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ELECTROCHEMICAL STUDIES OF HYDROLYTIC PRODUCTS FROM REFINED PALM OIL WITH IMMOBILIZED ENZYMES AT

MODIFIED CARBON ELECTRODES

Zahraa A. Jarjes

UNIVERSITI SAINS MALAYSIA

2012

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ELECTROCHEMICAL STUDIES OF HYDROLYTIC PRODUCTS FROM REFINED PALM OIL WITH IMMOBILIZED ENZYMES AT

MODIFIED CARBON ELECTRODES

By

Zahraa A. Jarjes

Thesis submitted in fulfillment of the requirements for the degree of

Doctor of Philosophy

July 2012

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Saya isytiharkan bahawa kandungan yang dibentangkan di dalam tesis ini adalah hasil kerja saya sendiri dan telah dijalankan di Universiti Sains Malaysia kecuali dimaklumkan sebaliknya. Tesis ini juga tidak pernah diserahkan untuk ijazah yang lain sebelum ini.

I declare that the content which is presented in this thesis is my own work which was done at Universiti Sains Malaysia unless informed otherwise. The thesis has not been previously submitted for any other degree.

Disaksikan oleh:

Witnessed by:

____________________________

Tandatangan calon:

Signature of student:

________________________________

Tandatangan Penyelia:

Signature of Supervisor:

Nama calon:

ZAHRAA A. JARJES

Name of student:

K/P / Passport No.:

A5946776

Nama Penyelia:

PROF. SULIMAN BIN AB GHANI

Name of Supervisor:

K/P / Passport No.:

520807045269

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ii

DEDICATION

This work is lovingly and respectfully in honor dedicated to my

wonderful husband, Muataz Basheer Aziz for his constant

encouragement and motivation, to my beloved parents for their

inspiration and to my forever supportive family members.

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iii

ACKNOWLEDGEMENTS

First of all, I would like to thank God for the strength he has given me, I would like to express my sincere gratitude to my supervisor Professor Sulaiman Ab Ghani for his supervision, support, encouragement, patience and confidence during this research.

Additionally, I am so grateful to him for his advices, understanding and behaviors to me like a father during the difficult and unfortunate days of my life. He made me complete this thesis. I am also grateful to my cosupervisor Professor for Mohamad Razip Samian for his valuable comments.

I am indebted to Universiti Sains Malaysia and also to the Ministry of Science, Technology and Innovation (MOSTI), Malaysia for Postgraduate Incentive Research Grant 1001/PKIMIA/842038. I am also thankful to the University for the Fellowship Scheme.

I also thank all academics and technical staffs of the School of Chemical Sciences, USM for their advice and help.

My sincere thanks to all my laboratory colleagues, who have become family, and made Malaysia a place that I can call home, special thanks for my dear friend Zaini haryati Mohd. Zain for all her love, patience and support during my study work.

Finally, I am grateful to my husband muataz, my children luqman and abdul rahman, my brother in low omar, my parents, Assad and Manal, to my sisters Essraa, Zainab, Assmaa, and brothers, Mohammad, abdo allah and bellal for their endless support and love during my study as in all stages of my life.

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iv

TABLE OF CONTENTS

Page

DEDICATION ii

ACKNOWLEDGEMENTS iii

TABLE OF CONTENTS iv

LIST OF TABLES ix

LIST OF FIGURES x

LIST OF ABBREVIATION xvi

ABSTRAK xviii

ABSTRACT xx

CHAPTER ONE: INTRODUCTION AND LITERATURE REVIEW 1

1.1 Palm Oil 1

1.1.1 Products of Palm Oil 4

1.1.2 Chemistry of Palm Oil 6

1.1.3 Edible and Non-Edible Uses of Palm Oil 8

1.1.4 Palm Oil in Biodiesel 9

1.1.5 Uses of Palm Oil Biomass 10

1.1.6 Electrochemical Studies Involving Palm Oil 10

1.1.7 Hydrolysis of Palm Oil 15

1.2 Biofuel Cells 20

1.3 Enzymes as a Biocatalyst 26

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v

1.3.1 Lipoxygenase 27 1.3.2 Glycerol Dehydrogenase 28

1.3.3 Alcohol Dehydrogenase 29 1.3.4 Aldehyde Dehydrogenase 31 1.4 NADH Oxidation Issue 32 1.5 Electropolymerization of Conductive Polymers 36 1.5.1 Polymethylene Green 37 1.5.2 Polyaniline 39 1.5.3 Poly(o-phenylene diamine) 41 1.5.4 Poly(4-vinyl pyridine) 43 1.5.5 Polypyrrole 45 1.6 Enzyme Immobilization 47 1.7 Enzyme Entrapment in Nafion 49

1.8 Electrochemical Characterization Techniques 50

1.8.1 Cyclic Voltammetry 50

1.8.2 Linear Sweep Voltammetry 52

1.8.3 Amperometry 53

1.9 Objectives 53

CHAPTER TWO: EXPERIMENTAL 55 2.1 Materials 55

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vi

2.2 Equipment 56

2.3 Procedures 57

2.3.1 Assay of Lipase Activity 57

(a) Effect of pH, Temperature and Time on Lipase Activity 58

(b) Effect of Substrate and Enzyme Weights on Lipase Activity 58

2.3.2 Electropolymerization 58

2.3.3 Preparation of Electrodes 59

(a) Preparation of Lipoxygenase Electrode 59

(b) Preparation of Glycerol Dehydrogenase Electrode 60

(c) Preparation of Alcohol and Aldehyde DehydrogenaseElectrode 61

2.3.4 Preparation of Oil Emulsion for Voltammetry Studies 61

CHAPTER THREE: RESULTS AND DISCUSSION 62

3.1 Lipase Activity 62

3.1.1 Effect of Oil Loading 62

3.1.2 Effect of Enzyme Loading 63

3.1.3 Effect of pH 65

3.1.4 Effect of Temperature 66

3.1.5 Effect of Incubation Time 68

3.2 Mechanism of Lipoxygenation of Fatty Acids 69

3.3 Optimization of the Experimental Conditions 73 3.3.1 Study of Lipase Dependence 73

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vii

3.3.2 Effect of Buffer 75

3.3.3 Effect of pH 77

3.3.4 Effect of Lipoxygenase Weight 78

3.3.5 Effect of Scan Rate 79

3.3.6 Optimization of the Nafion Membrane 80

3.4 Synthesis and Characterization of Polymers on Carbon Cloth Electrodes 81

3.4.1 Electropolymerization of the Monomers 81

3.4.2 Morphological Study 89

3.4.3 Monomer Concentration 90

3.4.4 Scan Rate 93

3.5 Electrocatalysis of NADH Oxidation 100

3.5.1 Electrocatalysis of NADH by PMG 100

3.5.2 Electrocatalysis of NADH by PANI 102

3.5.3 Electrocatalysis of NADH by PoPD 104

3.5.4 Electrocatalysis of NADH by P4VP 106

3.5.5 Electrocatalysis of NADH by PPy 107

3.6 Glycerol oxidation 108 3.6.1 Glycerol Oxidation by Unmodified Electrode GDH 110

3.6.2 Glycerol Oxidation by GDH/PMG Modified Electrode 111

3.6.3 Glycerol Oxidation by GDH/PANI Modified Electrode 113

3.6.4 Glycerol Oxidation by GDH/PoPD Modified Electrode 114

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viii

3.6.5 Glycerol Oxidation by GDH/P4VP Modified Electrode 115

3.6.6 Glycerol Oxidation by GDH/PPy Modified Electrode 116

3.7 Optimization of the Experimental Conditions 119

3.7.1 Effect of NAD+ Concentration 119

3.7.2 Effect of Buffer 120

3.7.3 Effect of pH 126

3.7.4 Effect of Amount of Lipase 127

3.8 Electrochemical Oxidation of Glycerol by Multiple Dehydrogenase Enzymes Immobilized on POPD Modified Carbon Electrode 128 3.9 Optimization of the Experimental Conditions 132

3.9.1 Effect of Buffer 132

3.9.2 Effect of pH 136

3.9.3 Effect of Scan Rate 137

3.9.4 Effect of Amount of Lipase 140

3.9.5 Effect of NAD+ Concentrations 141 3.10 Amperometric Studies 142

CHAPTER FOUR: CONCLUSIONS 144

4.1 Conclusions 144 4.2 Recommendations for Future Research 147 REFERENCES 148

ADDENDUM 173

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ix

LIST OF TABLES

Page

1.1 World Palm Oil Productions from 1980 to 2008 2

1.2 World Oil and Fat Production 3

1.3 Fatty Acid Distribution of Palm Oil 6

1.4 Triacyl Glycerol Compositions (%) of Malaysian Tenera Palm Oil 7

1.5 Important Commercially Available Lipases 18

1.6 Fuels and Enzymes Used in Enzymatic Biofuel Cells 25

3.1 The Optimum Conditions of Candida Rugosa Lipase Activity to Hydrolyzed Refined Palm Oil 69 3.2 The Current Density of Lipoxygenase Immobilized in Nafion modified with Different Ammonium Salts on Carbon Electrode Recorded in Oil Emulsion, at scan rate of 100 mVs-1. 80 3.3 Electrochemical Parameters Obtained from the Electropolymerization 89 3.4 The Relationships Between Cathodic and Anodic Peak Current and The

Square Root of Scan Rate

99

3.5 Electrochemical Parameters Obtained from Electrochemical Oxidation of Glycerol Liberated from the Hydrolysis of refined Palm Oil

118

3.6 The Effect of GDH or ADH Enzyme Concentrations on The Amperometric Response of GDH/PoPD-Modified Electrode and

ADH/PoPD-Modified Electrode in Oil Emulsion in The Presence of 1 mM of NAD+

143

3.7 The effect of ADH and AldDH Enzymes Amount on the Amperometric Responses of ADH/AldDH/PoPD-Modified Carbon Cloth in

Oil Emulsion in the Presence of 1 mM of NAD+

143

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LIST OF FIGURES

Page

1.1 Cross section of a fruit 4

1.2 Composition of palm oil mesocarp 5

1.3 Variety of palm oil-based food products 8

1.4 Biomass initiatives as renewable energy 11

1.5 Hydrolysis of triacyl glycerol by lipase 16

1.6 Three dimensional structure of Candida rugosa lipase 19

1.7 A generalized scheme of an enzymatic biofuel cell, where a fuel is oxidized at the anode and molecular oxygen is reduced at the cathode 23 1.8 Electrochemical oxidation reaction of NADH 32

1.9 Basic catalytic functionalities of many organic 2-electron-proton acceptors efficient for catalytic NADH oxidation (a-f). Structural formulae of some commonly used mediators for catalytic NADH oxidation: (g) Meldola blue (p- phenylene diimine). (h) N-methyl phenazinium (o-phenylene diimine), (i) (tetracyanoquinodimethane, (j) (tetrathiofulvalene) . 34 1.10 Chemical Structure of Methylene Green 37

1.11 Chemical structure of polyaniline 39

1.12 (a) PANI like and (b) Phenazine Like Structure of PoPD 41

1.13 Chemical structure of 4VP monomer 44

1.14 Chemical structure of polypyrrol 45

1.15 The chemical structure of Nafion 49

1.16 Cyclic voltammogram for a reversible redox couple 51

1.17 Linear sweep voltammetry 52

3.1 The effect of substrate weight on the activity of Candida rugosa lipase 63 3.2 The effect of enzyme weight on the activity of Candida rugosa lipase 64 3.3 The effect of pH on the activity of Candida rugosa lipase 66

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xi 3.4

The ef The effect of temperature on the activity of Candida rugosa lipase

67 3.5

The ef The effect of incubation time on the activity of Candida rugu rugosa lipase

68

3.6 Schematic mechanism of the oxygenation of lipids by lipoxygenase

71 3.7 The cyclic voltammogram of lipoxygenase-modified electrode recorded in oil

emulsion, at scan rate 100 mVs-1

72

3.8

The cl The cyclic voltammograms of lipoxygenase-modified carbon electreelectrode recorded in oil emulsion in the absence (i) and the presenpresence (ii) of lipase, scan rate 100 mVs-1

74

3.9 Log current density at lipoxygenase-modified electrode in oil emulsion at various weight of lipase (n=3)

75

3.10 The cyclic voltammograms of lipoxygenase-modified carbon electrode recorded in oil emulsion prepared using (i) sodium phosphate buffer, (ii) Potassium phosphate buffer, and (iii) phosphate buffer saline, scan rate 100 mVs-1.

76

3.11 The effect of various pH on the log current density of lipoxygenase-modified electrode

77

3.12 The effect of different weight of lipoxygenase on the log current density of lipoxygenase-modified electrode

78

3.13 Cylic voltammograms of lipoxygenase-modified electrode recorded in oil emulsion at scan rates from a to e (20, 40, 60, 100, and 150) mVs-1. Inset:

Relationships between cathodic peak current and the square root of scan rate.

79

3.14

Cyclic Cylic voltammogram obtained during electropolymerization of 0.4 mM methylene green and 0.1 M sodium nitrate in 10 mM sodis t sodium tetraborate (scan rate 50 mVs-1) and scanning up to 10 cycles

82

3.15

Cycli Cylic voltammogram obtained during electropolymerization of 50 mM ANI in 0.2 M p-toluene sulphonic acid and 0.5 M KCl. The voltammogram is cycled between Eap −0.2 and +1V (vs.Ag/AgCl), at scan rate 50 mVs-1and scanning up to 10 cycles

84

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xii 3.16

Cyclic Cylic voltammogram obtained during electropolymerization of 50 mM in oPD in 1 M H3PO4 and 0.5 M CaCl2. The voltammogram is cycled between Eap −0.7 and +1V (vs.Ag/AgCl), at scan rate 100 mVs-1 and scanning up to 10 cycles

86

3.17 Cyclic voltammogram obtained during electropolymerization of 3 mM 4VP in 0.1 M tetrabutyl ammonium perchlorate in acetonitrile, pH 3.0. The voltammogram is cycled between Eap −0.1 and +1V (vs.Ag/AgCl), at scan rate 50 mVs-1 and scanning up to 10 cycles.

87

3.18 Cyclic voltammogram obtained during electropolymerization of 50 mM PPy in 0.2 M p-toluene sulphonic acid and 0.5 M KCl. The

voltammogram is cycled between Eap −1 and – 0.4 V (vs.Ag/AgCl), at scan rate 50 mVs-1 and scanning up to 10 cycles

88

3.19 SEM micrographs of (a) PMG (b) PANI (c) PoPD (d) P4VP and (e) PPy 91 3.20 The effect of monomer concentration on the oxidation current the density of

electropolymerization of (a) PMG, (b) PANI and (c) PoPD

92

3.21

Cyclic Cyclic voltammograms of PMG at different scan rates from a to f (20, 50, 100, 150, 200, 250 mV s-1). Inset: Relationships between cathodic and anodic peak current and the square root of scan rate

94

3.22 Cyclic voltammograms of PANI at different scan rates from a to f (20, 50, 100, 150, 200, 250 mV s-1). Inset: Relationships between cathodic and

anodic peak current and the square root of scan rate

95

3.23

Cyclic Cyclic voltammograms of PoPD at different scan rates from a to f (20, 50, 100, 150, 200, 250 mV s-1). Inset: Relationships between cathodic and anodic peak current and the square root of scan rate.

96

3.24

Cyclic Cyclic voltammograms of P4VP at different scan rates from a to f (20, 50, 100, 150, 200, 250 mV s-1). Inset: Relationships between cathodic and anodic peak current and the square root of scan rate

97

3.25

Cyclic Cyclic voltammograms of PPy at different scan rates from a to f (20, 50, 100, 150, 200, 250 mV s-1). Inset: Relationships between cathodic and anodic peak current and the square root of scan rate.

98

3.26 Cyclic voltammograms for the PMG recorded in phosphate buffer

solutions (pH7) in the absence (CV1) and presence of : 0.1 mM (CV2) and 0.5 mM (CV3) of NADH. At scan rate 50 mV s-1.

101

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xiii 3.27

Cyclic Cyclic voltammograms for the PANI recorded in phosphate buffer solutions (pH7) in the absence (CV1) and presence of: 0.1 mM (CV2) and 0.5 mM (CV3) of NADH. At scan rate 50 mV s-1.

103

3.28 Cyclic voltammograms for the PoPD recorded in phosphate buffer solutions (pH7) in the absence (CV1) and presence of: 0.1 mM (CV2) and 0.5

mM (CV3) of NADH. At scan rate 50 mV s-1.

105

3.29 Cyclic voltammograms for the P4VP recorded in phosphate buffer solutions (pH7) in the absence (CV1) and presence of 0.5 mM (CV2) of

NADH. At scan rate 50 mV s-1

106

3.30 Cyclic voltammograms for the PPy recorded in phosphate buffer solutions (pH7) in the absence (CV1) and presence of 0.5 mM (CV2) of

NADH. At scan rate 50 mV s-1

107

3.31 Schematic reactions for the oxidation of glycerol 109

3.32 Cyclic voltammograms of GDH/unmodified electrode in (a) oil emulsion in the presence of 1 mM of NAD+ and (b) phosphate buffer solution (pH = 7). The scan rate is 100 mVs-1

110

3.33 Cyclic voltammograms of (a) GDH/unmodified electrode in oil emulsion in the presence of 1 mM of NAD+ (b) GDH/PMG modified electrode in oil emulsion in the presence of 1 mM of NAD+ (c) GDH/PMG modified electrode in phosphate buffer pH 7 and in the presence of 1 mM of NAD+

112

3.34 Cyclic voltammograms of (a) GDH/unmodified electrode in oil emulsion in the presence of 1 mM of NAD+(b) GDH/PANI modified electrode in oil emulsion in the presence of 1 mM of NAD+ (c) GDH/PANI modified electrode in phosphate buffer pH 7 and in the presence of 1 mM of NAD+

113

3.35 Cyclic voltammograms of (a) GDH/unmodified electrode in oil emulsion in the presence of 1 mM of NAD+ (b) GDH/PoPD modified electrode in oil emulsion in the presence of 1 mM of NAD+, (c) GDH/PoPD modified electrode in phosphate buffer pH 7 and in the presence of 1 mM of NAD+

114

3.36 Cyclic voltammograms of (a) GDH/ unmodified electrode in oil emulsion in the presence of 1 mM of NAD+ (b) GDH/P4VP modified electrode in oil emulsion and in the presence of 1 mM of NAD+ (c) GDH/P4VP modified electrode in phosphate buffer pH 7 and in the presence of 1 mM of NAD+

115

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3.37 Cyclic voltammograms of (a) GDH/PPy modified electrode in oil emulsion in the presence of 1 mM of NAD+(b) GDH/PPy modified electrode in phosphate buffer pH 7 and in the presence of 1 mM of NAD+

117

3.38 The oxidative current density at GDH/polymers modified carbon cloth electrode in oil emulsion at various concentrations of NAD+, (n = 3)

120

3.39 Cyclic voltammograms of GDH/PMG modified electrode in oil emulsion and in the presence of 1 mM of NAD+ prepared using (a) sodium phosphate buffer, (b) phosphate buffer saline (PBS), and (c) potassium phosphate buffer. The scan rate is 50 mVs-1.

122

3.40 Cyclic voltammograms of GDH/PANI modified electrode in oil emulsion in the presence of 1 mM of NAD+ prepared using (a) sodium phosphate buffer, (b) phosphate buffer saline (PBS), and (c) potassium phosphate buffer. The scan rate is 50 mVs-1.

123

3.41

Cyclic Cyclic voltammograms of GDH/PoPD modified electrode in oil emulsion in the presence of 1 mM of NAD+ prepared using (a) sodium phosphate buffer, (b) phosphate buffer saline (PBS), and (c) potassium phosphate buffer. The scan rate is 50 mVs-1.

124

3.42

Cyclic Cyclic voltammograms of GDH/P4VP modified electrode in oil emulsion in the presence of 1 mM of NAD+ prepared using (a) sodium phosphate buffer, (b) phosphate buffer saline (PBS), and (c) potassium phosphate buffer. The scan rate is 50 mVs-1.

125

3.43 The oxidative current density at GDH/polymers modified carbon cloth electrode in oil emulsion at various pH, (n = 3).

126

3.44 The oxidative current density at GDH/polymers modified carbon cloth electrode in oil emulsion at various weight of lipase, (n = 3)

127

3.45 Cyclic voltammograms of (a) PoPD-modified electrode, (b) ADH/PoPD- modified electrode and (c) ADH/AldDH/PoPD-modified electrode, all recorded in oil emulsion in the presence of 1 mM of NAD+. The scan rate is 50 mVs-1.

130

3.46 Cyclic voltammograms of (a) PoPD-modified electrode (b) GDH/PoPD- modified electrode, all recorded in oil emulsion in the presence of 1 mM of NAD+. The scan rate is 50 mVs-1.

131

3.47 Linear sweep voltammograms of GDH/PoPD-modified electrode in oil emulsion and in the presence of 1 mM of NAD+ prepared using (a) potassium phosphate buffer, (b) sodium phosphate buffer and (c) phosphate buffer saline (PBS). The scan rate is 50 mVs-1.

133

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3.48 Linear sweep voltammograms of ADH/PoPD-modified electrode in oil emulsion and in the presence of 1 mM of NAD+ prepared using (a) potassium phosphate buffer, (b) sodium phosphate buffer and (c) phosphate buffer saline (PBS). The scan rate is 50 mVs-1.

134

3.49 Linear sweep voltammograms of ADH/AldDH/PoPD -modified electrode in oil emulsion and in the presence of 1 mM of NAD+ prepared using (a) potassium phosphate buffer, (b) sodium phosphate buffer and (c) phosphate buffer saline (PBS). The scan rate is 50 mVs-1.

135

3.50 The oxidative current density at GDH, ADH, and ADH/AldDH/PoPD-modified carbon cloth electrode in oil emulsion in the presence of 1 mM of NAD+ at various pH (n = 3)

136

3.51 Cyclic voltammograms of GDH/PoPD-modified carbon cloth electrode in oil emulsion in the presence of 1 mM of NAD+ at scan rates from a to f (20, 50, 100, 150, 200, 250 mV s-1). (Inset: The relationship between cathodic and anodic peak current and the square root of scan rate)

137

3.52 Cyclic voltammograms of ADH/PoPD-modified carbon cloth electrode in oil emulsion in the presence of 1 mM of NAD+ at scan rates from a to f (20, 50, 100, 150, 200, 250 mV s-1). (Inset: The relationship between cathodic and anodic peak current and the square root of scan rate)

138

3.53 Cyclic voltammograms of ADH-AldDH/PoPD -modified carbon cloth electrode in oil emulsion in the presence of 1 mM of NAD+ at scan rates from a to f (20, 50, 100, 150, 200, 250 mV s-1). (Inset:The relationship between cathodic and anodic peak current and the square root of scan rate)

139

3.54 The oxidative current density at GDH, ADH, and ADH/AldDH/PoPD- modified carbon cloth electrode in oil emulsion in the presence of 1 mM of NAD+ at various weight of lipase (n = 3)

140

3.55 The oxidative current density at GDH, ADH, and ADH/AldDH/PoPD- modified carbon cloth electrode in oil emulsion at various concentrations of NAD+ (n = 3)

141

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xvi

LIST OF ABBREVIATIONS

ADH Alcohol Dehydrogenase

AldDH Aldehyde Dehydrogenase º C Degree Celsius

CVs Cyclic Voltammograms

DTAB Dodecyltrimethylammonium bromide e Electron

Eap Applied Potential Epa Anodic Peak Potential Epc Cathodic Peak Potential

g Gram

GDH Glycerol Dehydrogenase

h Hour

Ipa Anodic Peak Current Ipc Cathodic Peak Current j Current Density

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xvii kDa Kilo Dalton

LSV Linear Sweep Voltammetry NADH β-nicotinamide adenine dinucleotide PANI Polyaniline

PMG Polymethylene green PoPD Poly(o-phenylene diamine) PPy Polypyrrole PQQ Pyrroloquinoline quinine

PTSA p-toluene sulphonic acid PVA Polyvinyl alcohol

P4VP Poly(4-vinyl pyridine) SEM Scanning Electron Microscopy

SLO Soybean Lipoxygenase

TGs Triglycerides V Volt

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xviii

KAJIAN ELEKTROKIMIA HASIL HIDROLISIS MINYAK SAWIT TERTAPIS DENGAN ENZIM TERIMOBILKAN PADA ELEKTROD KARBON

TERUBAHSUAI ABSTRAK

Kerja ini menerangkan kesan pH, suhu, masa pengeraman, kepekatan substrat dan berat enzim pada hidrolisis minyak sawit tertapis menggunakan lipase komersial dari yis Candida rugosa. keputusan menunjukkan bahawa hidrolisis adalah optimum pada pH 7.5, suhu 37° C, masa eraman 60 minit dan berat enzim dan substrat 0.1 dan 2 g, masing-masing. Asid lemak yang dibebaskan telah digunakan sebagai substrat untuk lipoksigenase yang dipegunkan di permukaan membran Nafion di atas elektrod karbon

yang telah diubah suai. Arus yang dijana telah dikaji dengan voltammetri berkitar.

Didapati penampan kalium pada pH 7 dan 0.4 mg lipoksigenase adalah penting untuk menghasilkan ketumpatan arus yang lebih tinggi.

Pengelektrooksidaan gliserol (yang diperolehi pada keadaan optimum hidrolisis minyak sawit tertapis, menggunakan lipase komersial) bersama nikotinamida adenina dinukleotida (NAD+) menggunakan enzim gliserol dehidrogenase yang dipegunkan dalam membran Nafion yang telah dimodifikasi dengan ammonium pada elektrod kain karbon terubahsuai. Disebabkan oleh voltan lampau yang besar yang dihadapi untuk pengoksidaan NADH pada elektrod, lima jenis polimer iaitu polimetilena hijau (PMG), polianilina (PANI), poli(orto-fenilena diamina) (PoPD), poli(4-vinil piridina) (4-VP), dan polipirola (PPy) telah digunakan untuk menjana semula NAD+ dan mengulang-alik elektron daripada NADH kepada elektrod. Secara amnya, proses redoks gliserol adalah quasi-berbalik dalam julat potensi -0.5 sehingga +0.6 V lawan Ag/AgCl. Kesan pH, kepekatan NAD+, dan berat lipase telah dikaji untuk mencapai ketumpatan arus yang

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xix

lebih tinggi. Pengoksidaan elektrokimia gliserol telah juga dikaji menggunakan dua sistem enzim berlainan, iaitu 1) sistem monoenzim, yang melibatkan alkohol dehidrogenase dipegunkan dalam lapisan Nafion pada permukaan elektrod kain karbon yang diubahsuai dengan PoPD, dan 2) sistem bienzim yang melibatkan alkohol dehidrogenase dan aldehid dehidrogenase yang disekatgerak pada elektrod seperti yang diterangkan tadi. Pembangunan sistem kedua ini bertujuan untuk mengkaji kemungkinan pengoksidaan gliserol berperingkat. Kajian voltammetri ke atas julat potensi -1.5 sehingga +0.1 V berlawan Ag/AgCl, telah menunjukkan bahawa elektrod ini memaparkan, kebanyakannya proses elektrod jenis quasi-berbalik untuk pengoksidaan gliserol. Pelbagai parameter juga telah dikaji untuk mencari keadaan yang optimum.

Pada amnya, ketumpatan arus oksidatif maksimum semasa untuk kesemua elektrod adalah pada pH 7 dengan berat lipase 0.1 – 0.15 g dan kepekatan NAD+ 1 - 3 mM.

Penemuan ini dijangka berguna dalam pembentukan katod dan anod dalam sel biofuel menggunakan asid lemak dan gliserol daripada minyak kelapa sawit bertapis.

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ELECTROCHEMICAL STUDIES OF HYDROLYTIC PRODUCTS FROM REFINED PALM OIL WITH IMMOBILIZED ENZYMES AT MODIFIED

CARBON ELECTRODES

ABSTRACT

This work describes the effect of pH, temperature, incubation time, and substrate and enzyme weights on the hydrolysis of refined palm oil using the commercial lipase from the yeast Candida rugosa. It was found that the hydrolysis was optimum at pH 7.5, temperature 37 °C, incubation time 60 min and the enzyme and substrate weights of 0.1 and 2 g, respectively. The released fatty acids were used as a substrate for lipoxygenase immobilized on modified Nafion membrane carbon electrode and the current generated was studied by cyclic voltammetry. Potassium buffer at pH 7 and 0.4 mg of lipoxygenase was found to be crucial in order to produce a higher current density.

The electrooxidation of glycerol (obtained at optimum conditions of hydrolysis of refined palm oil using commercial lipase) in the presence of nicotinamide adenine dinucleotide (NAD+) using glycerol dehydrogenase enzyme immobilized in ammonium modified Nafion membrane on polymer modified carbon cloth electrodes. Due to the large overvoltage encountered for NADH oxidation at the electrode, five conducting polymers viz. polymethylene green (PMG), polyaniline (PANI), poly(ortho-phenylene diamine) (PoPD), poly(4-vinyl pyridine) (P4VP), and polypyrrole (PPy) were used to regenerate NAD+ and to shuttle electrons from the NADH to the electrode. In general, the redox processes of glycerol were of quasi-reversible over potential ranges of -0.5 to

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+0.6 V vs Ag/AgCl. The effect of pH, concentrations of NAD+, and weight of lipase has been studied to achieve a higher current density. The electrochemical oxidation of glycerol was also studied by another two enzymes system, monoenzyme system, alcohol dehydrogenase immobilized within Nafion layer on the surface of PoPD modified carbon cloth electrode, and a bienzyme system of alcohol dehydrogenase and aldehyde dehydrogenase immobilized similarly on the electrode as described earlier. The latter was aimed for a possible study of multi-step oxidation of glycerol. Voltammetric studies over potential ranges of – 1.5 to + 0.1 V vs. Ag/AgCl, have shown that these electrodes displayed mostly quasi-reversible type of electrode processes for the oxidation of glycerol. Various parameters have also been examined for experimental optimum conditions. In general, the maximum oxidative current density for all electrodes was obtained at pH 7 with weight of lipase 0.1- 0.15 g and concentration of NAD+ 1 - 3 mM.

The findings were expected to be useful in the fabrication of cathode and anodes in a biofuel cell using fatty acids and glycerol from refined palm oil.

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1

CHAPTER ONE

INTRODUCTION AND LITERATURE REVIEW

1.1 Palm Oil

In 1434, a Portugese sailor, Gil Eannes first reported about oil palms (Elaeis guineensis). Oil palms are grown mainly in the western part of Africa, Indonesia, Malaysia, and recently in Brazil and Colombia. Oil palm trees grow up to 20 m in height and they grow best at temperature of 24-27oC. Oil palm trees need a humid climate and the cultivated oil palm carry fruit from their fourth year onward and can be harvested for 40-50 years. Malaysia is blessed with good weather conditions which prevail throughout the year make it useful for palm oil cultivation. The first commercial oil palm estate in Malaysia was set up in 1917 at Tennamaran Estate, Selangor. In 1960s, oil palms were commercially cultivated in large scale to avoid over dependence on natural rubber which is major product during earlier years. Since then, palm oil industry has expended fast and has emerged as the most remunerative agricultural commodity, overtaking natural rubber (Norazlan, 2006).

Palm oil is the second most traded vegetable oil crop in the world after soy, Malaysia is among the largest producers and exporters of palm oil in the world, accounting for 52% or 26.3 million tonnes of the total world oils and fats exports in year 2006 (Sumathi et al., 2008). Table 1.1 shows the major palm oil producers, while Table 1.2 details world production of oils and fats.

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Table 1.1 World Palm Oil Productions from 1980 to 2008 (Ong et al., 2011) Palm oil output / 1000 metric tones

1980 1985 1990 1995 2000 2005 2008

Malaysia 2576 4133 6088 8123 10,842 14,962 17,735

Indonesia 691 1243 2413 4220 7050 14,070 19,100

Nigeria 433 386 580 660 740 800 860

Thailand - - 232 354 525 685 1160

Colombia - - 226 388 524 661 800

Papua New Guinea - - 145 223 336 310 400

Cote D‟Ivoire - - 270 285 278 260 330

Brazil 12 29 66 75 108 160 420

Others 875 1041 1000 5994 5191 1826 2100

World total 4587 6832 11,020 20,322 25,594 33,733 42,904

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Table 1.2 World Oil and Fat Production (Ong et al., 2011) Volume / 1000 metric tones

1980 1985 1990 1995 2000 2005 2008

Palm oil 4543 6832 11,020 20,322 25,594 33,733 40,200

Soyabean oil 13,382 13,974 16,097 15,119 21,743 33,575 35,700

Rapeseed oil 3478 6066 8160 10,936 14,496 16,205 19,900

Sunflower seed oil 5024 6564 7869 7003 9808 9661 10,800

Tallow & grease 6283 6518 6813 7013 8071 8211 8585

Lard 4691 4989 5509 5141 6580 7568 7740

Butter fat 5746 6315 6500 4834 5829 6665 7123

Cottonseed oil 2992 3942 3782 3312 3815 4989 4400

Groundnut oil 2864 3575 3897 4325 4382 4523 4445

Palm kernel oil 571 868 1450 1877 2620 3975 5300

Coconut oil 2716 2627 3387 3253 3147 3257 3500

Olive oil 1701 1796 1855 1863 2513 2916 3000

Others 3695 4430 4552 4618 4995 5100 5840

World total 57,686 68,496 80,891 89,615 113,591 140,378 160,471

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4 1.1.1 Products of Palm Oil

There are two main products produced by the oil palm fruit namely crude palm oil obtained from the mesocarp and crude palm kernel oil obtained from the endosperm (kernel). Figure 1.1 shows a cross section of palm oil fruit. The main by-product and wastes produced from the processing of palm oil are the empty fruit bunches, palm oil mill effluent, sterilizer condensate, palm fibre, and palm kernel shell (Yusoff, 2006).

Figure 1.1 Cross section of a fruit.

Mesocarp

Kernal

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Mesorcarp accounts for about 60% of the total composition of oil palm fruit.

Figure 1.2 shows the composition of mesocarp where the oil accounts for 39% of the overall composition. Crude palm oil is obtained from the mesocarp part of oil palm fruit after undergoing several processes such as sterilization, stripping, extraction, and purification (Norazlan, 2006).

Figure1.2 Composition of palm oil mesocarp (Norazlan, 2006)

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6 1.1.2 Chemistry of Palm Oil

Crude palm oil is semi solid at room temperature; the oil is bright orange-red in colour due to its high content of carotene. Like all oils triglycerides (TGs) are the major constituents of palm oil. Each glycerol molecule is esterified with three fatty acids. The minor components of palm oil are phosphatides, sterols, carotenoids, tocopherols, tocotrienols, and trace metals. The major fatty acids in palm oil are myristic, palmitic, stearic, oleic, and linoleic (Sundram et al., 2003). The fatty acid distribution of the palm oil is given in Table 1.3.

Table 1.3 Fatty Acid Distribution of Palm Oil (Noor et al., 2003)

Name Fatty acid

chain length

% w/w Formulae Molecular

weight

Lauric C12:0 0.2 CH3(CH2)10COOH 200.31

Myristic C14:0 1.1 CH3(CH2)12COOH 228.36

Palmitic C16:0 44.0 CH3(CH2)14COOH 256.42

Stearic C18:0 4.5 CH3(CH2)16COOH 284.47

Oleic C18:1 39.2 C17H33COOH 282.44

Linoleic C18:2 10.1 C17H31COOH 280.43

Others - 0.9 - -

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Palm oil contains saturated and unsaturated fatty acids in about equal amounts.

Most of the fatty acids are present as TGs. There are 7% to 10% of saturated TGs, mainly tripalmitin. The fully unsaturated TGs represent 6 to 12%. The Sn-2 position has specificity for unsaturated fatty acids. Therefore, more than 85% of the unsaturated fatty acids are located in the Sn-2 position of the glycerol molecule. The triacylglycerols in palm oil partially define most of the physical characteristics of the palm oil such as melting point and crystallisation behaviour (Kifli, 1981). Table 1.4 shows the percentage distribution of individual TGs of palm oil.

Table 1.4 Triacyl Glycerol Compositions (%) of Malaysian Tenera Palm Oil (Kifli, 1981)

No Double Bond 1 Double Bond 2 Double Bonds 3 Double Bonds > 4 Double Bonds

MPP 0.29 MOP 0.83 MLP 0.26 MLO 0.14 PLL 1.08 PMP 0.22 MPO 0.15 MOO 0.43 PLO 6.59 OLO 1.71

PPP 6.91 POP 20.02 PLP 6.36 POL 3.39 OOL 1.76 PPS 1.21 POS 3.50 PLS 1.11 SLO 0.60 OLL 0.56

PMO 0.22 PPL 1.17 SOL 0.30 LOL 0.14 PPO 7.16 OSL 0.11 OOO 5.38

PSO 0.68 SPL 0.10 OPL 0.61 SOS 0.15 POO 20.54 MOL -

SPO 0.63 SOO 1.81 OPO 1.86

OSO 0.18 PSL -

Others 0.16 0.34 0.19 0.15 0.22 Total 9.57 33.68 34.12 17.16 5.47 Note: M, myristic; P, palmitic, S, stearic; O, oleic; L, linoleic

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8 1.1.3 Edible and Non-Edible Uses of Palm Oil

Palm oil is used in both edible and non-edible applications and 90 percent of palm oil and its products are used for edible purposes. Palm oil is used extensively in food preparation and manufacturing mainly as cooking and frying oils, or shortenings and margarine. Figure 1.3 provides examples on a number of palm-based food applications. The remaining 10% of palm oil and its products are used for non-edible applications, mainly in the soap and in oleochemical industries (Sarmidi et al., 2009).

Figure 1.3 Variety of palm oil-based food products (Hai, 2002)

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9 1.1.4 Palm Oil in Biodiesel

Biodiesel is a clean-burning diesel fuel produced from vegetable oils, animal fats, or grease. Biodiesel as a fuel gives much lower toxic air emissions than fossil diesel. In addition, it gives cleaner burning and has less sulphur content, and thus reducing emissions. Due to its origin from attractive resources, it is expected to compete with petroleum products in the future (Hayyan et al., 2010).

Biodiesel can be chemically defined as a methyl ester which is prepared from triglycerides in vegetable oils by transesterification with methanol (Meher et al., 2006).

Biodiesel has been extensively researched using many different types of vegetable oils.

Among these sources, palm oil is the cheapest vegetable oil due to its higher yield of approximately 5000 kg per hectare, compared with other vegetable oils, the maximum of which, for coconut oil is around 2250 kg per hectare. Therefore, it would be economically instinctive to believe palm oil as the feedstock for biodiesel production (Al-Zuhair et al., 2007).

In Malaysia, palm oil is used in the production of biodiesel (palm oil methylester or palm oil diesel) for buses and cars due to the absence of sulphur and nitrogen.

Furthermore, the use of palm oil is also advocated because it is assumed to dramatically reduce CO2 emissions (Reijnders & Huijbregts, 2008).

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10 1.1.5 Uses of Palm Oil Biomass

Huge quantities of biomass by-products, such as empty fruit bunches fibers, shells, fronds, and trunks are produced. These biomasses can be converted into many value added products. Palm oil biomasses can be transformed into three types of biomass energies, which are bio products, biofuels, and bio power (Kalinci et al., 2011).

Figure 1.4 shows the scope of biomass initiatives. Biofuels are used in the transportation sector. There are three types of biofuels that are bioethanol, bio diesel, and bio methanol.

Palm oil based biofuels are „„environmentally friendly‟‟ compared with fossil fuels which could cause the damage to the environment through emission of large quantities of green-house gases and pollutants (Chew & Bhatia, 2008). Biopower is the use of biomass to generate electricity. There are six major types of biopower systems: Direct firing, cofiring, gasification, pyrolysis, anaerobic digestion, and small modular systems.

Most of the bio power plants use direct fired systems. In addition, gas and liquid fuels can be produced from biomass through pyrolysis. In pyrolysis biomass is heated in the absence of oxygen. The biomass then turns into a liquid called pyrolysis oil, which burns like petroleum to generate electricity (Bazmia et al., 2010).

1.1.6 Electrochemical Studies Involving Palm Oil

The usage of palm oil in the electrochemical field has been investigated by different authors. Different electrochemical techniques have been used to treat and electroanalyze palm oil such as voltammetry, potentiometry, and chronopotentiometry, for different objectives. One of these objectives is to generate electricity using fuel cells.

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Figure 1.4 Biomass initiatives as renewable energy (Sumathi, et al., 2008) Bio-products

Materials Adsorbent

Products Fuels Chemicals

Materials Heat & Power Bio-fuel

Enzymatic Hydrolysis Lignin product Biomass

Residue Harvesting Energy Crops

Bio-power Pyrolysis Gasification

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An electrochemical determination of vitamin E in palm oil has been reported (Atuma, 1975). The method involves saponification, extraction of the unsaponifiable material and direct voltammetric determination. Two indicator electrodes (glassy carbon and carbon paste) have been employed in this work for comparison purposes. The most important advantage of this method is its precision and the rapidity of analysis. The determination of the copper content of crude and hydrogenated palm oils has been investigated using a copper(II) ion-selective electrode with the direct potentiometric method. The method does not suffer from matrix effects and the reproducibility is reasonable. The apparatus assembly is simple, the running cost is low, and the capital cost is less for this method (Fung & Fung, 1978).

The determination of copper and lead in palm oil has also been achieved by stripping chronopotentiometry technique. The metal ions were concentrated as their amalgams on the glassy carbon surface of a working electrode that was coated with a thin mercury film. An ultrasonic bath was used for the extraction of copper and lead from the oil samples. The concentration of trace metals is an important criterion for the assessment of oil qualities with regard to freshness, maintenance properties, storage, and their influence on human nutrition and health (Cypriano et al., 2008).

The electroanalytical possibilities of using Nafion–coated probes in cooking palm oil without any form of sample modification or dilution by conducting solvents has been investigated (Surareungchai & Kasiwat, 2000).

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The electrochemical probe consists of co planar platinum working and counter electrodes, and a silver quasi reference electrode, was coated with a film of Nafion. It was demonstrated that the probe could be used to perform direct analysis of the antioxidant tert-butyl hydroquinone in cooking palm oils using differential pulse voltammetry.

An electro-Fenton pretreatment and biological oxidation has been used for the removal of recalcitrant contaminants present in palm oil effluent obtained from a food processing industry. Low molecular weight fatty acids were obtained at the end of electro-Fenton pretreatment and it was further degraded to CO2 by biological oxidation.

Electro-Fenton enables the successful increment of biodegradability index of the wastewater and plays an important role in wastewater management (Babu et al., 2010).

Furthermore, amperometric enzyme electrodes have been constructed by adsorbing anionic royal palm tree peroxidase on spectroscopic graphite electrodes. The resulting H2O2-sensitive biosensors were characterized both in a flow injection system and in batch mode to evaluate its main bioelectrochemical parameters. The results indicate a uniquely superior characteristic of the biosensors, which due to the high stability of this enzyme in presence of H2O2 with an extremely high thermal and pH- stability (Alpeeva et al., 2005).

An optical thin-film biosensor chip-based analytical technique has been proven to be a rapid, simple, specific, and sensitive method suitable for the detection of trace amounts of species-specific DNA from palm oil and has been demonstrated to be effective with other different vegetable oils (Bai et al., 2011).

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The multiwalled carbon nanotubes were synthesised by utilising palm oil as organic carbon sources at 700°C and then inserted electrochemically with Lithium. The charge/discharge test of Lithium/ multiwalled carbon nanotubes cells was performed under a galvanostatic mode. The irreversible capacity of the multiwalled carbon nanotubes was found to be relatively large due to formation of the passivation layer on the tube surface (Kudin et al., 2009).

The supercritical water gasification of wet biomass from empty fruit bunch palm as a hydrogen production has been used to generate electricity using fuel cell.

Supercritical water behaves like a non - polar organic single-phase solvent. It has been applied in the production of hydrogen rich fuel gas from wet biomass. The empty fruit bunch which has high moisture is just waste produced from a palm oil factory. The hydrogen produced by this process has been utilized to generate electricity using fuel cell which is an electrochemical device that produces electricity from a combined chemical reaction and electrical charge transport. Therefore, it offers potential benefits, efficiency, no emissions, and greenhouse gas reduction (Utomo et al., 2006).

Palm oil biofuel cell has never been tried before but other edible oils (soy bean oil) biofuel cell have been reported (Kerr & Minteer, 2008).

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15 1.1.7 Hydrolysis of Palm Oil

Triglyceride the main component of natural oil or fat is converted stepwise into diacylglycerol, monoacylglycerol, and glycerol by hydrolysis accompanied with the liberation of fatty acid at each step (Beisson et al., 2000). Hydrolysis of oil and fat is an important industrial operation; the products glycerol and fatty acids are widely used as raw materials in food, cosmetic, and pharmaceutical industries (Snape & Nakajima, 1996).

The Colgate-Emery process has been used for the hydrolysis of oil, in which pressurized steam at high temperature has been employed to hydrolyze ester bonds (Bamebey & Brown, 1948). This process not only consumed energy, but also affects the properties of fatty acids in the triacylglycerol mixtures, produce undesirable compounds such as ketones and hydrocarbons, and also undesirable colour impurities which have to be separated from the products (Al-Zuhair et al., 2003).

Recently, enzymatic hydrolysis of triglycerides has gained increasing attention, which can be carried out at room temperature and atmospheric pressure making it energy efficient in comparison with the steam splitting process (Murty et al., 2002).

Furthermore, enzymes are biodegradable and consequently are less polluting than chemical catalysts (Cavalcanti-Oliveira et al., 2011). Lipases are a class of hydrolyses enzymes that are primarily responsible for the hydrolysis of acylglycerides (Sharma et al., 2001).

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Lipases (EC 3.1.1.3) are serine hydrolases that do not require any cofactors (Singh et al., 2008). It catalysed reactions that take place at the interface between the aqueous phase containing the enzyme and the oil phase (Lee et al., 2006). Lipase is a polypeptide chain folded into two domains, the C terminal domain and the N-terminal domain which contain the active site with a hydrophobic tunnel from the catalytic serine to the surface that can accommodate a long fatty acid chain. The catalytic mechanism of lipases is centred on the active site serine. The nucleophilic oxygen of the active site serine forms a tetrahedral hemiacetal intermediate with the triacylglyceride. The ester bond of the hemiacetal is hydrolysed and the diacylglyceride is released. The active site serine acyl ester subsequently reacts with a water molecule. The acyl enzyme is then cleaved and the fatty acid is dissociated (Öztürk, 2001). A typical reaction catalyzed by lipases is shown in Figure 1.5.

Figure 1.5 Hydrolysis of triacylglycerol by lipase

Fatty acids Glycerol

Triacyl glycerol

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The exclusive features of lipases such as substrate specificity, regio-specificity and chiral selectivity drew great attention in both physiological and biotechnological aspects. Their main applications are in organic chemical processing, detergent formulations, synthesis of biosurfactants, the oleochemical industry, the dairy industry, the agrochemical industry, paper manufacturing, nutrition, cosmetics, and pharmaceutical processing (Salihu et al., 2011).

Lipases can be derived from animal, bacterial, and fungal sources so they all tend to have similar three-dimensional structures (Saxena et al., 2003). Many pure lipases, often obtained by recombinant technology and be purchased from enzyme suppliers.

Table 1.5 summarizes commercially available lipases.

To date the lipase from yeast Candida rugosa is the most industrially used enzymes due to its high activity both in hydrolysis as well as synthesis, versatile catalytic reactions and broad specificities (Ratledge & Tan, 1990; Redondo et al., 1995).

The three-dimensional structure of Candida rugosa lipase (Öztürk, 2001) can be seen in Figure 1.6.

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Table 1.5 Important Commercially Available Lipases (Doucet, 2007)

Origin Code Applications

mammalian origin

human pancreatic lipase HPL human gastric lipase HGL

porcine pancreatic lipase PPL organic synthesis, digestive

aid guinea pig pancreatic lipase GPL-RP2

fungal origin

Candida rugosa CRL organic synthesis

Candida antarctica B CAL-B organic synthesis

Rhizomucor miehei RML cheese manufacturing

Aspergillus oryzae AOL cheese manufacturing

Penicillium camembertii PEL oleochemistry

Rhizopus delemar RDL oleochemistry

Rhizopus oryzae

(phospholipase A1 activity)

ROL oleochemistry

Rhizopus arrhizus RAL oleochemistry

bacterial origin

Pseudomonas glumae PGL detergent enzyme, organic

Burkholderia cepacia PCL/BCL synthesis

Pseudomonas mendocina PML organic synthesis

Chromobacterium viscosum CVL detergents

Bacillus thermocatenulatus BTL-2 organic synthesis

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Figure 1.6 Three dimensional structure of Candida rugosa lipase (Öztürk, 2001)

The hydrolysis of palm oil, palm olein, and palm stearin by the commercial lipase from Candida rugosa has been reported (Khor et al., 1986); palm oil and palm olein were found to be hydrolyzed at the same rate under the same conditions, whereas palm stearin was hydrolyzed much more slowly. The kinetics of Candida rugosa lipase hydrolysis of palm oil in lecithin/isooctane reverse micellar system has been also studied (Knezevic et al., 1998), the reaction was found to obey Michaelis-Menten kinetics for the initial conditions. Five commercial lipases were compared for their ability to

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hydrolyze palm olein in organic solvent in a two-phase system (H-Kittikun et al., 2000).

The results indicated that lipase from (Candida rugosa) showed the highest specific activity and achieved nearly complete hydrolysis.

The effects of enzymatic hydrolysis on crude palm olein by lipase from Candida rugosa have been also investigated (You & Baharin, 2006). Crude palm olein was hydrolyzed first to produce an oil rich in free fatty acids, then a comparison has been also made between crude palm olein and hydrolyzed crude palm olein for properties such as melting point, percentage of free fatty acids produced and viscosity. Hydrolyzed crude palm olein is found to be preferred in this hydrolysis process.

In this study we described the optimum conditions for the hydrolysis of refined palm oil using commercial Candida rugosa lipase. The effect of oil loading, enzyme loading, pH, temperature, and incubation time was investigated. The liberated fatty acids and glycerol from the hydrolysis of refined palm oil was used then as a substrate for different enzymes immobilized in ammonium modified Nafion membrane on the carbon electrode for potential biofuel cell applications.

1.2 Biofuel Cells

The energy-demanding bioelectronic devices require small power sources that are able to maintain operation over long periods of time at natural conditions.

Miniaturized biofuel cells in the future can be used as alternative energy supply sources for nano-microelectronic devices and biosensors (Ramanavicius et al., 2005). The concept of biofuel cells has been recognized for almost one century since the first

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microbial biofuel cell was demonstrated in 1912. In the 1960s, NASA showed great interest in power generation from human wastes on the space shuttles (Kim et al., 2006).

The production of electrical power from low-cost biofuels is an essential challenge in the energetics, since biofuels are renewable, sustainable, and reduce the demand for common fuel sources (Pizzariello et al., 2002). The development of ever more miniaturized and complicated implantable biomedical devices represent the driving force for the design of small, reliable, biocompatible, and low power source systems biofuel cells are being developed for such applications (Barrière et al., 2006). Biofuel cells are able of utilizing naturally existing biomass as fuel. They are an important substitute to conventional fuel cells and batteries that are besieged by non renewability, non implantability, size/weight, operating conditions (high temperature, acidity, and toxicity), waste issues, and logistics (Bilge et al., 2009).

Biofuel cells belong to a special class of fuel cells where biocatalysts such as microorganisms or enzymes are employed instead of metallic inorganic catalysts (Sokic- Lazic & Minteer, 2008). Biofuel cells function by coupling two reactions, the oxidation of a biofuel (by microbes or enzymes) at the anode and the reduction of molecular oxygen to water at the cathode. The electrons travel from the anode to the cathode of the biofuel cell, where they are finally accepted by molecular oxygen. The availability of electrons and protons at the cathode foster the reduction of molecular oxygen to water.

For an efficient system, the only byproducts of a biofuel cell should be carbon dioxide and water. This is an attractive feature of biofuel cell since the byproducts are non toxic.

Biofuel cells would, therefore, be more environmentally friendly (Justin, 2001).

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Biofuel cells reported in the literature are within two distinct categories: Biofuel cells that utilize the chemical pathways of living cells (microbial fuel cells) and those that employ isolated enzymes. Microbial fuel cells can achieve high efficiency in terms of conversion of chemical energy into electrical energy; however, problems associated with this approach include low volumetric catalytic activity of the whole organism and low power densities due to slow mass transport of the fuel across the cell wall (Moore et al., 2004a).

During the past decades, oxidoreductases enzymes have concerned the efforts of many research groups in the environmental and biotechnological field due to their huge potential to eradicate pollutants and catalyze a great range of redox processes with no hazardous side effects (Fernández-Sánchez et al., 2002). The application of redox enzymes for the targeted oxidation and reduction of particular fuel and oxidizer substrates at the electrode supports and the generation of the electrical current output is used for the development of biofuel cells (Ramanavicius et al., 2008). In Figure 1.7 a schematic view of a biofuel cell is presented.

The first enzyme-based biofuel cell was reported in 1964 using glucose as the fuel (Yahiro et al., 1964). Glucose is an ideal renewable fuel because it is produced by photosynthesis in plants such as sugar cane or corn which make it an attractive option as the fuel for portable fuel cells. Furthermore, glucose is plentiful in nature, cheap, environmentally benign, and easy to produce and handle (Ryu et al., 2010).

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Figure1.7 A generalized scheme of an enzymatic biofuel cell, where a fuel is oxidized at the anode and molecular oxygen is reduced at the cathode (Coman, 2009)

Semi- permeable membrane

Anode Cathode

Enzyme

Enzyme

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While glucose is the most common fuel used by enzymatic biofuel cells, it is not essentially the best for all applications (Sokic Lazic et al., 2010). There is a wealth of fuel choices such as pyruvate, its use was directed by its abundance and importance as a metabolic intermediate. Hydrogen as a non carbon containing fuel was known as a famous substrate for conventional fuel cells (Ivanov et al., 2010).

Other fuels that have been used in enzymatic fuel cells are aliphatic alcohols such as methanol which is an attractive alternative to dihydrogen as the anodic fuel due to its ready availability and easy to transport and store (Palmore et al., 1998). Ethanol has drawn further attention as a biofuel that can be produced by fermentation of biomass and is already commercially available for combustion engines. An enzymatic biofuel cell has been reported in which the anode and the cathode electrodes are both powered by ethanol and operate at ambient temperature (Ramanavicius, et al., 2008). Glycerol is also an attractive fuel due to its high energy density, low vapour pressure, and low toxicity opposed to the latter alcohols. It is also plentiful due to the fact that it is a byproduct of biodiesel production (Arechederra et al., 2007). Thus, the ability to oxidize such a higher order poly alcohols like glycerol would have a profound impact on the fuel cell market. The main fuels and the respective enzymes used for their bioelectrocatalytic oxidation are listed in Table 1.6.

Rujukan

DOKUMEN BERKAITAN

It was reported that when fluroxypyr was used for weed control in oil palm plantations, no residue was detected in crude palm oil and crude palm kernel oil irrespective of the

To study the catalytic activity of synthesized nanostructured composite materials as cracking catalysts for the conversion of used and crude palm oil and their selectivity

Enzymatic hydrolysis of oil-palm residues from oil palm trunk as a second-generation biofuel feedstock by potential lignocellulolytic fungal isolate,

a) To compare the operational parameters and biogas production between mesophilic and thermophilic anaerobic degradation processes under various hydraulic

1) To carry out the characterization of extracted starch from oil palm trunk for further modification. 2) To determine the compatibility of modified starch (CMS) from oil palm

In this study, spray characteristics of Refined Palm Oil (RPO) were studied in a Constant Volume Combustion Chamber (CVCC), and compared to conventional fuel such as gasoline

The present investigation focuses on hydrolysis of triglyceride to produce free fatty acids and glycerol from crude palm oil (CPO) using Candida rugosa lipase in batch

5.2 Effects of the substitution of dietary fish oil with crude palm oil and palm fatty acid distillate on growth, muscle fatty acids composition and the activities of hepatic