the New Zealand Foundation for
Research, Science & Technology (FRST) under contracts C02X0301 and
C02X0806. We thank The New Zealand King Salmon Co. Ltd for providing
salmon tissue samples, Sealord Group Ltd for providing hoki tissue samples,
and Renco New Zealand for providing calf pregastric esterase.
107
CONNECTING STATEMENT 3
The FA specificities established for the two fish lipases, described in the
previous chapter, demonstrated a potential application in dairy products. In
chapter V, the immobilization of Chinook salmon bile salt activated lipase is
described using two hydrophobic supports. The salmon lipase was selected for
immobilization as it had higher specific activity against p-NP esters (chapter
III) and was more effective at hydrolyzing cream (chapter IV) than hoki lipase.
108
CHAPTER V. HYDROPHOBIC IMMOBILIZATION OF A BILE SALT
ACTIVATED LIPASE FROM CHINOOK SALMON
(ONCORHYNCHUS TSHAWYTSCHA) (MANUSCRIPT 4)
Journal of Molecular Catalysis B: Enzymatic, 2011, under revision
Authors: Ivan Kurtovica,b
, Susan N. Marshallb* and Xin Zhao
a
Affiliations:
aDepartment of Animal Science, McGill University (Macdonald Campus)
21,111 Lakeshore Road, Ste. Anne de Bellevue (QC) Canada H9X 3V9
bThe New Zealand Institute for Plant & Food Research Limited, PO Box 5114,
Port Nelson, Nelson, New Zealand
[email protected]; [email protected]
* Corresponding Author: Dr. Susan N. Marshall
The New Zealand Institute for Plant & Food Research Limited
PO Box 5114, Port Nelson, Nelson, New Zealand
Tel.: + 643 9897611, Fax: + 643 5467049;
E-mail address: [email protected]
5.1. Abstract
Immobilization of a bile salt activated lipase from Chinook salmon was
achieved on two hydrophobic supports. Salmon lipase immobilized on octyl-
Sepharose had approximately 40-fold higher activity (on a dry weight basis)
against a tributyrin emulsion than the same lipase immobilized on Lewatit VP
OC 1600. It also had approximately 10-fold higher activity than Candida
antarctica Lipase B immobilized on Lewatit (Novozym 435). Salmon lipase-
octyl-Sepharose was highly active against both ghee and fish oil emulsions,
but Novozym 435 and salmon lipase-Lewatit had very low activities against
the fish oil emulsion. The vast discrepancies in lipolytic activity between the
two forms of immobilized fish lipase, and Novozym 435, are attributed in part
to the conformation of the bound lipases, but most importantly to the different
hydrophobicities of the lipase-carrier complexes, which determine the
substrate access for hydrolysis. Immobilization appeared to raise the
temperature optimum of the salmon lipase against tributyrin. The thermal
109
stability and bile salt requirement of the lipase were not affected by
immobilization. The results demonstrate a successful immobilization of an
active and functional fish bile salt activated lipase and its potential for low
temperature modifications of lipids in emulsions.
Keywords: bile salt activated lipase; Chinook salmon; immobilization;
Lewatit VP OC 1600; octyl-Sepharose
5.2. Introduction
Enzymes are versatile, efficient and precise biocatalysts. The soluble lipases
are easy to apply and highly useful in applications such as flavour
development in food products and detergents. However, for applications on an
industrial scale such as organic synthesis and modifications of fats and oils, it
is necessary to immobilize the enzymes. Immobilization separates the enzyme
from the reaction products, allows for multiple re-use and continuous
processing. Immobilization often improves the activity and stability
parameters of an enzyme, as well as altering its substrate specificity and
enantioselectivity (Malcata et al. 1990; Fernandez-Lafuente et al. 1998; Holm
and Cowan 2008; Hanefeld et al. 2009).
Supports with densely-packed, strongly hydrophobic, lipid-like groups (e.g.
octyl-agarose) have been developed to allow preparation of immobilized lipase
derivatives that are efficient catalysts in high water-activity systems
(Fernandez-Lafuente et al. 1998). Lipase attachment to the hydrophobic
surface of the support is assumed to involve an extremely hydrophobic area
around the active site, with the enzyme undergoing interfacial activation
during immobilization. Interfacial activation is thought to involve an
interaction between the lipase and substrate at a lipid/water interface causing a
change in enzyme conformation which enhances activity (Derewenda et al.
1992). Immobilization on a lipid-like hydrophobic support such as octyl-
agarose produces the same effect. This adsorption is highly selective and
provides a means to immobilize and purify at the same time (Fernandez-
Lafuente et al. 1998), which is particularly useful for unstable enzymes such
as Chinook salmon lipase.
110
Microbial lipases are of the greatest interest for biocatalysis. Commercial
microbial lipases are produced extracellularly in high yield from cultures,
helping to make recovery convenient and relatively economical. In addition,
the microbial lipases can be genetically manipulated more easily than those
from animals and tend not to require additional factors such as calcium and
bile salts for activity (as indicated in the review by Kurtovic et al. 2009b).
Candida antarctica Lipase B (CAL-B) is one of the most versatile and
commonly used lipases in biotransformations (Chen et al. 2008; Cabrera et al.
2009). Novozym 435 (Novo 435, produced by Novozymes) is the immobilized
form of CAL-B on Lewatit VP OC 1600, a macroporous acrylic polymer
resin. The full immobilization protocol has not been disclosed, but the carrier
is known to be of a hydrophobic nature (poly(methyl methacrylate crosslinked
with divinylbenzene), (Chen et al. 2008) and the enzyme bound to it via
hydrophobic interactions. This has been demonstrated by incubating the resin
with detergents and organic solvents to successfully desorb CAL-B (Chen et
al. 2008; Cabrera et al. 2009). Novo 435 will operate in aqueous media (i.e.
emulsions), but it is most valued for its high activity in non-aqueous media
such as oils and organic solvents (Robles Medina et al. 1999; Deng et al. 2005;
Holm and Cowan 2008; Adamczak et al. 2009; Karabulut et al. 2009),
suggesting that the lipase-carrier complex also has hydrophobic character.
We have previously purified and characterized the digestive lipase from
Chinook salmon, Oncorhynchus tshawytscha (Kurtovic et al. 2010). The
enzyme is a carboxyl ester lipase (often called bile salt activated lipase) and
requires calcium ions and sodium cholate for optimum activity. However, it
should be noted that the salmon lipase can operate at a lower rate without bile
salts, so these could be omitted from a food product or other application, if
necessary. The versatility of bile salt activated lipases towards catalyzing
many different types of reactions (Wang and Hartsuck 1993; Hui and Howles
2002; Kurtovic et al. 2009a) is of significant potential benefit from an
applications standpoint. The salmon lipase exhibits the highest activity against
a medium chain (C10:0) ester of p-nitrophenol, and the activity at lower
111
temperatures is relatively high compared to other fish and mammalian
carboxyl ester lipases (Kurtovic et al. 2010).
To the best of our knowledge, immobilization of an active and functional bile
salt activated lipase from any source has not been demonstrated to date. This
manuscript describes the immobilization of a bile salt activated lipase from
Chinook salmon on two different hydrophobic supports, octyl-Sepharose and
Lewatit VP OC 1600. Catalytic performance of the immobilized salmon
lipases against several emulsified lipid substrates is compared with that of
Novo 435.
5.3. Materials and methods
5.3.1. General
Bile salt activated lipase was extracted from the pyloric ceca of farmed
Chinook salmon (The New Zealand King Salmon Co. Ltd, Nelson, New
Zealand). The ceca were frozen at -80°C immediately after collection
(Kurtovic et al. 2010). Ghee (clarified butter) was purchased from a local
store. Fish oil extracted from New Zealand hoki (Macruronus
novaezealandiae) was supplied by SeaDragon Marine Oils Ltd (Nelson, New
Zealand). Benzoyl-DL-arginine-p-nitroanilide (BA-p-NA), octyl-Sepharose
CL-4B (Oct-S), p-nitrophenyl palmitate (p-NPP), and tributyrin were
purchased from Sigma Chemical Co. (St. Louis, MO, USA). P-
aminobenzamidine-cellulose (p-ABA-cellulose) was supplied by Life
Technologies (Invitrogen New Zealand Ltd, Nelson, New Zealand). Lewatit
VP OC 1600 (Lew) was supplied by Lanxess (Bayer, Leverkusen, Germany)
and Novozym 435 FG (Novo 435) was from Novozymes (Bagsvaerd,
Denmark).
5.3.2. Lipase extraction and purification with p-ABA-cellulose
Frozen pyloric ceca were crushed, lyophilised, delipidated sequentially with
several organic solvents, and air-dried to yield the pyloric ceca powder
112
(Kurtovic et al. 2010). The ceca powder was stirred in 20 mM Tris buffer (pH
7.2) for 30 min at 21°C. The ratio of buffer (mL) to powder (g) was 20:1. The
supernatant (crude extract) was recovered by centrifugation at 10,000 g for 15
min using a Beckman Avanti® J-25 I centrifuge (Beckman Coulter, Inc.,
Fullerton, CA, USA). This supernatant was batch loaded onto a trypsin affinity
resin, p-ABA-cellulose, which had been pre-equilibrated with Tris buffer then
collected on a sintered glass funnel under vacuum. The ratio of crude extract
(mL) to damp resin (g) was 6:1. Purification with the affinity resin was carried
out to remove trypsin and, to a lesser extent, other serine proteases (Cohen et
al. 1981; Yang et al. 2009). Binding was carried out at 8°C for 1 h, using a
continuous rotating mixer. The supernatant (void) was recovered by
centrifugation at 3,000 g for 5 min. Subsequently, the resin was washed once
with the Tris buffer for 5 min and the wash recovered. The void and wash
were combined (p-ABA v-w) as the lipase solution.
5.3.3. Immobilization
Oct-S and Lew were washed with copious amounts of Milli-Q water,
equilibrated with Tris buffer, and collected on a sintered glass funnel under
vacuum. Equal volumes of the lipase solution (mL) were mixed with each
damp resin (g) in a 7:1 ratio. Binding, using a continuous rotating mixer, was
carried out at 21°C for 2 and 4.5 h for Oct-S and Lew, respectively. Each
supernatant (void) was recovered by centrifugation at 3,000 g for 5 min.
Subsequently, each resin was washed with Tris buffer for 5 min, and the
combined void-wash was retained. The resins were washed with the buffer
two more times and the washes discarded. The resins were then rinsed and
soaked in Tris buffer containing 20 mM calcium chloride and 5 mM sodium
cholate for 1.5 h, with gentle end-over-end mixing at 21°C. After recovery on
a sintered glass funnel, the resins were dried over silica gel under vacuum for
15 h at 21°C. The protein loading on the supports (mg protein/g support) was
calculated from the difference between lipase load and combined void-wash
fractions.
113
5.3.4. Trypsin activity assay
Throughout the course of immobilization trypsin activity was assayed at 30°C
using BA-p-NA as substrate (Erlanger et al. 1961). The reaction mixture
comprised an aliquot of the sample (25–100 µL) plus 1 mM BA-p-NA in 0.1
M Tris buffer (pH 8.2, containing 10 mM calcium chloride and 1% (v/v)
DMSO) to make a final reaction volume of 3 mL. The sample was replaced
with 20 mM Tris buffer (pH 7.2) to establish blank-rate activity. The release of
p-nitroaniline from BA-p-NA was measured at 410 nm in a Unicam (model
UV4) UV-visible spectrophotometer (Thermo Electron Corporation, Waltham,
MA, USA) fitted with a peltier temperature controller. One unit (U) of activity
was defined as 1 µmol p-nitroaniline released/min under the assay conditions,
using the extinction coefficient of 8,800 M-1
cm-1
(Erlanger et al. 1961).
5.3.5. Lipase activity assays during immobilization
Two methods were used to measure the lipase activity throughout the course
of immobilization. The first was a spectrophotometric method, utilizing the
hydrolysis of p-NPP in an aqueous medium at 30°C and pH 8 (Kurtovic et al.
2010). The second method was titrimetric, utilizing the hydrolysis of tributyrin
(modified from Committee on Codex Specifications 1981). Reactions were
carried out at 30°C with 70 mL aqueous substrate emulsion (5 mM tributyrin,
0.18% (w/v) sodium caseinate and 0.015% (w/v) lecithin) in a stirred pH stat
vessel (718 STAT Titrino, Metrohm, Herisau, Switzerland). Calcium chloride
(2 mM) and sodium cholate (5 mM) were added at the start of the assay from
0.5 and 0.25 M stock solutions (in Milli-Q water), respectively. Each reaction
was monitored for 10–45 min, depending on the sample (dry resins containing
immobilized lipases required 30–45 min assays for reproducibility). The
released free fatty acids were titrated with 0.01 M NaOH with the pH stat set
at 8.0. The slope (mL 0.01 M NaOH/min) in the linear region of the titration
curve (typically the initial rate of reaction) was used to calculate the activity.
Blank-rate activity was determined in the absence of the sample. One unit of
activity was defined as 1 µmol butyric acid released/min under the assay
conditions.
114
5.3.6. Immobilized lipase activity against natural substrates
Immobilized lipases (dry resins) were tested for activity against a ghee
emulsion and a fish oil emulsion and compared with Novo 435. The assays
were titrimetric, conducted as for the tributyrin assay, except that the stat pH
was either 9.0 (immobilized salmon lipases) or 6.75 (Novo 435), the optimum
pH for the respective lipase. For the ghee emulsion, reactions were carried out
at 35°C with 70 mL aqueous substrate emulsion (35% ghee, 2.2% sodium
caseinate, 0.4% lecithin and 0.1% gum arabic, all w/v). For the fish oil
emulsion, reactions were carried out at 30°C with 70 mL aqueous substrate
emulsion containing 5% (w/v) fish oil and 5% (w/v) gum arabic (based on
Desnuelle et al. 1955). Calcium chloride (2 mM) and sodium cholate (5 mM)
were included in the assays for immobilized salmon lipases. Each reaction was
monitored for 30–90 min. The initial rate of reaction was used to calculate the
activity in all cases. Blank-rate activity was determined with the respective
resin only in place of the immobilized lipase sample. One unit of activity was
defined as 1 µmol fatty acid released/min under the assay conditions. All
assays were carried out in duplicate.
5.3.7. Protein determination
Protein concentrations were measured using a method based on that of Lowry
et al. (1951) with modifications to the concentration of reagents and
incubation times. Bovine serum albumin (BSA) was used as the standard.
Modified standard curves in the presence of either SDS or Triton X-100 were
used when desorbing the immobilized proteins/lipase from the supports using
detergents (2%, w/v).
5.3.8. Properties of free and immobilized lipase
Properties were assessed with the 5 mM tributyrin assay, described in section
5.3.5. Samples were p-ABA v-w (free lipase) and salmon lipase-Oct-S
(immobilized lipase). To assess the effect of pH on activity, the substrate
emulsion was adjusted to the required pH (ranging from 6.0 to 10.0) prior to
sample addition. To study the effect of temperature on activity, the substrate
emulsion was equilibrated at temperatures from 20 to 50°C for 10 min prior to
115
sample addition. The effects of temperature on lipase stability were
investigated by incubating the sample at various temperatures (20 to 60°C) for
30 min using a digital block heater, then rapidly cooling on ice to 0°C, prior to
assay. The effect of a primary bile salt on activity was measured in the absence
and presence of 5 mM sodium cholate. Calcium chloride was present in all
assays. In addition to using tributyrin as substrate, activity of the immobilized
lipase only was measured against the ghee emulsion (described in section
5.3.6) in both the absence and presence of cholate. All assays were carried out
at optimum pH for each sample.
5.4. Results and discussion
5.4.1. Lipase immobilization on hydrophobic supports
The removal of trypsin/serine proteases with p-ABA-cellulose was necessary
to prevent the proteolytic degradation of lipase during immobilization. This
step was crucial for the retention of lipase activity in the immobilized form,
since protease inhibitors (e.g. benzamidine), bile salts and glycerol were
omitted from the extraction buffer. Although these compounds protect the
lipase from inactivation (Blackberg and Hernell 1993; Kurtovic et al. 2006;
Kurtovic et al. 2010), they were not included in the buffer as they can interfere
with the hydrophobic interactions necessary for lipase binding (unpublished
results). This is especially the case with bile salts, which act as surfactants and
can form strong interactions with hydrophobic supports (Armstrong and Carey
1982).
The activity results are divided into two parts. Based on the activities against
BA-p-NA, trypsin – the most abundant digestive protease in Chinook salmon
– was removed almost completely by the affinity resin (Table 5.1).
Table 5.1
Activities (U) against BA-p-NA (measurement of trypsin activity) for the
purification of Chinook salmon lipase on p-ABA-cellulose
Sample Total volume (mL) U/mL Total U Recovery (%)
116
Crude extract 51.0 ± 0.5 1.442 ± 0.001 73.5 ± 0.7 100
p-ABA v-w a 85.0 ± 0.5 0.051 ± 0.001 4.3 ± 0.1 5.9 ± 0.2
Assays were carried out at 30°C; Values are mean ± SE (n = 2).
a v-w = combined void-wash fractions.
With p-NPP as substrate, low free-lipase activity was obtained in Oct-S and
Lew v-w fractions, indicating that the lipase had bound to both supports (Table
5.2). However, the immobilized salmon lipases also exhibited very low
activities against this synthetic substrate. Our studies indicate that
conformational changes in the enzyme and/or increased hydrophobicity
following immobilization adversely affected the substrate access. Similarly,
the activity of Novo 435 against p-NPP was <2-fold that of salmon lipase-Oct-
S.
Tributyrin appeared to be a more appropriate substrate to measure the amount
of lipase (activity) bound on the supports, especially Oct-S (Table 5.2). When
immobilized on this resin, the estimated total recovery of salmon lipase
activity (including the unbound and bound lipase) was >60%. In contrast, it
seems that the conformational orientation of the bound lipase and the
hydrophobic character of the Lew resin did not favour either p-NPP or
tributyrin. The change in properties of the fish lipase due to the immobilization
meant that it was difficult to estimate the amount of lipase bound on the Lew
resin with either of these substrates. Salmon lipase-Oct-S had higher activity
against tributyrin than the same lipase immobilized on Lew or Novo 435. On a
dry weight basis, this activity was approximately 40- and 10-fold higher than
those of salmon lipase-Lew and Novo 435, respectively.
Table 5.2
Lipase activities (U) against p-NPP and tributyrin for the immobilization of
Chinook salmon lipase on two hydrophobic supports (Oct-S and Lew)
Sample Total
volume (mL)
U/mL or g a Total U Recovery (%)
p-NPP Tributyrin p-NPP Tributyrin p-NPP Tributyrin
Load 42.5 ± 0.5 1.737 ±
0.002
13.2 ± 0.2 73.8 ±
0.9 561 ± 9
100 100
Oct-S v-w b 63.0 ± 0.5 0.054 ±
0.001 3.1 ± 0.4
3.40 ±
0.07 190 ± 20
4.6 ±
0.1 35 ± 4
117
Lew v-w 67.0 ± 0.5 0.053 ±
0.001 0.20 ± 0.05
3.52 ±
0.04 13 ± 3
4.8 ±
0.1 2.3 ± 0.5
Salmon lipase-Oct-S
(dry) 0.490 ± 0.005
g
0.42 ±
0.04 330 ± 2
0.21 ±
0.02 163 ± 2
0.28 ±
0.03 29.0 ± 0.6
Salmon lipase-Lew
(dry) 2.310 ± 0.005
g
0.0024 ±
0.0002 8.6 ± 0.4
0.006 ±
0.001 20 ± 1
0.010 ±
0.001 3.5 ± 0.2
Novo 435 (dry) c -
0.77 ±
0.09 32 ± 2
- -
- -
Oct-S/Lew damp weight was 6.0 g; assays were carried out at 30°C; Values
are mean ± SE (n = 2).
a U/mL for liquid samples; U/g for immobilized lipases.
b For all v-w samples, v-w = combined void-wash fractions.
c Included for comparison.
Considerably more protein bound to Oct-S than Lew (207.0 and 60.1 mg/g of
dry resin, respectively). However, this is only true for the dry forms of the
immobilized lipases. Due to its agarose backbone, Oct-S is considerably more
hydrophilic than Lew and swells to approximately 12 times its dry weight
when wet, whereas Lew absorbs a lot less water. Thus, in a practical sense
where damp resins are used (i.e. in our experiments), higher protein loading
was observed for Lew (25.9 compared to 17.3 mg/g of damp resin for Oct-S).
These results are in agreement with the results for CAL-B binding onto octyl-
agarose and Lew (Cabrera et al. 2009).
The complete removal of salmon proteins with either 2% Triton X-100, a non-
ionic detergent, or 2% SDS, an anionic detergent, was not achieved with either
support. The lipase could not be visualized by SDS-PAGE (data not shown).
This suggests a relatively tight binding of the salmon lipase on both carriers.
The tight binding is a desirable characteristic that would allow multiple reuse
cycles with minimal leaching of the enzyme.
Bile salt activated lipases appear to bind more strongly than microbial lipases
as CAL-B was desorbed completely from octyl-agarose and Lew using 1–2%
Triton X-100 (Cabrera et al. 2009). Bile salt activated lipases are relatively
large and complex proteins. They have a large hydrophobic patch surrounding
the active site and two bile salt binding sites, one of which is a large active site
loop (Blackberg and Hernell 1993; Wang et al. 1997; Moore et al. 2001).
When the loop is in a closed form, the large hydrophobic patch is not
118
completely shielded (Moore et al. 2001). In contrast, microbial lipases have a
smaller active site lid that upon closure completely covers the hydrophobic
residues around the active site. They also lack any bile salt binding sites
(Jaeger et al. 1999; Moore et al. 2001; Sharma et al. 2001). The structural
differences between the two types of lipases are likely to result in stronger
hydrophobic interactions between the salmon lipase and the hydrophobic
supports compared with CAL-B.
5.4.2. Immobilized lipase activity against natural lipid substrates
The catalytic performance of the immobilized salmon lipases was examined
with two lipids with very different fatty acid profiles, and compared with that
of Novo 435. Of all the natural fats, milkfat/ghee is the most complex and
contains a relatively high proportion of saturated, short chain (9%, w/w) and
medium chain (17%, w/w) fatty acids (Balcão and Malcata 1998; Lindmark
Mansson 2008). In contrast, the majority of fatty acids in hoki oil are long
chain, with a considerable proportion (18–22%, w/w) of n-3 PUFAs (McLean
and Bulling 2005). The percentage of ghee in the emulsion (35%) was
equivalent to the fat content in the standardized dairy cream sold in New
Zealand. For a representative substrate containing a considerable proportion of
short chain fatty acids, ghee emulsion was used instead of cream as the fat
does not separate in the assay mixture. The assay with ghee was carried out at
35°C, the minimum temperature required to prevent the substrate from
solidifying. A 35% fish oil emulsion made in the same manner as the ghee
emulsion resulted in very low or no activity with all the samples. The
optimized final emulsion contained 5% fish oil and gum arabic.
With ghee as substrate, the specific activity of the immobilized CAL-B (Novo
435) was higher than that of the salmon lipase immobilized on Lew, but lower
than that of the fish enzyme immobilized on Oct-S (Table 5.3). When
examined for the chain length selectivity against saturated fatty acids, Novo
435 was most active against C12–C16 fatty acids (Karabulut et al. 2009).
Although ghee contains ~ 45% saturated C12–C16 fatty acids (Lindmark
119
Mansson 2008), the emulsion used in our study does not appear to be an
optimal substrate for Novo 435.
Table 5.3
Activity (U) of the immobilized Chinook salmon lipases and Novo 435 against
ghee and fish oil emulsions
Lipase U/g
Ghee Fish oil
Salmon lipase-Oct-S (dry) 69 ± 4 47 ± 2
Salmon lipase-Lew (dry) 29 ± 3 5 ± 1
Novo 435 (dry) 44 ± 5 2.5 ± 0.4
Assays were carried out at 35/30°C (ghee/fish oil);
Values are mean ± SE (n = 2).
It has been demonstrated that Novo 435 is efficient at both cleaving and
synthesizing triglycerides (TGs) containing long chain unsaturated fatty acids
from sunflower, cod liver and microalgal oils in non-aqueous media
(Haraldsson 1999; Robles Medina et al. 1999; Deng et al. 2005). However, in
our study Novo 435 was unable to hydrolyze the TGs in a fish oil emulsion.
This was also the case for salmon lipase-Lew. Significant activity against the
emulsion was only achieved with salmon lipase-Oct-S. There are several
reasons for the difference in activity between Oct-S- and Lew-bound enzymes.
The reasons are likely to include the micelle size in the emulsified substrate
and enzyme conformation. However, it also seems reasonable to conclude that
the hydrophobicity or hydrophilicity of the resins plays a significant part.
The conformation of the bound lipase and its orientation on the carrier directly
affect the catalytic performance. For salmon lipase-Oct-S, the attachment
likely involves the large hydrophobic patch surrounding the active site of the
lipase, which requires the active site loop to open fully (Moore et al. 2001).
The lipase may undergo a partial interfacial activation during immobilization
(Fernandez-Lafuente et al. 1998) because the hydrophobic groups on the resin
act as a substrate analogue. The bound lipase conformation would therefore
120
favour a relatively higher activity against the substrates. In addition, Oct-S is
more hydrophilic than Lew so the emulsified lipid substrate will have a higher
affinity towards Oct-S in an aqueous medium (Hanefeld et al. 2009). This is in
agreement with salmon lipase-Oct-S having a higher activity against the
emulsified substrates (specifically tributyrin and fish oil) than salmon lipase-
Lew.
For salmon lipase-Lew, the hydrophobic or hydrophilic character of the resin
is likely to have a greater influence on the catalytic performance than the
bound lipase conformation. Physical entrapment (Chen et al. 2008) and
weaker hydrophobic adsorption (Chen et al. 2008; Cabrera et al. 2009) without
interfacial activation are the likely modes of attachment of salmon lipase on
Lew. Lew consists of a crosslinked polymer and lacks the well-defined and
dense planar hydrophobic surface created by the octyl residues in Oct-S
(Fernandez-Lafuente et al. 1998). Lew, the more hydrophobic carrier, does not
favour the partition of the emulsified substrate from the aqueous reaction
medium to the resin, which could explain the lower activities observed in our
study.
In summary, salmon lipase-Oct-S is the least hydrophobic lipase-carrier
complex, followed by salmon lipase-Lew, and then Novo 435, and these
differences are mostly responsible for the differences in activity of these
enzymes against the emulsified substrates.
5.4.3. Effect of immobilization on pH and thermal properties and bile salt
requirement of Chinook salmon lipase
Immobilization may alter the properties of an enzyme and therefore its use in
applications so it was important to determine if the salmon lipase had different
properties as a result of the immobilization process. Of the two immobilized
salmon lipases, only salmon lipase-Oct-S was investigated as it had higher
activity against tributyrin, as well as the natural substrates (Tables 5.2 and
5.3).
The free form of salmon lipase was most active within a narrow pH range of
8.0–8.5 (Fig. 5.1). Similarly, the highest activity for the immobilized lipase
121
was measured at pH 8.0–9.0. A previous study on porcine pancreatic lipase
reported an increase in pH optimum after immobilization (Kartal et al. 2009).
Fig. 5.1. Activity of free (– – –) and immobilized (—) Chinook salmon lipase
against tributyrin as a function of pH. Duplicate measurements were taken at
each pH.
An increase in the temperature optimum of the immobilized lipase was seen
compared with the free form (Fig. 5.2). The immobilized lipase had the
highest activity at 45°C, whereas the free form was most active at 35°C. The
increase in temperature optimum is likely due to the stabilization of enzyme
structure (i.e. decreased enzyme flexibility), and has been reported for many
immobilized enzymes (Kartal et al. 2009). Although both the free and
immobilized forms have high hydrolytic activity (>60%) at lower
temperatures, the immobilized form is less sensitive to temperature changes up
to the optimum temperature for hydrolysis.
20
40
60
80
100
5 6 7 8 9 10 11
Rel
ati
ve
act
ivit
y (
%)
pH
122
Fig. 5.2. Activity of free (– – –) and immobilized (—) Chinook salmon lipase
against tributyrin as a function of temperature. Duplicate measurements were
taken at each temperature.
Improved thermal stability of an enzyme is often one of the outcomes of the
immobilization process (Holm and Cowan 2008; Hanefeld et al. 2009). In the
present study, the thermal stability of the salmon lipase appeared unchanged
upon immobilization (Fig. 5.3). Both forms of the lipase lost a significant
amount of activity when incubated above 30°C. The thermal properties of the
immobilized salmon lipase make it particularly useful for low temperature
applications. An example of such an application is the production of long
chain, highly unsaturated fatty acids, where mild reaction conditions are
necessary to minimize the oxidation of fatty acids (Haraldsson 1999).
60
70
80
90
100
10 20 30 40 50 60
Rel
ati
ve
act
ivit
y (
%)
Temperature ( C)
123
Fig. 5.3. Effect of temperature on the stability of free (– – –) and immobilized
(—) Chinook salmon lipase when incubated in duplicate for 30 min prior to
assay. Residual activities are relative to the un-incubated sample and were
measured against tributyrin.
The immobilized fish lipase was soaked in buffer containing calcium ions and
cholate before drying. Despite this treatment, it was found that the
immobilized salmon lipase required additional bile salts for optimum activity
against tributyrin. However, the activity in the absence of cholate was still
high for both the free and immobilized forms (74–76% of the cholate-inclusive
activity). The high activity in the absence of cholate is not unexpected as bile
salt activated lipases are known to have basal activity against smaller
substrates, such as the short chain monoacid TGs (Wang and Hartsuck 1993).
Cholate was also required for optimum activity of the immobilized salmon
lipase in a ghee emulsion. In the absence of cholate the activity was 55–60%
of the optimum. The relatively lower activity of the immobilized lipase in the
absence of cholate against ghee, compared with that against tributyrin, is due
to lack of basal activity against the bulkier, water-insoluble TGs in ghee.
5.5. Conclusions
This study presents a successful immobilization of Chinook salmon bile salt
activated lipase. When immobilized on Oct-S, the lipase had higher activity
0
20
40
60
80
100
120
10 20 30 40 50 60
Res
idu
al
act
ivit
y (
%)
Temperature ( C)
124
against tributyrin than the same lipase immobilized on Lew (~ 40-fold higher)
and CAL-B immobilized on Lew (Novo 435, ~ 10-fold higher activity).
Salmon lipase-Oct-S also had high activity against both ghee and fish oil
emulsions. Salmon lipase-Lew only showed significant activity against the
ghee emulsion. The discrepancies in activities between the two forms of
immobilized fish lipase are attributed in part to the conformation of the bound
lipase, but most importantly to the different hydrophobicities of the lipase-
carrier complexes, which determine the substrate access for hydrolysis.
Immobilization appeared to raise the temperature optimum of the salmon
lipase against tributyrin, but did not affect the thermal stability of the enzyme.
In addition, the bile salt requirement of the lipase was not affected by
immobilization.
The immobilized fish lipase (salmon lipase-Oct-S) hydrolyzed natural
substrates containing a wide range of fatty acids. One was a substrate with a
relatively high proportion of short chain fatty acids (ghee), and the other
contained high amounts of long chain n-3 PUFAs (fish oil). This versatility in
substrate utilization and activity at low temperatures show that Chinook
salmon lipase immobilized on Oct-S has potential for applications which
require the modification of lipids in aqueous media.
Further studies on the immobilization of fish bile salt activated lipases are in
progress, relating in particular to the improvement of catalysis in an oil
medium for the production of free fatty acids and alkyl esters.
5.6. Acknowledgements
This study was supported by funds from the New Zealand Foundation for
Research, Science & Technology (FRST) under contracts C02X0301 and
C02X0806. We thank The New Zealand King Salmon Co. Ltd for providing
salmon tissue samples, Lanxess for providing Lewatit VP OC 1600 resin,
Novozymes for providing Novozym 435, SeaDragon Marine Oils Ltd for
providing the hoki oil, and Dr. Graham Down of Life Technologies
(Invitrogen NZ Ltd) for providing the cellulose resin and sharing his extensive
knowledge of resin chemistry.
125
CHAPTER VI. GENERAL DISCUSSION AND CONCLUSION
Enzymes have been touted as the future of sustainable industrial processing as
they promise lower energy usage, fewer waste products, and greater reaction
specificity than traditional chemical processing techniques. Like their
microbial and mammalian counterparts, fish lipases have the potential to
reduce energy costs in industrial processes, purify specific FAs, deliver
structured lipids for nutraceutical use, and produce particular flavours in food
products. Marine lipases have the potential to deliver different activities to
those of currently available commercial lipases sourced from microorganisms
and the few mammalian sources. However, one of the main challenges for the
commercial exploitation of the fish enzymes is the conformational instability
that makes extraction and processing very difficult. In this thesis, the activity
of lipases from two commercial fish species in New Zealand, Chinook salmon
and New Zealand hoki was investigated with the particular goal of improving
the enzyme stability and allowing the novel activities of the two lipases to be
exploited. There is potential to extract these enzymes from underutilised waste
streams of fish processing in New Zealand and develop high-value enzyme
products, especially through immobilization.
Lipase activity assay development was carried out prior to the purification
studies. For the spectrophotometric assay using p-NPP, the initial assays were
based on previous published research on both microbial and fish lipases (e.g.
Winkler and Stuckmann 1979; Gjellesvik et al. 1992; Pencreac'h and Baratti
1996). Using the published methods, very low activities were measured in the
hoki crude extract. This led to various improvements in the assay, the most
effective being the inclusion of calcium ions and sodium cholate. The
requirement for cholate for optimum activity is not unique to salmon and hoki
lipases as other bile salt activated lipases also require primary bile salts for
optimum activity. The lack of bile salt inclusion in previous studies is most
likely due to the use of a ‗standard‘ rather than an optimized assay. The final
outcome in the present work was a quick, sensitive and reliable assay for fish
bile salt activated lipases.
126
It was important to stabilize the pyloric ceca for highest yield of lipase activity
as it enabled easier and more reliable monitoring of the enzyme recovery
during the purification. The use of lyophilization and delipidation protocols
achieved this goal. Water removal prior to the removal of lipid was crucial for
the activity preservation. When the ceca were delipidated without first
lyophilizing them the loss of stability was attributed to the removal of a water
monolayer around the lipase, together with the free water, by the hydrophilic
solvents (i.e. acetone and 1-butanol) used during delipidation (Zaks and
Klibanov 1988; Klibanov 1989).
The instability of the two fish lipases during extraction and purification was
partially overcome through modifications to the extraction/purification buffer.
The inclusion of Na cholate, glycerol and protease inhibitors stabilized the
lipase. Whilst protease inhibitors were used during fish lipase purification in
previous studies (Gjellesvik et al. 1992; Iijima et al. 1998), the inclusion of
cholate and glycerol was not very common. Protein precipitation with PEG
1000, another less commonly used technique during enzyme purification, was
included in the present study.
Lyophilization treatments carried out during the purification protocol also
contributed to improved lipase stability. The treatment was used to concentrate
the affinity fraction, as well as the final purified lipase preparations, SDL and
HDL. The treatment produced a viscous liquid instead of a powder,
maintaining the lipase in a soluble form. The combination of stabilization
buffers and lyophilization made possible the purification and characterization
studies on the two lipases.
An affinity chromatography protocol using cholate-Affi-Gel 102 was
optimized for the fish lipases. For the removal of low affinity bound proteins,
0.4% Na cholate was chosen as a compromise between satisfactory removal of
non-lipase proteins and minimal removal of lipase. Lipase was removed
completely with 1.2% cholate, while some of the more tightly bound proteins
were left behind. Both the elution of lipase with 2 or 2.5% Na taurocholate
(Gjellesvik et al. 1992) and the removal of non-specifically bound proteins
with 0.5–1% Na deoxycholate (Wang 1980; Wang and Johnson 1983) were
127
unsuccessful. This was due to the compacting of the resin which impeded
buffer flow.
In the present study, the final degrees of purification for the two enzymes were
lower than those in some of the previously published studies on fish lipases
(Mukundan et al. 1985; Gjellesvik et al. 1992; Iijima et al. 1998). The
difference was attributed to the relatively high specific activity in the starting
material in the present study, which was due to the stabilization of the starting
material and optimization of the extraction buffer. The relatively lower levels
of purification and poor stability indicated that the fish lipases would need to
be immobilized before commercial use could be contemplated.
The visualization of purified fish lipase activity in a polyacrylamide gel using
a substrate overlay (zymography) was very exciting, as it has not been
reported previously. This is particularly significant considering the low
stability of the two lipases. As was the case with lipase assay optimization,
stability was achieved with the inclusion of cholate and calcium in the running
buffer. The successful zymographic analysis provides other researchers with a
valuable tool for studying fish lipases.
Marine lipases are at present underutilized because not enough is known about
their characteristics. The research presented in this thesis has expanded
significantly the knowledge of fish bile salt activated lipases. The
classification of the two digestive lipases as ‗bile salt activated‘ adds to the
growing body of evidence that many fish digestive lipases of marine origin
belong to this type (Table 2.1). Both Chinook salmon and hoki lipases shared
some characteristics with other fish lipases. These include the pH and
temperature optima for activity and low thermal stability. But they also
differed from most fish lipases in that they were more acid stable and
exhibited maximal activity against a medium chain ester of p-NP (cf. mullet
lipase was most active against the palmitate ester, Aryee et al. 2007). The
stability at slightly acid to neutral pH values, together with the high activity
against a medium chain ester of p-NP and a relatively high activity at low
128
temperatures presents opportunities for the applications of the salmon and hoki
lipases in food (dairy) products.
Maximal activity against a medium chain ester of p-NP indicated a potential
for the hydrolysis of lipids containing short and medium chain FAs, such as
milkfat. Therefore, the lipases from salmon and hoki were evaluated as flavour
modifying agents in dairy products. The fish enzymes gave a result similar to a
well-established commercial lipase used in dairy products (calf PGE) in terms
of the total amount and types of FAs released over the course of the reaction.
Like PGE, the fish enzymes released mainly short chain FAs from cream
triglycerides, with the highest specificity towards the key dairy product flavour
and odour compounds, butanoic and hexanoic acids. Furthermore, the
differences in FFA composition between PGE and salmon lipase-treated
creams were not significant.
It has been shown previously that marine lipases have specificity towards the
long chain PUFAs that are abundant in fish oil and other marine lipids (Patton
et al. 1975; Patton et al. 1977; Gjellesvik 1991; Gjellesvik et al. 1992; Koven
et al. 1994b; Koven et al. 1997; Olsen et al. 1998). Thus, it was quite
interesting that the salmon and hoki lipases had specificity towards short chain
FAs in cream TGs, instead of the longer chain, saturated and monounsaturated
FAs. The high reactivity towards short chain FAs is attractive from an
applications standpoint. If used in commercial dairy products, different flavour
profiles between salmon and hoki lipase-modified dairy lipids would be
expected due to slight differences in the short chain FA specificity of the two
lipases. There may be an opportunity to use fish lipases in commercial dairy
applications, especially if they can deliver particular and desirable flavours not
provided by the enzymes in use now. To make this successful there are
barriers to overcome. Despite the raw material (pyloric ceca) being both cheap
and abundant, there are significant costs associated with developing large-
scale, food-grade extraction processes, although the work in this thesis
indicates that extraction is certainly possible. There may also be issues with
consumer acceptance.
129
Although the free forms of the fish lipases are suitable for use in food (dairy)
products, applications such as bioindustrial modifications of fats and oils (e.g.
the production of FFA concentrates or TGs enriched with PUFAs) require the
lipases to be immobilized on a permanent support. Immobilization separates
an enzyme from the reaction products, allows for multiple re-use and
continuous processing (Fernandez-Lafuente et al. 1998; Holm and Cowan
2008). The salmon lipase was bound to two supports through hydrophobic
interactions. Immobilization allowed the lipase to be used effectively for the
hydrolysis of a variety of emulsified lipid substrates, and would facilitate use
in continuous large-scale processes. Immobilization also helped overcome the
lipase instability problem, both during use and in storage. The immobilized
lipase in dry form can be stored at room temperature. The study provided
some novel insights into the immobilization of a bile salt activated lipase and
factors that need consideration, such as: overall hydrophobicity of the
immobilized lipase-carrier complex; mode of attachment and the
conformation/orientation of the lipase as the result of such attachment; and its
predicted effect on catalytic efficacy (substrate partitioning from the bulk
reaction medium towards the lipase-carrier support). A feature of the salmon
lipase bound onto the hydrophobic supports is the strong binding that should
minimize leaching and allow multiple reuse cycles.
Salmon lipase-octyl-Sepharose had high activity in the aqueous medium
containing emulsified lipid substrates. The immobilized lipase was used to
effectively hydrolyze two natural substrates containing short or long chain
FAs: ghee, with a relatively high proportion of short chain FAs, and fish oil,
with high amounts of long chain n-3 PUFAs. This versatility in substrate
utilization and activity at low temperatures show that Chinook salmon lipase
immobilized on octyl-Sepharose has potential for applications for lipid
modifications in aqueous media.
Immobilization of a bile salt activated lipase from any source has not been
reported previously. Bile salt activated lipases are relatively large and complex
proteins that contain a large active site lid (Blackberg and Hernell 1993; Wang
et al. 1997; Moore et al. 2001). The lipase needs to be immobilized in an
130
active conformation (with the lid open). Another barrier to immobilization is
the need for multi-step purification and cofactors to improve the stability of
the lipases, making the process unpractical and costly. Lipases sourced from
microorganisms are immobilized routinely as they are easier to extract/purify
and can be readily manipulated via genetic tools to suit application demands.
Nevertheless, the work in this thesis indicates that immobilization of the
salmon lipase on supports with densely-packed, strongly hydrophobic, lipid-
like groups is certainly possible. The salmon lipase can be adsorbed and
purified at the same time as the support is highly selective.
Both the free and immobilized forms of the salmon enzyme displayed relative
insensitivity to temperature change and high activity at lower temperatures
(Figs. 3.9 and 5.2). The salmon enzyme is thus particularly suited for low
temperature applications, such as cleavage of TGs to long chain, highly
unsaturated FA concentrates, where mild reaction conditions are necessary to
minimize FA oxidation (Haraldsson 1999).
Despite their potential as catalysts, and as relatively high-tech products to add
value to fisheries by-products, fish lipases present challenges for extraction
and use that have so far resisted commercial exploitation. The enzymes are
unstable and their concentrations in tissue cannot match those of the microbial
enzymes. Fish bile salt activated lipases need calcium to function and operate
more efficiently in the presence of bile salts. These properties have limited
their potential use in commercial applications. However, the present research
has shown that digestive lipases from Chinook salmon and hoki have
properties that are different to currently available commercial enzymes. These
properties may offer advantages in some applications. Whilst these enzymes
are labile and hard to extract, the stabilization buffers and immobilization
protocols developed through this work may allow their use in industrial
processes.
131
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APPENDIX I. DETAILS OF METHODS AND RESULTS AND
ADDITIONAL EXPERIMENTS FOR CHAPTER III
Appendix I contains the supplementary material omitted from the manuscript
for brevity.
Methods
Buffer formulation trials for higher extraction activity and lipase stability
A series of trials was carried out in order to obtain higher lipase activity in the
crude extract and improve the stability of the enzyme. Crude extracts were
prepared using ceca powder (g) to buffer (mL) ratio of 1:30 in all experiments,
unless noted otherwise. The pH of each extract was measured (to check for
buffering efficiency of the chosen buffer) and re-adjusted if necessary. Lipase
activity was measured against p-NPP. All measurements were done in
duplicate.
Effect of protease inhibitors on hoki lipase activity:
Since lipases need to be protected from protease digestion during the course of
purification, tests were carried out with the commonly used protease
inhibitors, benzamidine, PMSF and EDTA, in order to check their effect on
lipase activity. Crude extract was prepared in 25 mM Tris buffer (pH 8) and
incubated with an inhibitor (1–5 mM final concentration) for 30 min at 4°C
and the residual activity was measured. All incubations were done in
duplicate. Control incubation was either purified water (for benzamidine and
EDTA) or 2-propanol (for PMSF).
Effect of benzamidine concentration and slightly acidic pH extraction on hoki
lipase stability:
Crude extract was prepared in either 25 mM Tris buffer (pH 8.5, with 5 mM
CaCl2 and 0, 2, 5 or 10 mM benzamidine) or 25 mM Bis-Tris buffer (pH 6,
with 5 mM CaCl2). The extracts were kept unfrozen at 0°C. Besides lipase
activity, trypsin activity against BA-p-NA (Kurtovic et al. 2006), and
157
chymotrypsin activity against N-Succinyl-Gly-Gly-Phe-p-NA (Achstetter et al.
1981) were also measured. The activities were monitored over 72 h.
Effect of bile salts and 1% (w/v) glycerol on hoki lipase activity/stability:
Crude extract was prepared in 25 mM Tris buffer (pH 8.5, with 5 mM CaCl2, 2
mM benzamidine and either 10 mM Na cholate, 10 mM Na deoxycholate, 1%
glycerol, or 10 mM Na cholate and 1% glycerol) and kept at 0°C. Lipase
activity was monitored over 24 h.
Effect of a divalent metal cation chelating agent (EDTA) and Na cholate
concentration on hoki lipase activity/stability:
Crude extract was prepared in either 25 mM Tris buffer (pH 8.5, with 5 mM
CaCl2, 2 mM benzamidine and 10 mM Na cholate) or pH 8.5 TBE (25 mM
Tris, 2 mM benzamidine, 1 mM EDTA, with 5, 10 or 20 mM Na cholate). The
extracts were kept at 0°C and lipase activity was monitored over 72 h.
Lipase extraction at high pH:
In an attempt to further reduce the protease activity in the crude extract,
extraction of salmon lipase was carried out with pH 10, 10.5 and 11 buffers.
The crude extract of salmon lipase was used instead of hoki as it had higher
activity, allowing more reliable monitoring during storage. The buffers
contained 25 mM glycine, 2 mM benzamidine, 1 mM EDTA and 20 mM Na
cholate. Extraction was performed in the same manner as with pH 8.5 TBE
containing 20 mM Na cholate. Besides lipase activity, trypsin activity
(Kurtovic et al. 2006) and elastase activity against N-Succinyl-Ala-Ala-Ala-p-
NA (Bieth et al. 1974) were also measured.
Buffer trials after removal of Na cholate:
Since Na cholate interfered with purification of lipase by affinity
chromatography, it was omitted from the extraction buffer. A series of further
tests were then carried out on the salmon lipase with modified extraction
buffers in an attempt to improve lipase extraction and maintain stability.
Protein concentration in all crude extracts was measured with the Lowry
assay. Lipase stability at 4°C was monitored over 96 h.
158
For the first set of trials, the crude extract was prepared in 100 mM TBE (pH
8.5), the same buffer containing 5% glycerol, or the same buffer containing
0.1 M NaCl. Glycerol levels of 10, 15, 20 and 25% in the extraction buffer
were used in the second set of trials. For the third set of trials, the crude extract
was prepared in 100 mM TBE (pH 8.5, containing 20% glycerol) using ceca
powder to buffer ratios of 1:10, 1:30 or 1:50. In the fourth set of trials,
phosphate (Na2HPO4) was compared to Tris as the buffering agent. Crude
extract was prepared in either pH 8.0 TBE20 or pH 7.8 PBE20 using ceca
powder to buffer ratios of 1:10 and 1:30. The pH values <8.5 were used as the
buffering range of phosphate is 6–8.
Lipase assays
Three assay methods were tested with hoki crude extract as the sample:
The first was a direct titration method using an autotitrator in the pH-STAT
mode (Committee on Codex Specifications 1981). The assays were done at
25°C. Tributyrin, triolein, olive oil and cholesterol esters (palmitate or oleate)
were used separately as the substrate. The released FAs were titrated with 0.01
M NaOH with the pH stat set at 8.0. For tributyrin assays, different
concentrations of substrate (17 (original), 10, 5 and 1 mM), emulsifiers (Na
caseinate, 0.04–0.6% (w/v) and lecithin, 0.003–0.05% (w/v)), calcium ions (1–
10 mM), Na cholate (0.1, 0.5, 1, 5 and 10 mM), taurocholate and deoxycholate
(5 mM) and different sample volumes (0.2–2 mL) were tested. Regarding
triolein, olive oil and cholesterol ester assays, several emulsifiers/detergents
(e.g. lecithin, Tweens, stearic acid), dispersing agents/stabilizers (e.g. Na
caseinate, albumin, gum arabic) and solvents (e.g. 1-butanol, 2-propanol,
dichloromethane) were tested as aids for the preparation of substrate mix.
Polyols (glycerol and Trigol) were used in some assays in an attempt to
improve the emulsion by making the medium more hydrophobic. Most of the
assays contained calcium ions and Na cholate (details in ‗Results and
discussion‘, Table A1.5). Ultrasonic homogenization was utilized in a few
instances to improve the emulsion. One unit (U) of activity was defined as 1
μmol FA (e.g. butyric acid from tributyrin) produced/min.
159
The second was the p-NPP assay. The following were tested during assay
optimization: nine solvents (methanol, ethanol, ethylene glycol, 2-propanol,
DMSO, 1-butanol, heptane, dichloromethane and chloroform) for stock
substrate preparation and compatibility with the Tris buffer; several final
substrate concentrations (0.125, 0.25, 0.5 and 1 mM) and sample volumes
(0.02, 0.05, 0.1 and 0.3 mL); the effects of calcium ions (10 and 50 mM), Na
cholate (5 and 10 mM), NaCl (0.05 to 0.15 M), emulsifiers (0.1 and 0.5%
Tween 80 or lecithin), gum arabic (0.01 and 0.1%) and magnetic stirring
during the assay.
A spectrophotometric/turbidimetric assay using olive oil as the substrate
(Iizuka et al. 1991) was the third assay. The assays were done at 30°C. The
increase in turbidity from released FAs was monitored at 340 nm. The
intention for testing this assay was to obtain a quick and sensitive method for
lipase activity using a natural long chain TG substrate. Stock substrate
solutions were initially prepared in 1-butanol and ethanol (Iizuka et al. 1991).
The variations tested were: stock substrate solution prepared in 2-propanol;
different stock substrate concentrations; and several final substrate
concentrations. The effects of individual bile salts, calcium ions, bicarbonate,
albumin, glycerol, and magnetic stirring during the assay were also examined.
One unit of activity was defined as absorbance change of 0.001/min. All
assays were carried out in duplicate.
Protein determination
The following methods were compared: Lowry (Lowry et al. 1951); enhanced
Lowry (Stoscheck 1990); biuret (Gornall et al. 1949); and absorbance at 280
nm (A280), with and without A260 (correction for nucleic acids, (Groves et al.
1968)). Bovine serum albumin was used as the standard in all assays.
Results and discussion
Buffer formulation trials for higher extraction activity and lipase stability
The tests with protease inhibitors showed that <20% of lipase activity was lost
after incubations with benzamidine and EDTA, independent of inhibitor
160
concentration. Incubation with PMSF resulted in 30% less lipase activity.
Therefore, benzamidine and EDTA were used in the subsequent extraction
buffers.
The effect of benzamidine concentration and slightly acidic pH extraction on
hoki lipase stability is shown in Table A1.1. The lipase activity was not
retained in all extracts. Chymotrypsin activity was present in all crude extracts
(especially 0 and 2 mM benzamidine) and significant levels remained even
after 72 h. Trypsin activity was also present in 0 and 2 mM benzamidine and
pH 6 extracts, up to 72 h. The protease activity contributed to the decline in
lipase activity over the storage period. Although 5 mM benzamidine was
adequate for complete trypsin inhibition, 27% less lipase activity was
extracted with 5 mM compared to 0 and 2 mM benzamidine. Therefore, 2 mM
benzamidine was used in the subsequent extraction buffers.
Table A1.1
Effect of benzamidine concentration and slightly acidic pH extraction on
relative lipase activity (%) in the crude extract from hoki pyloric ceca, with
storage at 0°C for 72 h
Storage
time (h)
Extraction buffer
pH 6 0 mM benz 2 mM benz 5 mM benz 10 mM benz
0 51 100 100 73 66
2.3 39 83 90 66 58
24 16 12 36 28 35
72 7 7 5 5 7
The effect of bile salts and 1% glycerol on hoki lipase stability is shown in
Table A1.2. Similar to the results in Table A1.1, <40% of lipase activity was
retained after 24 h. Although 10 mM deoxycholate had the same effect on
lipase stability as 10 mM cholate, cholate was more effective at lipase
extraction. In addition, when deoxycholate was added to the extraction buffer,
a finely dispersed white precipitate (Ca deoxycholate) was formed. Therefore,
cholate was used in the subsequent extraction buffers.
161
Table A1.2
Effect of bile salts and glycerol on relative lipase activity (%) in the crude
extract from hoki pyloric ceca, with storage at 0°C for 24 h
Storage
time (h)
Extraction buffer
10 mM
cholate
10 mM
deoxycholate
1%
glycerol
10 mM cholate
and 1% glycerol
0 100 76 47 99
1 92 71 45 92
2 87 74 43 87
24 37 38 23 29
The effect of EDTA and Na cholate concentration on hoki lipase stability is
shown in Table A1.3. Full lipase stability was achieved for 24 h at 0°C with
the samples containing EDTA and lacking calcium ions. Lack of Ca ions
reduces the activity of serine proteases (Kurtovic et al. 2006). Although Ca
ions are necessary for optimum lipase activity, they have a negative effect on
stability. On the other hand, cholate protects the lipase from protease attack
(Blackberg and Hernell 1993; Loomes 1995). Besides chelating Ca ions,
EDTA is able to bind other divalent metal cations in the crude extract and
reduce any inherent metalloprotease activity. Slightly higher activity and
stability were obtained with 20 mM than with 10 mM cholate. Overall, highest
activity and stability of hoki ceca lipase was obtained with pH 8.5 TBE
containing 20 mM Na cholate.
Table A1.3
Effect of EDTA and Na cholate concentration on relative lipase activity (%) in
the crude extract from hoki pyloric ceca, with storage at 0°C for 72 h
Storage
time (h)
Extraction buffer
Ca and
10 mM cholate
EDTA and
5 mM cholate
EDTA and
10 mM cholate
EDTA and
20 mM cholate
0 92 93 97 100
1 83 92 100 100
162
2 80 85 87 94
24 60 95 106 116
72 12 35 38 40
Lipase extraction at high pH:
Lipase extraction at pH 10 produced only 10% of activity typically extracted
at pH 8.5 and the activity decreased on storage at 4°C. Extraction at pH values
>10 resulted in lower lipase activities. Significant trypsin and elastase
activities were present in all three extractions, indicating that the proteases
were not fully denatured at high pH values. Overall, extraction of salmon
lipase at pH ≥10 was not effective.
Buffer trials after removal of Na cholate:
The following results were obtained from the first three sets of trials: the
highest specific activity and stability were achieved with 100 mM TBE (pH
8.5, containing 5% glycerol) extraction; the activity and stability were
improved with 20% glycerol in the extraction buffer; the highest specific
activity was obtained with the 1:30 ratio, whereas stability after >4 h was
slightly higher (~5%) with the 1:10 ratio, and the crude extract obtained with
the 1:50 ratio had the lowest specific activity and stability of the lipase.
The effect of PBE20 extraction on salmon lipase specific activity and stability
is shown in Table A1.4. The crude extracts obtained with the 1:10 ratio had
slightly higher stabilities than those obtained with the 1:30 ratio for both
PBE20 and TBE20. However, the pH value of the crude extract obtained with
the 1:10 ratio was 7.7 instead of 8.0 for the Tris extract and 7.1 instead of 7.8
for the phosphate extract. Thus, 1:30 was a more appropriate ratio. Overall,
PBE20 and TBE20 extracts had similar specific activities. However, lipase
stability was higher in the phosphate extracts. PBE20 was therefore used in the
ultimate purification protocol for both salmon and hoki lipases (Fig. 3.1).
Table A1.4
Comparison of specific lipase activity (U/mg) in the crude extract from
Chinook salmon pyloric ceca obtained with TBE20 and PBE20 buffers, with
storage at 4°C for 96 h
163
Storage
time (h)
Extraction buffer and ratio
TBE20, 1:10 TBE20, 1:30 PBE20, 1:10 PBE20, 1:30
0 0.28 0.40 0.25 0.35
2 0.28 0.31 0.29 0.35
24 0.25 0.20 0.28 0.29
36 0.23 0.12 0.26 0.21
96 0.03 0.03 0.09 0.09
Lipase assays
The titrimetric assay using tributyrin was improved approximately three-fold
when compared to the starting conditions (no CaCl2 and bile salts; tributyrin
and emulsifier concentrations as per reference). Highest activity (27 U/mL)
was obtained with 5 mM tributyrin, 5 mM Na cholate, 2 mM CaCl2, 0.18%
(w/v) Na caseinate and 0.015% (w/v) lecithin in water (Table A1.5). Although
10 mM CaCl2 was initially found to be required for optimum activity, it
produced too high blank rate of base addition, most likely due to the formation
of Ca(OH)2 precipitate. When ≤2 mM CaCl2 was used the blank rate was
negligible and the optimum activity was largely unaffected. Sample volumes
of 0.5–1 mL gave the most reproducible results. The activity against tributyrin
was >10-fold higher compared to that against olive oil/triolein and activity
against cholesterol esters was very low.
Table A1.5
Lipase activities of crude extract from hoki pyloric ceca obtained with the
titrimetric assay
Substrate and other assay components Activity (U/mL)
Tributyrin (17 mM) with 0.6% Na caseinate and 0.05%
lecithin
9.0
Tributyrin (5 mM) with 0.18% Na caseinate and 0.015%
lecithin
11.0
+ 2 mM CaCl2 12.5
+ 2 mM CaCl2 and 1 mM Na cholate 23.0
164
+ 2 mM CaCl2 and 5 mM Na cholate 27.0
+ 2 mM CaCl2 and 5 mM Na taurocholate 16.0
+ 2 mM CaCl2 and 5 mM Na deoxycholate 4.0
Olive oil/triolein (5 mM) with 0.18% Na caseinate and
0.015% lecithin + 2 mM CaCl2 and 5 mM Na cholate
1.2
Olive oil/triolein (20 mM) with 0.3% Na caseinate and
0.3% lecithin + 20 mM CaCl2 and 20 mM Na cholate and
10 μM albumin; sonicated
1.8
Cholesteryl palmitate (5 mM) with 0.18% Na caseinate
and 0.015% lecithin + 2 mM CaCl2 and 5 mM Na cholate
0.7
Values are average of duplicate determinations.
Regarding the p-NPP assay, solubility of p-NPP was very low in methanol,
ethanol, ethylene glycol and DMSO, making these solvents unsuitable for
stock substrate preparation. In contrast, p-NPP readily dissolved in heptane,
dichloromethane and chloroform. However, these solvents were incompatible
with the Tris buffer, even at <2% (v/v). Adequate p-NPP solubility and buffer
compatibility was achieved with both 2-propanol and 1-butanol. Addition of
Tween 80 to the buffer inhibited lipase activity significantly. Similarly,
lecithin did not emulsify the substrate effectively as it appeared to cause
precipitation, making the assay mixture cloudy and unsuitable for
spectrophotometric measurements. Addition of NaCl to the buffer caused a 5–
10% decrease in lipase activity. The highest activity and fully linear reaction
rate profile (for at least 3 min) was obtained with 0.25 mM final p-NPP
concentration (from 15 mM stock in 2-propanol), 20 mM CaCl2 and 5 mM Na
cholate in 20 mM Tris-HCl (pH 8.0, with 0.01% gum arabic). The assay was
improved ~100-fold compared to the original one (no CaCl2 and bile salts). It
was also found that using p-NPP stock older than 24 h resulted in lower
activities, due to spontaneous substrate breakdown. This assay is quick,
consistent and more sensitive than the titrimetric one. It is also particularly
suitable for lipase since a long chain FA ester of p-NP is used in order to
exclude non-specific esterase activity. Overall, the p-NPP assay was the most
reliable and convenient method for monitoring the activity during the
purification procedure.
165
All turbidimetric assays produced non-linear reaction profiles after ~30 s (i.e.
the rate graphs of dAbs vs. time were concave). This assay was found
unreliable as it produced inconsistent results which were difficult to interpret
accurately.
Protein determination
The Lowry method was the most reliable due to the most linear standard curve
(Fig. A1.1), high sensitivity and reproducible results. The enhanced Lowry
method, although most sensitive, did not produce very linear standard curve
(R2 = 0.90) and had greater variations within the replicate measurements. The
biuret method required relatively large sample volumes due to low sensitivity,
which was inconvenient when using purified samples. In addition, there was a
problem of chemical interferences (e.g. by EDTA and Na cholate) with the
assay when larger sample volumes were used. Lastly, measuring the
absorbance at 280 nm, both with and without the correction for nucleic acids
(A260) gave consistently higher protein concentration estimates compared to
the other three methods and was thus considered unreliable.
Fig. A1.1. Standard curve for the Lowry protein assay.
R² = 0.9995
R² = 0.9994
0
0.2
0.4
0.6
0.8
1
1.2
0 100 200 300 400 500
A5
40
Protein (μg)
166
Hoki lipase and other omitted results
The following tables and figures (relating mainly to hoki lipase) were omitted
from the manuscript for brevity. However, all results were included in the
discussion.
Table A1.6
Purification scheme for hoki digestive lipase
Purification step Total
volume (mL)
Protein conc.
(mg/mL)
Total protein
(mg)
Total activity
(U)
Specific
activity (U/mg)
Activity
recovery (%)
Purification
(-fold)
Crude extract 28 ± 0 6.1 ± 0.3 167 ± 7 21 ± 2 0.13 ± 0.01 100 1
PEG 1000 fraction 34 ± 1 4.6 ± 0.1 155 ± 5 17 ± 1 0.11 ± 0.01 83 ± 12 0.9 ± 0.1
Dialysate 63 ± 6 2.1 ± 0.2 134 ± 5 15 ± 1 0.11 ± 0.01 73 ± 4 0.9 ± 0.1
Affinity fraction 34 ± 3 0.06 ± 0.01 2.1 ± 0.5 1.3 ± 0.1 0.63 ± 0.13 47 ± 1* 4.9 ± 0.7
Affinity fraction
conc.
9 ± 1 0.22 ± 0.03 1.9 ± 0.3 1.2 ± 0.1 0.65 ± 0.09 47 ± 2* 5.1 ± 0.4
GF fraction 19 ± 2 0.023 ± 0.005 0.43 ± 0.06 0.40 ± 0.03 0.94 ± 0.21 48 ± 3* 7.4 ± 1.3
These results are compiled from three separate purifications.
*Recovery back-calculated as if all material from previous step was used for
further purification.
Table A1.7
Preparation of SDL and HDL
Sample Total volume
(mL)
Total protein
(mg)
Total
activity (U)
Specific activity
(U/mg)
Activity
recovery (%)
S GF pooled 52 0.55 2.22 4.1 100
S GF dialysate 57 0.49 1.94 3.9 88
S GF lyophilized
(SDL)
14 0.47 1.85 4.0 84
H GF pooled 32 0.70 0.66 0.95 100
H GF dialysate 35 0.67 0.63 0.93 95
H GF lyophilized
(HDL)
9 0.63 0.60 0.94 90
167
0
0.01
0.02
0.03
0.04
0.05
5 10 15 20 25 30
A2
80
(
) or
Lip
ase
act
ivit
y (
U/m
l) (
)
tube number
Fig. A1.2. A representative elution profile of hoki lipase on Sephacryl S-300
HR. Tubes 17-22 (indicated by dashed line) were pooled as the GF fraction.
Fig. A1.3. IEF-PAGE of gel filtration concentrates: purified hoki (lanes 1 and
2, 6.5 µg each) and salmon digestive lipase (lanes 4, 5 and 6; 7.5, 7.5 and 2.6
µg, respectively); pI standards (lane 3).
Lanes 2 and 4 were contaminated with proteins from the standards and were
excluded from the analysis. This contamination of the lanes immediately
9.6
8.0
7.5
7.1
7.0
6.5
6.0
5.1
4.45
pI1 3 5 6
168
adjacent to the standards was observed with all four repeats of the gels and
appears to be the result of lengthy separation time using high voltages.
Fig. A1.4. Acyl-chain specificity of hoki lipase with p-nitrophenyl esters.
Activities are relative to p-NPC. b = basal activity in the absence of Na
cholate. Values are means ± SD (n = 3).
Fig. A1.5. Effect of bile salts on hoki lipase activity against p-NPC. Activities
are relative to 5mM cholate (standard assay). Values are means ± SD (n = 3).
Calculation of kinetic parameters
The following graphs were used to derive the kinetic parameters of the two
lipases.
169
Fig. A1.6. Michaelis-Menten plot for the hydrolysis of p-NPC by Chinook
salmon lipase at 35°C. Values are means ± SD (n = 3). R2= 0.9928.
Fig. A1.7. Michaelis-Menten plot for the hydrolysis of p-NPC by hoki lipase
at 35°C. Values are means ± SD (n = 3). R2= 0.9966.
The Michaelis-Menten plots for both lipases suggested a hyperbolic
relationship between velocity and substrate concentration. Therefore,
modifications to the Michaelis-Menten equation that result in a linear
relationship can be used to estimate the kinetic parameters.
0
0.02
0.04
0.06
0.08
0.1
0.12
0 50 100 150 200 250
v o(U
/mL
)
[p-NPC], (µM)
0
0.01
0.02
0.03
0.04
0.05
0.06
0 50 100 150 200
v o(U
/mL
)
[p-NPC], (µM)
170
Fig. A1.8. Lineweaver-Burke plot for the hydrolysis of p-NPC by Chinook
salmon lipase at 35°C. Values are means ± SD (n = 3).
From this plot, Vmax = 0.151 ± 0.024 U/mL and Km = 91.6 ± 33.2 µM. The
large variations in the parameters highlight the weakness of this plot, where
the errors at very low substrate concentrations are greatly exaggerated. A
better plot is the Hanes:
Fig. A1.9. Hanes plot for the hydrolysis of p-NPC by Chinook salmon lipase
at 35°C. Values are means ± SD (n = 3).
y = 588.35x + 6.8014
R² = 0.9993
y = 459.87x + 7.8575
R² = 0.9951
y = 716.83x + 5.7452
R² = 0.9967
0
20
40
60
80
100
120
140
160
-0.02 0.02 0.06 0.1 0.14 0.18 0.22
1/v
o(m
L/U
)
1/[p-NPC], (µM-1)
y = 7.2168x + 566.99
R² = 0.9898
y = 6.9497x + 491.02
R² = 0.9707
y = 7.4838x + 642.96
R² = 0.9805
0
500
1000
1500
2000
2500
-100 -50 0 50 100 150 200 250
[p-N
PC
]/v o
(µM
mL
/U)
[p-NPC], (µM)
171
With Hanes plot, the experimental errors retain a constant contribution over
the whole range of substrate concentrations. From this plot, Vmax = 0.139 ±
0.005 U/mL and Km = 78.3 ± 7.6 µM. Using the protein concentration of 0.035
± 0.002 mg/mL for SDL (Table A1.7) and salmon lipase Mw of 54.9 ± 0.4
kDa (Fig. 3.5), [lipase] = 0.63 ± 0.04 µM and turnover number (k2) = 3.674 ±
0.25 s-1
. Thus, the final values of kinetic parameters for the salmon lipase
were: Km = 78 ± 8 µM and k2 = 3.7 ± 0.3 s-1
.
For hoki lipase:
Fig. A1.10. Lineweaver-Burke plot for the hydrolysis of p-NPC by hoki lipase
at 35°C. Values are means ± SD (n = 3).
Vmax = 0.080 ± 0.004 U/mL and Km = 85.3 ± 26.9 µM.
y = 1059.5x + 12.61
R² = 0.9983
y = 772.64x + 13.217
R² = 0.9976
y = 1346.3x + 12.003
R² = 0.9947
0
20
40
60
80
100
120
140
160
-0.02 0 0.02 0.04 0.06 0.08 0.1 0.12
1/v
o(m
L/U
)
1/[p-NPC], (µM-1)
172
Fig. A1.11. Hanes plot for the hydrolysis of p-NPC by hoki lipase at 35°C.
Values are means ± SD (n = 3).
Vmax = 0.071 ± 0.003 U/mL and Km = 68.1 ± 13.1 µM. Using the protein
concentration of 0.074 ± 0.004 mg/mL for HDL (Table A1.7) and hoki lipase
Mw of 44.6 ± 0.4 kDa (Fig. 3.5), [lipase] = 1.70 ± 0.09 µM and k2 = 0.709 ±
0.049 s-1
. Thus, the final values of kinetic parameters for hoki lipase were: Km
= 68 ± 13 µM and k2 = 0.71 ± 0.05 s-1
.
Additional experiments
Other purification techniques tested using hoki lipase extracts
Methods
Lipase precipitation at acidic pH values
Precipitation of lipase (and other proteins) was carried out at pH 4 and 5.
Dilute HCl was slowly added with mixing to the crude extract until the
required pH was reached. The samples were then centrifuged and pellets were
re-suspended in the original volume of extraction buffer (TBE, pH 8.5, with 20
mM Na cholate). Lipase, trypsin (Kurtovic et al. 2006), chymotrypsin
(Achstetter et al. 1981) and elastase (Bieth et al. 1974) activities were
measured in the crude extract and the re-suspended pellets.
y = 14.175x + 972.86
R² = 0.9934
y = 13.583x + 746.45
R² = 0.9912
y = 14.768x + 1199.3
R² = 0.9794
0
500
1000
1500
2000
2500
3000
3500
4000
4500
-100 -50 0 50 100 150 200
[p-N
PC
]/v o
(µM
mL
/U)
[p-NPC], (µM)
173
Ammonium sulphate fractionation
This procedure is used frequently during enzyme purification. Stepwise
fractionation of the crude extract was started at 15%, then increased to 25, 35
and finally 45% of saturation. In addition, a one-step treatment (0–45 or 0–
63% of saturation) was also attempted. Lipase and protease activities were
monitored throughout the procedure.
Ion exchange chromatography
Experiments were first carried out in a batch-wise manner to find the pH at
which the lipase binds to an anion (DEAE-cellulose, medium grade) or cation
(SP-cellulose, medium grade) exchanger (both resins supplied by Invitrogen
New Zealand Ltd, Nelson, New Zealand). The pH range was 5–9 for DEAE
and 4–8 for SP with 0.5 pH unit intervals. The resins were pre-washed with 1
M NaCl and rinsed thoroughly with Milli-Q water until the conductivity of the
wash was ~0.1 mS. The resins were then equilibrated to correct pH values
with several buffer washes. The buffers used were Bis-Tris (pH 5–7) and Tris
(pH 7.5–9) for DEAE trials, and citrate (pH 4–6) and phosphate (pH 6.5–8) for
SP trials. All buffers had conductivities of <5 mS. Sample (crude extract
prepared in 2 mM benzamidine and 1 mM EDTA solution without any
buffering agent) was added to the wet resins and mixed for 30 min at 4°C. The
ratio of sample (mL) to resin (g) was 2:1. Crude extract and resin supernatant
(unbound fraction) were tested for lipase activity. Control experiments omitted
ion exchange resins. All measurements were done in duplicate.
DEAE-cellulose column chromatography:
The objective of this experiment was to test lipase binding efficiency, elution
conditions, and activity recovery. The experiment was done at 4°C. DEAE-
cellulose (6 g) was packed into an 8 mL column and equilibrated with the
loading buffer (pH 8.5 TBE with 20 mM Na cholate). Crude extract (9 mL)
was prepared in the same buffer and applied onto the column at 6 mL/h
followed by loading buffer. The ratio of sample (mL) to resin (g) was 1.5:1
(instead of 2:1) in order to improve lipase binding. The flow-through and
elution fractions were monitored for protein concentration at 280 nm. Stepwise
gradient elution was then performed at 80–100 mL/h with elution buffers
174
containing NaCl (in 50 mM increments) in the loading buffer. The resin was
soaked in 200 mM NaCl elution buffer for 15 h. Elution was continued with
250 mM NaCl elution buffer at 16 mL/h. Further elutions were carried out
with NaCl concentrations up to 2 M.
QAE-cellulose column chromatography:
Purification with QAE-cellulose (medium grade, Invitrogen New Zealand Ltd)
was performed as an alternative to DEAE-cellulose, in a manner similar to
DEAE experiment. The experiment was done at 8°C, with protein monitoring
(A280) of the fractions. PEG 1000 fraction (4 mL) was applied to the column,
followed by loading buffer (as in DEAE experiment) at a flow rate of 11
mL/h. Lipase elution was attempted with 0.1, 0.2 and 1 M NaCl in the loading
buffer, at 40-70 mL/h. The resin was also soaked with 1 M NaCl for 15 h to
facilitate further lipase desorption.
Hydrophobic interaction chromatography
Two hydrophobic interaction chromatography trials with butyl- and phenyl-
cellulose (Invitrogen New Zealand Ltd, Nelson, New Zealand) were carried
out with both crude extract and dialysate. TBE buffer (pH 7 or 9) containing 1
M NaCl or 4 M KCl was used as the loading buffer and elution was carried out
with the same buffers in the absence of salt. The experiments were carried out
in a batch-wise manner and binding time was 1 h.
Results and discussion
Lipase precipitation at acidic pH values
Both supernatants had very low or no measurable lipase activity. However,
only 36% of starting activity was recovered from the pH 4 pellet, while the
lipase recovery from the pH 5 pellet was 57%, suggesting lipase denaturation
at both pH values, as a result of precipitation process. The protease activity
was relatively high (>60%) in both re-suspended pellets, indicating that they
co-precipitated with the lipase. Overall, this technique did not result in any
lipase purification.
175
Ammonium sulphate fractionation
The lipase precipitated at every stage of stepwise fractionation. The combined
15, 25 and 35% pellets contained ~50% of starting activity, which meant that
neither the supernatants nor the pellets had adequate activity levels required
before further purification steps. With a one-step treatment (0–45%
saturation), 65–80% of activity was recovered in the re-suspended pellet.
However, the pellet also contained most of the trypsin and elastase activities
and ~50% of the starting chymotrypsin activity. Very similar results were
obtained when ammonium sulphate was added to 63% of saturation.
Ion exchange chromatography
The pH binding experiments indicated that 30 and 40% of lipase did not bind
to DEAE-cellulose at pH 8–9 and 6–7.5, respectively. In contrast, >60% of
lipase activity was recovered in the unbound fractions from SP-cellulose at pH
6–8. Lipase binding was not improved with >30 min contact time. Binding
was not achieved at pH 4–6 for both ion exchangers due to protein
precipitation. The results indicated that DEAE-cellulose would be most
suitable to use with a loading buffer of pH ≥8.
DEAE-cellulose column chromatography:
The first 14 mL of flow-through contained 61% of applied lipase activity. This
was re-applied onto the column. The second flow-through fraction (49 mL)
contained 65% of activity applied originally, indicating that there was no
further binding upon re-application. Very low activity (2–3% of applied
activity) was recovered in each of the four elution fractions (25–40 mL each)
up to 200 mM NaCl. Another 2% of activity was recovered after soaking the
resin with 200 mM NaCl elution buffer. A further 2% of activity was obtained
from the elutions with ≥250 mM NaCl. Overall, lipase binding was low (≤33%
of applied activity), and <50% of bound lipase activity was recovered.
Regarding QAE-cellulose chromatography, ~65% of protein and 60% of lipase
activity loaded on the column were measured in the flow-through fraction,
indicating low lipase binding. No lipase activity was eluted with 0.1, 0.2 and 1
M NaCl, even after soaking the resin for 15 h. Protein peaks were observed
176
during every elution, however the peaks had no lipase activity. Overall, no
purification of lipase was achieved through QAE-cellulose procedure.
It was decided to discontinue further IE chromatography trials as the amount
of lipase bound was low and recoveries of the bound lipase were inconsistent.
Hydrophobic interaction chromatography
The unbound fraction contained <50% of the applied activity. However, no
lipase activity was detected in the eluted fraction in both trials. The results
indicated strong binding of lipase to the hydrophobic resins. Consequently,
hydrophobic interaction chromatography was not used in the final purification
scheme. Nevertheless, hydrophobic supports were used for the immobilization
of Chinook salmon lipase (Chapter V).
177
APPENDIX II. DETAILS OF METHODS AND RESULTS AND
ADDITIONAL EXPERIMENTS FOR CHAPTER V
Appendix II contains the supplementary material omitted from the manuscript
for brevity. Three sections from the manuscript are expanded.
Methods
Trypsin and lipase activity assays for immobilized lipases
A modified BA-p-NA/p-NPP assay was used for immobilized lipases (dry
resins). Since these samples were insoluble, a stopped assay was used instead
of the kinetic one. The sample (50–150 mg) was weighed accurately into a 15
mL centrifuge tube and the assay buffer was added. For ‗blank‘ activity
measurements, the respective resin only was used in place of the immobilized
lipase sample. After equilibrating the sample at 30°C in a shaking water bath,
the reaction was started by adding an aliquot of the substrate stock solution to
make a final reaction volume of 3 mL. When a light yellow colour was clearly
visible (generally 10–20 min), the reaction was stopped by pouring the
reaction mixture into a 10 mL disposable syringe to which a 0.2 µm disposable
filter cartridge (Sartorius, Goettingen, Germany) had been attached. The
filtrate was collected free of any resin sample and its absorbance measured at
410nm. A410 values for each sample were less than 1.00, which ensured the
substrate was non-limiting (zero order kinetics).
Protein desorption from lipase supports
Each immobilized salmon lipase was mixed with a 2% (w/v) detergent
solution (SDS or Triton X-100 in water). The ratio of detergent solution (mL)
to lipase sample (g) was ≥20, which ensured the resin was fully swollen.
Desorption was carried out for 1 h 45 min at 21°C, with end-over-end mixing.
The supernatant (desorbed protein solution) was recovered by centrifugation
and used in Lowry assay and SDS-PAGE. The solution was concentrated with
10 kDa MWCO Vivaspin concentrators prior to loading on the gel.
178
Results and discussion
Protein desorption
Triton X-100 was selected for bound salmon protein removal as it has been
used to remove CAL-B from several hydrophobic supports, including octyl-
agarose and Lewatit VP OC 1600 (Chen et al. 2008; Cabrera et al. 2009).
However, Triton X-100 was found to be incompatible with SDS-PAGE (lanes
containing Triton X-100 showed streaking after staining), and SDS was used
as an alternative detergent.
The protein balance for salmon lipase immobilization on each support is
summarized in Table A2.1.
Table A2.1
a) Protein balance for the immobilization of Chinook salmon lipase on Oct-S
Sample Total
volume
(mL)
Protein
mg/mL or g a
Total
protein
(mg)
Recovery
(%)
Crude extract 39 12.9 502
p-ABA v-w 59 6.9 410 81.5
Load for Oct-S 48 6.9 333
Oct-S v-w 75 3.4 254 76.3
Salmon lipase-Oct-S (dry)
– SDS desorption 0.38 g
12.9 4.9 1.5
Salmon lipase-Oct-S (dry)
– Triton X-100 desorption 0.38 g
8.0 3.1 0.9
b) Protein balance for the immobilization of Chinook salmon lipase on Lew
Sample Total
volume
(mL)
Protein
mg/mL or g a
Total
protein
(mg)
Recovery
(%)
Crude extract 51 14.1 719
p-ABA v-w 85 7.14 607 84.4
Load for Lew 42.5 7.14 303
179
Lew v-w 67 2.46 165 54.3
Salmon lipase-Lew (dry)
– SDS desorption 2.31 g
10.3 23.8 7.8
Salmon lipase-Lew (dry)
– Triton X-100 desorption 2.31 g
9.0 20.7 6.8
a mg/mL for liquid samples; mg/g for immobilized lipases.
Only a fraction of the expected bound salmon protein was desorbed from both
supports. More protein was desorbed from Lew than from Oct-S, suggesting
relatively weaker hydrophobic interactions between Lew and salmon
proteins/lipase. Of the two detergents, SDS was slightly more effective at
protein desorption from both supports.
When analyzed by SDS-PAGE, a faint band corresponding to Mw of ~55 kDa
was present in the crude extract and p-ABA v-w (fraction containing proteins
that did not bind to trypsin affinity resin). However, this band was not seen in
any of the other fractions (Fig. A2.1). The band is likely to be the salmon
lipase. The fraction containing proteins desorbed from trypsin affinity resin
with 0.1 M acetic acid (Ac-p-ABA) revealed one major band corresponding to
Mw of ~28 kDa, indicative of trypsin (Kurtovic et al. 2006). Many proteins
with different Mws did not bind to either Lew or Oct-S, as revealed by the
presence of multiple bands in Lew v-w and Oct-S v-w fractions. A band with
Mw of ~120 kDa in SDS-Oct-S (proteins desorbed from salmon lipase-Oct-S
with SDS) could be a lipase dimer. The absence of a lipase monomer in SDS-
Oct-S and SDS-Lew is due to low protein desorption from the supports.
180
Fig. A2.1. 12.5% acrylamide SDS-PAGE of fractions from the immobilization
of Chinook salmon lipase on Oct-S and Lew. Molecular weight standards
(kDa) are in the outside lanes. Samples containing Triton X-100 caused
streaking and were omitted.
a Proteins desorbed from trypsin affinity resin with 0.1 M acetic acid.
Additional experiments
Hydrolysis of fish (hoki) oil with immobilized salmon lipases and Novozym
435 in non-aqueous media
Methods
Hydrolysis in solvent-free medium (pure oil)
Each immobilized lipase (0.175 g salmon lipase-Oct-S, 0.5 g salmon lipase-
Lew, 0.5 g Novo 435) was added to 10 g hoki oil. The reaction was carried out
at 35°C using rotating mixer for salmon lipases and control (oil only), and at
60°C using shaking water bath for Novo 435. Samples of the reaction products
were taken at the start and after 2 h. Milli-Q water (200 µL) was then added to
each reaction mixture. Another sample was taken at 4 h reaction time. Solid
calcium chloride and sodium cholate were then added to a final concentration
of 5 mM to the reaction mixtures containing immobilized salmon lipases and
control. Further samples from all reaction mixtures were taken at 8 and 22 h
reaction time. TLC of all samples was carried out. The mobile phase contained
200
116
97.4
66.2
45
31
21.5
Crude ext p-ABA v-w Ac-p-ABAa Lew v-w SDS-Lew Oct-S v-w SDS-Oct-S
181
hexane and diethyl ether (70:12.4, v/v). Lipid classes were visualized with
primulin, a fluorescent dye.
Hydrolysis in hexane
The protocol was based on that of Desai et al. (2006). Each immobilized lipase
(0.175 g salmon lipase-Oct-S, 0.5 g salmon lipase-Lew, 0.5 g Novo 435) was
added to 6 mL hexane containing 30 mg (~5 mM) hoki oil. The reaction was
carried out at 30°C using rotating mixer for salmon lipases and control (oil in
hexane only), and at 60°C using shaking water bath for Novo 435. Samples of
the reaction products were taken at the start and after 3 h. Calcium chloride
(60 µL of 0.5 M stock solution in water) and sodium cholate (120 µL of 0.25
M stock solution in water) were then added to a final concentration of 5 mM to
the reaction mixtures containing immobilized salmon lipases and control. Only
water (180 µL) was added to the reaction mixture containing Novo 435.
Further samples were taken at 7 and 20 h reaction time. TLC of all samples
was carried out as indicated previously.
Results and discussion
Fig. A2.2 shows the results of the hydrolysis of fish oil in hexane. Equivalent
results were obtained for the hydrolysis in solvent-free medium. In both
media, FFAs were produced by Novo 435, but not with immobilized salmon
lipases. The hydrophobicity or hydrophilicity of lipase-carrier complexes plays
a significant part. Salmon lipase-Oct-S is relatively hydrophilic and salmon
lipase-Lew is also not hydrophobic enough for oil/hexane medium. Thus, the
substrate affinity towards immobilized salmon lipases is low. On the other
hand, Novo 435 is likely to be the most hydrophobic of the three complexes,
hence its high rate of FFA production. An additional treatment during the
preparation of Novo 435 (as was pointed out by (Cabrera et al. 2009)) is
proposed. The additional treatment may make the lipase-carrier complex more
hydrophobic and/or change the enzyme conformation.
182
Fig. A2.2. TLC of the products of hoki oil hydrolysis with immobilized
salmon lipases and Novo 435 in hexane. Total reaction time was 20 h. The
mixed standard contained monolein, diolein, triolein and ethyl nonadecanoate.
FFA/FAEE standard contained saturated FAs of various chain lengths and
ethyl nonadecanoate.
Control Salmon lipase-Lew Salmon lipase-Oct-S Novo 435 FFA/FAEE
0 7 20 3 7 20 Mixed Std 3 7 20 3 7 20 h Std
FAEE
Sterol
Esters
TG
FFA
DG
MG
