BIOCONVERSION OF OLD OIL-PALM TRUNK RESIDUES VIA ENZYMATIC HYDROLYSIS BY Penicillium rolfsii c3-2(1) IBRL USING REDUCING
SUGARS AS AN INDICATOR
by
LEE KOK CHANG
Thesis submitted in fulfillment of the requirements for the degree of
Doctor of Philosophy
APRIL 2015
ii
ACKNOWLEDGEMENTS
I would like to express my deep gratitude and sincere appreciation to my main
supervisor, Professor Dr. Darah Ibrahim, School of Biological Sciences, Universiti
Sains Malaysia for her invaluable and sound guidance, continued encouragement,
enthusiasm and tireless efforts throughout my studies. I am deeply grateful to her for
taking so much of her valuable time to discuss the finer points of the thesis with me
in order to complete this work in the present form. My deepest appreciation also goes
to my field supervisor, Dr. Takamitsu Arai for his beneficial guidance, unceasing
support and constructive reviews, patience and contributed experiences throughout
my studies.
A special sincere thank also goes to Ministry of Higher Education Malaysia allowing
me to pursue my postgraduate academic degree with the support of MyBrain15
program (MyPhD) as financial support to complete my study. I would like to thank
Japan International Research Center for Agricultural Sciences (JIRCAS) for the
greatest supports provided to carry out this entire project. My honest appreciation
also extended to Institute of Postgraduate Studies of Universiti Sains Malaysia for
providing financial supports through Postgraduate Research Grant Scheme, so that
my research works were completed smoothly.
Next, I would like to extend my sincere gratitude towards Dr. Akihiko Kosugi, Dr.
Yoshinori Murata, Panida Prawitwong, Akihiko Hirooka, Deng Lan and other
members or friends from JIRCAS who had rendered their helps throughout my
research. Again, with a deep sense of honor, I wish to thank En. Johari, Kak Jamilah
iii
and En. Rizal at Microscopy Unit of School of Biological Sciences for their patient
guidance for teaching me on SEM sample preparations. I am exceedingly grateful to
Dr. Leh Cheu Peng, Yin Hui and Ying Ying from School of Technology Industry
USM for providing all the laboratory facilities and assisting me throughout my
research. Special thank also goes to Li Yee for her sincere assistance and guidance as
well.
I would like to acknowledge all my labmates especially Syarifah Ab Rashid, Tong
Woei Yenn, Azza, Chee Keong, Wani and everyone who has been part of the team in
Industrial Biotechnology Research Laboratory for their constructive ideas and helps,
good companionship and also sharing the good memories together that will never be
forgotten.
Last but not least, I would like to take this opportunity to express my deepest
gratitude to my family and my wife Teo Kah Cheng, for their great patience,
encouragement, supports, love and understanding throughout the long journey for
completing my study. And finally to all whose names did not appear here, thanks a
lot.
iv
TABLE OF CONTENTS
PAGE
ACKNOWLEDGEMENTS ii TABLE OF CONTENTS iv LIST OF TABLES xiii LIST OF FIGURES xv LIST OF PLATES xviii LIST OF ABBREVIATIONS xix LIST OF PUBLICATIONS AND CONFERENCE PAPERS xxi ABSTRAK xxiii
ABSTRACT xxvi
CHAPTER ONE: INTRODUCTION 1 1.1 The potential of oil-palm trunk biomass as an alternative source for
production of lignocellulolytic enzymes 1
1.2 Microorganisms and their lignocellulolytic enzymes 4 1.3 Biotechnological applications 4 1.4 Objectives of research 5 1.5 Scope of study 5 CHAPTER TWO: LITERATURE REVIEW 7 2.1 Lignocellulosic Biomass 7 2.1.1 Structure and composition of lignocellulosic biomass 7 2.1.1.1 Cellulose 12 2.1.1.2 Hemicellulose 13 2.1.1.3 Lignin 15 2.2 Pretreatment of lignocellulosic biomass 16
v
2.2.1 Physical pretreatments 23 2.2.1.1 Mechanical comminution 23 2.2.1.2 Pyrolysis 23 2.2.1.3 Extrusion 24 2.2.1.4 Irradiation 25 2.2.2 Chemical pretreatments 26 2.2.2.1 Ozonolysis 26 2.2.2.2 Alkaline pretreatment 27 2.2.2.3 Acid pretreatment 29 2.2.2.4 Organosolvation 30 2.2.3 Physicochemical pretreatment 31 2.2.3.1 Steam explosion 31 2.2.3.2 Liquid hot water pretreatment 33 2.2.3.3 Ammonia fiber explosion (AFEX) 34 2.2.4 Biological pretreatment 35 2.3 Enzymatic hydrolysis of lignocellulosic biomass 37 2.3.1 Degradation of cellulose 37 2.3.2 Degradation of hemicellulose 41 2.3.3 Degradation of lignin 43 2.4 Microbial degradation of lignocellulosic biomass 46 2.4.1 Degradation by actinomycetes and bacteria 47 2.4.2 Degradation by fungi 48 2.5 Fermentation strategies 51 2.5.1 Separate hydrolysis and fermentation (SHF) 51 2.5.2 Simultaneous saccharification and fermentation (SSF) 54
vi
2.5.3 Consolidated bioprocessing (CBP) 56 2.6 Biotechnological application of lignocellulolytic enzymes 59 2.6.1 Pulp and paper industry 59 2.6.2 Bioconversion of lignocellulosic materials to biofuel 60 2.6.3 Animal feed 62 2.6.4 Food industry 63 2.7 Concluding remarks 63 CHAPTER THREE: GENERAL MATERIALS AND METHODS 65 3.1 Source of microorganisms 65 3.1.1 Fungal isolates from soil samples 65 3.1.2 Stock fungal cultures from IBRL 65 3.2 Maintenance of fungal isolates 65 3.3 Preparation of oil-palm trunk residues 66 3.4 Nutrient medium preparation for fungal growth 66 3.5 Enzyme production and protein determination 67 3.6 Enzyme assays 68 3.6.1 Determination of oil-palm trunk residues activity 68 3.6.2 Determination of total cellulase activity 68 3.6.3 Determination of cellobiase activity 69 3.6.4 Determination of arabinase activity 69 3.6.5 Determination of avicelase (exoglucanase) activity 70 3.6.6 Determination of carboxymethyl cellulase (CMCase) activity 70 3.6.7 Determination of β-glucanase activity 71 3.6.8 Determination of laminarinase activity 71 3.6.9 Determination of xylanase activity 71
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CHAPTER FOUR: ISOLATION, SCREENING AND SELECTION FOR LIGNOCELLULOSE-HYDROLYTIC FILAMENTOUS FUNGI USING OIL-PALM TRUNK RESIDUES
73
4.1 Introduction 73 4.2 Materials and methods 74 4.2.1 Preparation of oil-palm trunk residues 74 4.2.2 Samples for fungal source 74 4.2.2.1 Soil samples 74 4.2.2.2 Stock fungal cultures from IBRL 74 4.2.3 Isolation of potential lignocellulolytic fungal isolates 75 4.2.3.1 Nutrient medium preparation 75 4.2.3.2 Serial dilution of soil samples 75 4.2.3.3 Incubation in multi-plates and observation of
potential fungal growth 75
4.2.4 Maintenance of fungal isolates 75 4.2.5 Screening of potential fungal Isolates by submerged
fermentation 76
4.2.5.1 Primary screening by determination of specific
activity on oil-palm trunk residues 76
4.2.5.2
Secondary screening by determination of reducing sugar production from hydrolysis of oil-palm trunk residues
76
4.2.6 Morphological observation of selected potential
lignocellulolytic fungus 77
4.2.6.1 Media and growth conditions 77 4.2.6.2 Preparation of inoculum for cultivation on standard
agar media 78
4.2.6.3 Micromorphological observation by using light
microscope 78
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4.2.6.4 Microscopic examination by Scanning Electron Microscope
79
4.2.7 Molecular identification 80 4.2.7.1 DNA extraction, PCR amplification and DNA
sequencing 80
4.3 Results and discussion 81 4.3.1 Isolation and selection of potential fungal isolates for
degradation of oil-plam trunk residues 81
4.3.2 Morphological identification of selected potential
lignocellulolytic fungus 85
4.3.2.1 Morphological features on different agar media 85 4.3.2.2 Microscopic features of isolate c3-2(1) 93 4.3.2.3 Identification key 99 4.3.3 Molecular identification 101 4.4 Conclusion 106 CHAPTER FIVE: LIGNOCELLULOLYTIC ENZYMES
PRODUCED BY Penicillium rolfsii c3-2(1) IBRL FOR HYDROLYSIS OF OIL-PALM TRUNK RESIDUES
107
5.1 Introduction 107 5.2 Materials and methods 109 5.2.1 Preparation and alkaline-pretreatment of oil-palm trunk
residues 109
5.2.2 Compositional analysis of oil-palm trunk residues 110 5.2.3 Microorganisms and culture conditions 110 5.2.4 Enzyme assays 111 5.2.5 Detection of halozone on agar medium 111 5.2.6 Analysis of crude enzymes by SDS-PAGE 112
ix
5.2.7 Determination of optimal pH, temperature and thermal stability of crude enzyme
113
5.2.8 Time-course production of cellulolytic and hemicellulolytic
enzymes using oil-palm trunk residues by Penicillium rolfsii c3-2(1) IBRL
114
5.2.9 Saccharification experiment 114 5.2.10 Adsorption experiments and effect of hydrolysis by lignin 115 5.2.11 Microscopic observation of changes of plant cell structure in
oil-palm trunk residues due to enzymatic hydrolysis 116
5.2.12 Statistical analysis 116 5.3 Results and discussion 117 5.3.1 Compositional analysis of unpretreated and alkaline
pretreated oil-palm trunk residues 117
5.3.2 Enzyme activities and SDS-PAGE protein profiles analysis
of P. rolfsii c3-2(1) IBRL induced by different carbon sources
121
5.3.3 Optimal conditions of lignocellulolytic activities for P. rolfsii
c3-2(1) IBRL 128
5.3.4 Time-course production of cellulolytic and hemicellulolytic
enzymes by P. rolfsii c3-2(1) IBRL 131
5.3.5 Saccharification of alkaline-pretreated oil-palm trunks
residues with different enzyme dosage of crude enzyme from P. rolfsii c3-2(1) IBRL compared to commercial enzymes
135
5.3.6 Effect of lignin residues on enzymatic hydrolysis of oil-palm
trunk residues 144
5.3.7 Scanning electron micrographs of oil-palm trunk residues
due to enzymatic hydrolysis by P. rolfsii c3-2(1) IBRL 153
5.4 Conclusion 155 CHAPTER SIX: PURIFICATION AND CHARACTERIZATION
OF CELLULASE-FREE XYLANASE AND LAMINARINASE FROM THE NEWLY ISOLATED Penicillium rolfsii c3-2(1) IBRL
157
6.1 Introduction 157
x
6.1.1 Xylanase 158 6.1.2 Laminarinase 159 6.2 Materials and methods 160 6.2.1 Microorganism and culture condition 160 6.2.2 Purification of xylanase and laminarinase 161 6.2.3 Enzyme assay and protein determination 162 6.2.4 Activity detection by zymography technique 162 6.2.4.1 Detection of xylanase activity 162 6.2.4.2 Detection of laminarinase activity 163 6.2.5 Effects of pH and temperature on enzyme activity 163 6.2.5.1 Effects of pH and temperature on xylanase activity 163 6.2.5.2 Effects of pH and temperature on laminarinase
activity 164
6.2.6 Effects of pH and temperature on enzyme stability 164 6.2.6.1 Effects of pH and temperature on xylanase stability 164 6.2.6.2 Effects of pH and temperature on laminarinase
stability 164
6.2.7 Determination of kinetic parameters 165 6.2.7.1 Kinetic parameters for purified xylanase 165 6.2.7.2 Kinetic parameters for purified laminarinase 165 6.2.8 Analysis of hydrolysis products 165 6.2.8.1 Analysis of xylanase hydrolysis products 165 6.2.8.2 Analysis of laminarinase hydrolysis products 166 6.2.9 Substrate specificity 167 6.2.9.1 Substrate specificity of xylanase 167 6.2.9.2 Substrate specificity of laminarinase 167
xi
6.2.10 Effect of xylanase supplementation on the enzymatic
hydrolysis of oil-palm trunk residues 168
6.3 Results and discussion 169 6.3.1 Purification of xylanase 169 6.3.2 Purification of laminarinase 177 6.3.3 Effect of pH and temperature on enzyme activity and its
stability 182
6.3.3.1 Effect of pH and temperature on xylanase activity
and its stability 182
6.3.3.2 Effect of pH and temperature on laminarinase
activity and its stability 186
6.3.4 Determination of kinetic parameters 190 6.3.4.1 Kinetic parameters for purified xylanase 190 6.3.4.2 Kinetic parameters for purified laminarinase 190 6.3.5 Analysis of hydrolysis products 193 6.3.5.1 Analysis of xylanase hydrolysis products 193 6.3.5.2 Analysis of laminarinase hydrolysis products 194 6.3.6 Substrate specificity 196 6.3.6.1 Substrate specificity of xylanase 196 6.3.6.2 Substrate specificity of laminarinase 198 6.3.7 Effect of xylanase supplementation on the enzymatic
hydrolysis of oil-palm trunk residues 199
6.4 Conclusion 202 CHAPTER SEVEN: CONCLUSION AND RECOMMENDATIONS
FOR FUTURE RESEARCH 203
7.1 Conclusions 203 7.2 Recommendation and future research 206
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REFERENCES 207 APPENDICES APPENDIX 1: STANDARD CURVE APPENDIX 2: CHROMATOGRAM APPENDIX 3: MEDIA APPENDIX 4: REAGENTS AND SOLUTIONS APPENDIX 5: SCREENING RESULTS APPENDIX 6: REFERENCE PAPER APPENDIX 7: CELLUCLAST 1.5L APPENDIX 8: ACCELLERASE 1500
xiii
LIST OF TABLES
PAGE Table 2.1 Composition of representative lignocellulosic feedstocks. 10 Table 2.2 Most promising pretreatment technologies (advantages and
disadvantages) 19
Table 2.3 Lignocellulosic enzymes produced by some filamentous
fungi with several agricultural residues. 52
Table 4.1 Source, type and location of samples. 83 Table 4.2 Culture of fungal isolate c3-2(1) on different standard
media, colony color and textures. 87
Table 4.3 Comparison of macroscopic characteristics of fungal isolate
c3-2(1) and P. rolfsii Thom 91
Table 4.4 Comparison of microscopic characteristics of fungal isolate
c3-2(1) and P. rolfsii Thom 97
Table 4.5 Some of the characteristics amongst the subgenus of
Aspergilloides, Penicillium, Biverticillium, and Furcatum based on their penicillus type.
98
Table 4.6 Top five of BLASTn Algorithm Search’s result for fungal
isolate c3-2(1) 105
Table 5.1 Compositional analysis of oil-palm trunk residues 118 Table 5.2 Filter paper unit (FPU/ml), cellobiase unit (CBU/ml) and
protein concentration of P. rolfsii c3-2(1) IBRL and two commercial enzymes
122
Table 5.3 Specific activity (U/mg protein) of composite
lignocellulolytic enzymes of P. rolfsii c3-2(1) IBRL and two commercial enzyme preparations on selected preparations.
122
Table 5.4 Production of cellulolytic and hemicellulolytic enzymes
with other different Penicillium sp. under submerged cultivations
133
Table 5.5 Comparison of the hydrolytic performance of cellulases
from various Penicillium species and T. reesei 141
Table 5.6 Effects of isolated lignins from various sources on
hydrolysis of different types of lignocellulosic materials 151
xiv
Table 6.1 Summary of purification scheme for the xylanase of P.
rolfsii c3-2(1) IBRL 173
Table 6.2 Summary of purification scheme for the laminarinase of P.
rolfsii c3-2(1) IBRL 180
Table 6.3 Relative substrate specificity of purified xylanase from P.
rolfsii c3-2(1) IBRL 197
Table 6.4 Relative substrate specificity of purified laminarinase from
P. rolfsii c3-2(1) IBRL 199
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LIST OF FIGURES
PAGE Figure 2.1 Diagrammatic illustration of the framework of
lignocellulose 8
Figure 2.2 Schematic structural formula cellulose (glucan),
hemicelluloses (homoxylan) and lignin (core lignin) 9
Figure 2.3 Schematic of aims of pretreatment on lignocellulosic
biomass 17
Figure 2.4 Energy requirements for ball milling municipal solid waste 24 Figure 2.5 Schematic diagram of the cellulosome 40 Figure 2.6 Sites of attack on a fragment of a glucuronoarabinoxylan by
microbial xylanolytic enzymes 42
Figure 2.7 Lignin biodegradation process by white rot fungi 45 Figure 2.8 Evolution of biomass processing configurations featuring
enzymatic hydrolysis 53
Figure 2.9 Alternative organism development strategies to obtain
organisms useful in processing cellulosic feedstocks 58
Figure 4.1 Time-course profile of selected potential fungal isolates
which were showing the capability on hydrolysis of oil-palm trunks residues evaluated by sugar production concentration (mM)
84
Figure 4.2 Consensus sequence for fungal isolate c3-2(1) 104 Figure 5.1 Sugars released from oil-palm trunk residues when acted
with crude enzymes of P. rolfsii c3-2(1) IBRL 119
Figure 5.2 Effect of alkaline pretreated oil-palm trunk residues
compared to unpretreated oil-palm trunk residues hydrolyzed by crude enzyme of P. rolfsii c3-2(1) IBRL
120
Figure 5.3 Effect of pH on oil-palm trunk residues lignocellulolytic
activity by P. rolfsii c3-2(1) IBRL enzyme preparation 129
Figure 5.4 Effect of temperature on oil-palm trunk residues
lignocellulolytic activity by P. rolfsii c3-2(1) IBRL enzyme preparation
129
xvi
Figure 5.5 Residual activity expressed as percentage of the maximum oil-palm trunk residues activity by P. rolfsii c3-2(1) IBRL
130
Figure 5.6 Activities on oil-palm trunk residues, avicel, filter paper,
carboxymethylcellulose (CMC), laminarin from Laminaria digitata and xylan from birchwood in submerged cultures of P. rolfsii c3-2(1) IBRL
132
Figure 5.7 Time course for hydrolysis of oil-palm trunk residues by
using P. rolfsii c3-2(1) IBRL enzyme and commercial enzymes based on hydrolysis of total sugar conversion (%)
136
Figure 5.8 Hydrolysis of oil-palm trunk residues by using P. rolfsii c3-
2(1) IBRL enzymes and commercial enzymes based on total sugar conversion (%) with different enzyme dosage
138
Figure 5.9 Absorption of P. rolfsii c3-2(1) IBRL enzymes and
commercial enzymes on Klason lignin residues after 1.5 h at 4°C.
145
Figure 5.10 Influence of increasing Klason lignin loading on the extent
of enzymatic hydrolysis of oil-palm trunk residues (10 mg/ml) by P. rolfsii c3-2(1) IBRL enzyme, commercial enzymes Celluclast 1.5L and Accellerase 1500
147
Figure 6.1 Anion exchange chromatography (RESOURCETM Q, 6 ml)
elution profile of protein and xylanase activity 170
Figure 6.2 Elution profile of xylanase purification by gel filtration
HiPrepTM 16/60 Sephacryl S-100 High Resolution 171
Figure 6.3 Hydrophobic-interaction chromatography of xylanase on
phenyl ResourceTM PHE column after gel-filtration column chromatography
172
Figure 6.4 Anion exchange chromatography (RESOURCETM Q, 6 ml)
elution profile of protein and laminarinase activity 178
Figure 6.5 Elution profile of laminarinase purification by gel filtration
HiPrepTM 16/60 Sephacryl S-100 High Resolution 178
Figure 6.6 Hydrophobic-interaction chromatography of laminarinase
on phenyl ResourceTM PHE column after gel-filtration column chromatography
179
Figure 6.7 Effect of pH on the activity and its stability of P. rolfsii c3-
2(1) IBRL xylanase 183
Figure 6.8 Influence of temperature on activity and its thermal stability
of P. rolfsii c3-2(1) IBRL xylanase 185
xvii
Figure 6.9 Effect of pH on the activity and its stability of P. rolfsii c3-
2(1) IBRL laminarinase 187
Figure 6.10 Influence of temperature on activity and its thermal stability
of P. rolfsii c3-2(1) IBRL laminarinase 188
Figure 6.11 Michaelis-Menten plot and Lineweaver-Burk plot for
determining the Vmax and Km values of xylanase of P. rolfsii c3-2(1) IBRL
191
Figure 6.12 Michaelis-Menten plot and Lineweaver-Burk plot for
determining the Vmax and Km values of laminarinase of P. rolfsii c3-2(1) IBRL
192
Figure 6.13 TLC analysis for hydrolysis products released from xylo-
oligosaccharides 194
Figure 6.14 TLC analysis for hydrolysis products released from
laminari-oligosaccharides 195
Figure 6.15 Effect of purified xylanase supplementation (0.1 mg/g
substrate) added with commercial enzymes Celluclast 1.5L and Accellerase 1500 on conversion of oil-palm trunk residues after 72 h hydrolysis
200
xviii
LIST OF PLATES
PAGE Plate 4.1 The powdery oil-palm trunk residues that was used as
substrate for isolation of lignocellulolytic fungi and autoclaved oil-palm trunk residues in the multi-plate’s well showing the growth of some fungal colonies as indicated with red arrows
82
Plate 4.2 Colonies of isolate c3-2(1) on CYA and MEA for front and
reverse view at 25°C, after 7 days 88
Plate 4.3 Colonies of isolate c3-2(1) on CYA, 37°C; G25N and
CREA, 25°C for front and reverse view after 7 days 90
Plate 4.4 Conidiophore of Penicillium sp. c3-2(1) bearing terminal
biverticillate stained with lactophenol cotton blue 94
Plate 4.5 SEM micrographs showing the (A) penicilli structure of
Penicillium sp. c3-2(1) 96
Plate 4.6 Genomic DNA of the fungal isolate c3-2(1) resolved by 1%
of agarose gel electrophoresis 102
Plate 4.7 PCR amplification product resolved by 1% of agarose gel
electrophoresis 103
Plate 5.1 Plates showing zone of clearances around the fungal colony
P. rolfsii c3-2(1) IBRL after 2 days incubation at 30°C from the effect of enzymatic hydrolysis on CMC, laminarin from Laminaria digitata, and xylan from birchwood agars.
125
Plate 5.2 SDS-PAGE of protein profiles of crude enzymes induced by
different carbon sources 127
Plate 5.3 The view of extracted powdery klason lignin from oil-palm
trunk residues 148
Plate 5.4 Scanning electron micrographs of oil-palm trunk residues
that were unpretreated and alkaline pretreated samples. 154
Plate 6.1 Development of xylanase zymogram 175 Plate 6.2 Development of laminarinase zymogram 181
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LIST OF ABBREVIATIONS
AAD Aryl-alcohol dehydrogenases AAO Aryl alcohol oxidase AFEX Ammonia fiber explosion ANF Antinutritional factors ANOVA Analysis of variance ATCC American Type Culture Collection BGL β-glucosidase bp Base pair BSA Bovine serum albumin CBD Cellulose-binding domain CBH Cellobiohydrolase CBM Carbohydrate-binding module CBP Consolidated bioprocessing CD Catalytic domain cm Centimeter CMC Carboxymethylcellulose CMCase Carboxymethylcellulase cP Centipoise CREA Creatine sucrose agar CYA Czapek yeast agar DCM Direct microbial conversion DNA Deoxyribonucleic acid dNTP Deoxyribonucleotide triphosphate DNS Dinitrosalicylic acid DP Degree of polymerization EG Endo-β-1,4-glucanase FPU/g Filter paper unit per gram FPU/ml Filter paper unit per milliliter g Gram g m-3 Gram per cubic meter G25N Glycerol nitrate agar h Hour HMF 5-hydroxy-2-methyl-furfural IBRL Industrial Biotechnology Research Laboratory ITS Internal transcribed spacer JIRCAS Japan International Research Center for Agricultural Sciences kDa Kilodalton kg Kilogram kHz Kilohertz kPa m2/g Kilo pascal meter square per gram LHW Liquid hot water LiP Lignin peroxidases MEA Malt extract agar mg Milligram mg/ml Milligram per milliliter
xx
min Minute µl Microliter mm Millimeter mM Milimolar mN m2/g Millinewton meter square per gram MnP Manganese peroxidases MOPS 3-morpholinopropanesulfonic acid MPa Megapascal MSM Minimum salts medium NBRC NITE Biological Resource Center NCBI National Center for Biotechnology Information Nm/g Newton meter per gram OsO4 Osmium tetroxide PCR Polymerase chain reaction PDA Potato dextrose agar psi Pound per square inch rpm Revolutions per minute rRNA Ribosomal ribonucleic acid SDS-PAGE Sodium dodecyl sulfate polyacrylamide gel electrophoresis SEM Scanning electron microscope SHF Separate hydrolysis and fermentation SSCF Simultaneous saccharification and co-fermentation SSF Simultaneous saccharification and fermentation TAPPI Technical Association of the Pulp and Paper Industry TEMED Tetramethylethylenediamine TLC Thin layer chromatography U/mg Unit per milligram U/ml Unit per milliliter w/v Weight per volume w/w Weight per weight
xxi
LIST OF PUBLICATIONS AND CONFERENCE PAPERS
1. Lee, K. C., Arai, T., Ibrahim, D., Prawitwong, P., Deng, L., Murata, Y., Mori, Y. and Kosugi, A. (2015). Purification and characterization of a xylanase from the newly isolated Penicillium rolfsii c3-2(1) IBRL. BioResources 10(1):1627-1643.
2. Lee, K. C., Arai, T., Ibrahim, D., Kosugi, A., Prawitwong, P., Lan, D., Murata, Y. and Mori, Y. (2014). Purification and characterization of a thermostable laminarinase from Penicillium rolfsii c3-2(1) IBRL. BioResources 9(1):1072-1084.
3. Prawitwong, P., Kosugi, A., Arai, T., Deng, L., Lee, K. C., Ibrahim, D.,
Murata, Y., Sulaiman, O., Hashim, R., Sudesh, K., Ibrahim, W. A. B., Saito, M. and Mori, Y. (2012) Efficient ethanol production from separated parenchyma and vascular bundle of oil palm trunk. Bioresource Technology 125:37-42.
4. Lee, K. C., Arai, T., Kosugi, A., Darah, I., Prawitwong, P., Mori, Y. (2012). Penicillium rolfsii, The potential lignocellulolytic fungus on hydrolysis of oil-palm residues from oil palm trunk as a second generation biofuel feedstock. The 2nd Annual International Conference Unsyiah & 8th IMTGT Uninet Biosciences Conference. Universitas Syiah Kuala, 22-24 November 2012.
5. Lee, K. C., Arai, T., Kosugi, A., Darah, I., Prawitwong, P., Mori, Y. (2012).
Fungal bioconversion of old oil-palm trunks by enzymatic hydrolysis on development of alternate energy source. The 2nd Annual International Conference Unsyiah & 8th IMTGT Uninet Biosciences Conference. Universitas Syiah Kuala, 22-24 November 2012.
6. Lee, K. C., Arai, T., Kosugi, A., Darah, I., Prawitwong, P., Murata, Y., Mori,
Y. (2012). Degradation activity of Penicillium rolfsii strain isolated from the soil in Malaysia against the sap extraction residue of oil palm trunks. Annual Meeting of Japan Society for Bioscience, Biotechnology, and Agrochemistry (JSBBA) 2012. Kyoto Women’s University, 23-25, March 2012.
7. Lee, K.C., Darah, I. and Arai, T. (2013). Purification and characterization of xylanase and laminarinase produced by Penicillium rolfsii. Proceedings of the 8th Annual PPSKH Postgraduate Colloqium, Shool of Biological Sciences, Universiti Sains Malaysia. School of Biological Sciences, USM, Penang 5-6th June 2013.
xxii
8. Lee, K.C., Darah, I. and Arai, T. (2012). Enzymatic hydrolysis of oil-palm residues from oil palm trunk as a second-generation biofuel feedstock by potential lignocellulolytic fungal isolate, Penicillium rolfsii. Proceedings of the 7th Annual PPSKH Postgraduate Colloqium, Shool of Biological Sciences, Universiti Sains Malaysia. School of Biological Sciences, USM, Penang,7-8th November 2012.
9. Lee, K.C., Darah, I. and Arai, T. (2010). A study on the involvement of various enzyme in the degradation of parenchyma tissue of oil palm trunk by fungal isolates. Proceedings of the 4th Annual PPSKH Postgraduate Colloqium, Shool of Biological Sciences, Universiti Sains Malaysia. School of Biological Sciences, USM, Penang,15th December 2010.
xxiii
BIOPENUKARAN RESIDU BATANG KELAPA SAWIT TUA MELALUI HIDROLISIS BERENZIM OLEH Penicillium rolfsii c3-2(1) IBRL
MENGGUNAKAN GULA PENURUN SEBAGAI PENANDA
ABSTRAK
Produktiviti minyak yang semakin menurun daripada pokok kelapa sawit tua selepas
25 tahun penanamannya telah menyebabkan pembuangan sisa batang kelapa sawit
yang banyak daripada aktiviti penanaman semula. Batang kelapa sawit didapati
mengandungi sap yang tinggi dengan kandungan gula yang tinggi. Residu batang
kelapa sawit yang telah diperah sapnya merupakan bahan sisa yang akan dibuang
dalam kuantiti yang besar. Biojisim residu batang kelapa sawit dalam bentuk
buangan agro-industri merupakan sumber yang berpotensi untuk dijadikan sebagai
bahan mentah bagi penukaran ke bahan api bio, dan bahan-bahan kimia tambahan
yang bermutu tanpa persaingan dengan sumber makanan. Pencilan kulat berfilamen
yang dilabelkan sebagai c3-2(1) telah disaring dan dipilih menunjukkan aktiviti
enzim yang tinggi terhadap residu batang kelapa sawit. Dengan menggunakan kaedah
pencirian morfologi, pemerhatian melalui mikroskop dan identifikasi molekul,
pencilan c3-2(1) telah dikenalpasti sebagai Penicillium rolfsii. Pencilan tersebut
dinamakan sebagai Penicillium rolfsii c3-2(1) IBRL. Aktitviti spesifik terhadap
residu batang kelapa sawit yang lebih tinggi telah ditunjukkan dengan menggunakan
ampaian yang diperoleh daripada P. rolfsii c3-2(1) IBRL berbanding dengan enzim
komersial, di mana 2 hingga 3 kali ganda aktiviti yang lebih tinggi berbanding
dengan Celluclast 1.5L (Sigma) dan 3 hingga 4 kali ganda aktiviti yang lebih tinggi
berbanding dengan Accellerase 1500 (Genencor) telah ditunjukkan. Selain itu, P.
rolfsii c3-2(1) IBRL juga menunjukkan aktiviti enzim yang lebih tinggi terhadap
xilan, arabinan dan laminarin berbanding dengan enzim komersial. Hidrolisis untuk
xxiv
residu batang kelapa sawit dalam keadaan optimum didapati pada pH 5.0 dan suhu
50°C, dengan kestabilan haba yang lebih tinggi bagi enzim mentahnya. Selepas 48-
72 jam proses sakarifikasi biojisim, peningkatan sebanyak 1 hingga 1.5 kali ganda
jumlah penukaran gula telah ditunjukkan oleh enzim daripada P. rolfsii c3-2(1) IBRL
berbanding dengan enzim komersial. Jumlah hasil penukaran gula maksimum, iaitu
kira-kira 56% telah diperolehi daripada hidrolisis berenzim oleh enzim mentah P.
rolfsii c3-2(1) IBRL terhadap residu batang kelapa sawit berbanding dengan jumlah
hasil penukaran gula sebanyak 43% oleh Celluclast 1.5L dan 40% oleh Accellerase
1500 apabila kepekatan enzim 14 FPU/g substrat ditambahkan untuk proses
sakarifikasi dalam tempoh 48 jam. Residu lignin yang dipencilkan daripada residu
batang kelapa sawit ini didapati memberi kesan terhadap hidrolisis biojisim dan
membuktikan bahawa P. rolfsii c3-2(1) IBRL berupaya menghasilkan enzim ‘ikatan-
lignin’ yang lemah, seterusnya menyumbang pada kecekapan yang lebih tinggi
terhadap hidrolisis residu batang kelapa sawit. Xilanase dan laminarinase telah
berjaya ditulenkan daripada P. rolfsii c3-2(1) IBRL. Xilanase ini didapati
mempunyai sifat-sifat fizikal dan kimia seperti berikut: kespesifikan substrat
terhadap xilan daripada birchwood, Vmax= 691.6 μmol/min/mg dan Km= 5.72 mg/ml;
pH optimum, 5.0; suhu optimum, 50°C; kestabilan suhu selepas rawatan xilanase
pada 50°C untuk 4 jam (residu aktiviti > 90%); berat molekul berdasarkan analisis
SDS-PAGE dianggarkan 35 kDa. Laminarinase juga telah diuji terhadap laminarin
yang diperolehi daripada Laminaria digitata sebagai substrat di mana Vmax= 372.2
μmol/min/mg dan Km= 0,0817 mg/ml; pH optimum, 5.0; suhu optimum, 70°C;
kestabilan suhu selepas rawatan laminarinase pada suhu ≤ 55°C selama 4 jam (residu
aktiviti > 90%); berat molekul berdasarkan analisis SDS-PAGE dianggarkan 75 kDa.
Xilanase yang ditulenkan didapati meningkatkan kesan sinergi bersama-sama dengan
xxv
enzim komersial dalam proses sakarifikasi terhadap residu batang kelapa sawit
berdasarkan jumlah penukaran gula yang dihasilkan dan ini menunjukkan ia
merupakan salah satu enzim yang berperanan utama untuk hidrolisis residu batang
kelapa sawit.
xxvi
BIOCONVERSION OF OLD OIL-PALM TRUNK RESIDUES VIA ENZYMATIC HYDROLYSIS BY Penicillium rolfsii c3-2(1) IBRL USING
REDUCING SUGARS AS AN INDICATOR
ABSTRACT
Due to the decreased oil productivity of old oil-palm trees after 25 years, large
quantities of trunks as the waste were generated from the replanting activities. The
felled old oil-palm trunk was found containing large quantity of sap with high
concentration of sugar contents. The oil-palm trunk residues which are the residual
substances after squeezing sap will be discharged in large quantity. These oil-palm
trunk residues in the form of agro-industrial waste constitute a potentially enormous
source of feedstock for bioconversion into biofuel, and other value-added chemicals
without competition with food sources. A filamentous fungus namely c3-2(1) was
screened and selected for its strong activities against oil-palm trunk residues. Fungal
isolate c3-2(1) was identified as Penicillium rolfsii by morphological characterization,
microscopical observations and confirmed by molecular identification. It was
designated as Penicillium rolfsii c3-2(1) IBRL. Oil-palm trunk residues-hydrolyzing
specific activity of the culture supernatant from P. rolfsii c3-2(1) IBRL was found
superior to those of commercial enzymes Celluclast 1.5L (Sigma) and Accellerase
1500 (Genencor) which exhibited 2 to 3-fold and 3 to 4-fold higher activity,
relatively. On the other hand, P. rolfsii c3-2(1) IBRL exhibited a greater xylan,
arabinan and laminarin-hydrolyzing activities than those commercial enzymes. The
optimal conditions for oil-palm residues hydrolysis was found at pH 5.0 and
temperature of 50°C, with higher thermal-stability of crude enzymes. After 48–72 h
of biomass saccharification, 1 to 1.5-fold higher total sugar conversion was
performed by enzyme of P. rolfsii c3-2(1) IBRL compared to commercial enzymes.
xxvii
Maximum total sugar conversion yield of approximately 56% was obtained from
enzymatic hydrolysis on oil-palm trunk residues by crude enzyme of P. rolfsii c3-
2(1) IBRL compared to total sugar conversion yield 43% from Celluclast 1.5L and
40% from Accellerase 1500 when 14 FPU/g substrate of enzyme loading added at 48
h reaction. The isolated lignin residual from oil-palm trunk residues affected the
biomass hydrolysis, which revealed that P. rolfsii c3-2(1) IBRL is capable to produce
weak ‘lignin-binding’ enzymes which might contribute to the higher efficiency
hydrolysis on oil-palm residues. Purification of xylanase and laminarinase were
successfully achieved from P. rolfsii c3-2(1) IBRL. The xylanase had the following
physical and chemical properties: substrate specificity on xylan from birchwood,
Vmax=691.6 µmol/min/mg and Km=5.72 mg/ml; optimum pH, 5.0; optimum
temperature, 50°C; temperature stability after the treatment at 50°C for 4 hr (residual
activity > 90%); molecular weight by SDS-PAGE analysis, about 35 kDa. The
laminarinase was tested on laminarin from Laminaria digitata as substrate in which
Vmax=372.2 µmol/min/mg and Km=0.0817 mg/ml; optimum pH, 5.0; optimum
temperature, 70°C; temperature stability after the treatment at ≤ 55°C for 4 hr
(residual activity > 90%); molecular weight by SDS-PAGE analysis, about 75 kDa.
Considerable increasing synergism effect was observed on added purified xylanase
with commercial enzymes during saccharification of oil-palm residues based on total
sugar conversion, suggesting it is one of the key enzymes for the hydrolysis of oil-
palm trunk residues.
1
CHAPTER ONE
INTRODUCTION
1.1 The potential of oil-palm trunk biomass as an alternative source for
production of lignocellulolytic enzymes
Energy crisis is one of the most serious threats towards the sustainability of human
kinds and civilization. Excessive global consumption of energy, such as fossil fuels,
particularly in large urban areas with expansion of human population and increase of
industrial prosperity, high levels of pollution and greenhouse gasses in the
atmosphere have increased drastically during the last few decades (Sarkar et al.,
2012). Furthermore, the shortage of fossil fuels parallel to the global consumption of
fuels and consequences of climate change induced by greenhouse gas emissions
(Singh et al., 2011; Sulaiman et al., 2012), have led the tremendous focus on using
lignocellulosic biomass for the production of cellulases and other lignocellulolytic
enzymes. Lignocellulosic biomass originated from agricultural and forestry residues
and herbaceous is abundantly available renewable carbon source which can be
converted for further usage of fuel and chemical production (Rahikainen et al., 2011),
concomitantly to mitigate dependence on depleting fossil oil (Jørgensen et al., 2007).
Each year, more than 40 million tonnes of inedible plant materials are produced from
agricultural residues and much of which are thrown away (Sanderson, 2011). For
instance, some of the readily available lignocellulosic biomass is left at the field as a
waste and is burned after harvesting through agricultural burning activity (Dawson
and Boopathy, 2007), and this included wastes from oil palm plantation and industry.
Each year, there are approximately more than 30 million tons of biomass in the form
2
of empty fruit bunches, oil-palm trunks and oil-palm fronds generated in the oil palm
industry in Malaysia (Sulaiman et al., 2012). For instance, oil-palm trees with low
productivity of palm oil after 20 to 25 years of age (Lim et al., 1997; Jung et al.,
2011); these trees are chopped down and trunks are left to rot in the field (Yamada et
al., 2010). Consequently, the felled palm trunks represent one of the most important
biomass resources in Malaysia and Indonesia (Sumathi et al., 2008; Shuit et al.,
2009). These oil-palm trunks which are considered as wastes could be made use for
conversion of fermentable sugars and biofuels, in helping societies less dependent
solely on oil (Rostrup-Nielsen, 2005). Furthermore, when these discarded, woody
bits of plants are converted into value-added renewable resources, it can then be
fulfilling the term of ‘second-generation’ biofuels which might gradually eliminate
the use of ‘first-generation’ biofuels without competition with edible food crops such
as sugar cane and corn (Graham-Rowe, 2011; Sanderson, 2011). Accumulation of
lignocellulosic materials in abundance in places where agricultural residues present a
disposal problems, results not only detrimental to the environment but also in loss of
potentially valuable material, such as production of bioethanol from fermentable
sugars that derived from lignocellulosic waste.
Oil-palm is one of the most active agricultural crops and becoming the most
attractive option for Malaysia due to great amount of agricultural waste being
produced every year (Goh et al., 2010), and it was reported that the largest portion of
total agricultural waste in Malaysia comes from oil palm plantation (Misson et al.,
2009). In the year 2007, Kelly-Yong et al. (2007) reported a total of 10,827 thousand
tons of trunks were generated based on the total area of 4,304,914 Ha oil palm
cultivation, where for every 25 years the chopped oil palm trees contribute to 2.515
3
tons of trunks generated from each hectare of oil palm cultivation. A report on the
performance of the Malaysian oil palm industry showed that the total oil palm
planted area was 4.85 million hectares in 2010 (Sulaiman et al., 2012). Due to the
availability of a wide plantation area, a significant amount of biomass could be
produced and further converted into value-added product. According to Goh et al.
(2010) , second generation of bioethanol was applicable on lignocellulosic materials
which can be used for bioethanol production to partly substitute fossil fuels in vehicle.
Due to the unpredictability of palm oil price in international market, the renewable
energy policy that proposed by Malaysian government is still not satisfactory even
though palm oil can be used as raw material supply to biofuel industry.
Lignocellulose is consisted of more than 60% of plant biomass generated on earth
which may be a potential feedstock for biofuels production, enzymes application and
other biochemical products (Tengerdy and Szakacs, 2003). Cellulose, hemicelluloses
and lignin are generally found intermeshed by strong chemical bondings, such as
non-covalent and covalent cross-linkages (Pérez et al., 2002). As lignocellulosic
biomass such as oil-palm trunk residues contains high amount of sugars in the form
of celluloses and hemicelluloses, it represents a promising feedstock for the
bioethanol production. Kosugi et al. (2010) found a large quantity of sap with the
high glucose content from the sap of the felled trunk. Two distinct components can
be differentiated in the oil-palm trunk residue, which are parenchyma and vascular
bundle (Akmar and Kennedy, 2001; Hashim et al., 2011). These two components are
found in almost equal proportions in the fiber residues (Hashim et al., 2011).
Therefore, ethanol fuel production from lignocellulosic biomass is advantageous as it
does not lead to competition for food resources (Lynd, 1996). Due to the renewable
4
and ever-present nature of lignocellulosic biomass without competition with food
crops, they are promising for bioethanol production and put a tremendous amount of
effort into the research aspect for bioconversion.
1.2 Microorganisms and their lignocellulolytic enzymes
The main products of lignocellulose degradation are sugar components which can be
used as a carbon or food source by numerous microorganisms. Fermentable sugars
can be produced mainly by fungi in its natural habitat via solid state fermentation
processes (Ibrahim, 2008). Many microorganisms including bacteria and fungi have
been found potentially to degrade cellulose and other plant cell wall fibres.
Lignocellulolytic enzymes-producing fungi are wide spread and most of them
contribute significantly to the decomposition of lignocellulosic residues in nature by
producing various lignocellulolytic enzymes (Dashtban et al., 2009; Sánchez, 2009).
Members of the Trichoderma genera such as T. viride, T. longibrachiatum, T. reesei
are notable for their high enzymatic productivity. On the other hand, genera of
Penicillium, such as P. verruculosum and P. funiculosum were reported for their
superior performance of cellulase preparations over Trichoderma enzymes (Gusakov,
2011). Several bacteria were also reported capable of producing lignocellulolytic
enzymes, such as Pseudomonas fluorescens, Escherichia coli, Bacillus subtilis and
Serratia marcescens (Sethi et al., 2013).
1.3 Biotechnological applications
Lignocellulose degradation is of utmost prominence for biotechnological conversion
of lignocellulosic materials into value-added products, which fostering the
development and application of enzymatic processes by a wide range of industries in
5
recent decades due to their high specificity, fast in action and often save raw
materials, energy and chemicals (Jegannathan and Nielsen, 2013). Various
bioproducts from the lignocellulose-degrading microorganisms and their applications
are widely reported (Kuhad and Singh, 1993; Subramaniyan and Prema, 2002; Sun
and Cheng, 2002), as lignocellulolytic enzymes that produced could be applied in
various industries including textile, detergents, pulp and paper, fodder, bioconversion,
environment, food, chemical and pharmaceutical.
1.4 Objectives of research
The objectives of the current research are as follow:
1) To isolate and to screen potential fungal isolates that can degrade the oil-palm
trunk residues by submerged fermentation system.
2) To characterize the potential oil-palm trunk residues-degrading fungal isolate
by morphological study and molecular level identification.
3) To characterize the crude lignocellulolytic enzymes produced by the selected
potential fungal isolate.
4) To purify the key enzymes for hydrolysis of oil-palm trunk residues.
1.5 Scope of study
Potential oil-palm trunk residues-degrading fungus was isolated from soil samples
which were collected from oil palm plantation areas in northern part of Peninsular
Malaysia. In these studies, laboratory fungal isolates were also used for determining
production of lignocellulose-degrading enzymes as well. Capability of the fungal
isolates with high potential in degrading oil-palm trunk residues by crude enzyme
were further investigated based on their protein production and specific activities on
6
oil-palm trunk residues by submerged fermentation. The fungal isolates with high
capability of degrading the oil-palm trunk residues were selected for further study.
Fungal isolate identification was based on the morphology of the fungal culture by
comparing their physical characteristics (mechanism of spore forming, size and shape
of spores) to those of reference materials, via microscopic observation. The selected
fungal isolates were also identified using molecular biological protocol by DNA
amplification and sequencing of the internal transcribed spacer (ITS) region.
Microscopic observations of changes on the oil-palm residues’ structure were
investigated. These observations included the analyses of sugars released during the
hydrolysis of lignocellulosic materials, which give insight information on enzyme
mechanisms at an ultrastructural level. Enzymes are unstable molecules with a
definite physic-chemical organization. Characterizations of the crude enzyme were
conducted on the basis of pH and temperature parameters. The effect of lignin on the
hydrolysis of oil-palm trunk residues was investigated and this was due to the
hindrance of lignin which might affect the efficiency of enzymatic hydrolysis on
lignocellulosic materials. Next, the saccharification experiments were carried out by
using crude enzyme produced by the selected potential fungal isolate compared to
commercial enzymes such as Celluclast 1.5L and Accellerase 1500. Since the
enzymes are proteinaceous in nature, standard extraction and purification procedures
for enzymes were the same as those used for proteins. It is included the purification
of enzymes by ammonium sulfate precipitation, extraction of enzyme, dialysis,
column chromatography and electrophoresis to obtain homogenous purified protein
fraction. Synergism effect of added purified enzymes together with commercial
enzymes was also investigated in the saccharification experiment.
7
CHAPTER TWO
LITERATURE REVIEW
2.1 Lignocellulosic biomass
Lignocellulosic biomass in the form of agricultural and forestry residues are the most
abundant and boundless or renewable natural resources. Lignocellulose mostly
consists of lignin, hemicelluloses and celluloses, in which can be derived from
woody and non-woody plants. Due to their valuable lignocellulosic chemical
properties, agricultural biomasses are the substrate of paramount importance of
biotechnological value. Improvement in many processes related to lignocellulose
biotechnology has gained great interest in pass few years.
2.1.1 Structure and composition of lignocellulosic biomass
The chemical components of lignocellulosic biomass varies from source to source
(Sitton et al., 1979; Sreenath et al., 1999; Lynd et al., 2005; Reddy and Yang, 2005),
which the major components comprise cellulose (35–50%), followed by
hemicellulose (20–35%), lignin (10–25%) as well as minimal fraction of protein,
essential oils and ash. The structural components of lignocellulose are illustrated in
Figure 2.1 and Figure 2.2. These complex and native structural components of
biomass are generally contributing to the availability and resistance for enzymatic
hydrolysis. Lignin polysaccharide matrix is usually found bounding surround
cellulose fiber, whereas the structural integrity of cell walls is attributed to the
formation of both covalent and non-covalent linkages from xylan. The composition
of hardwoods and softwoods are significantly different. The lignin content of
softwoods is generally higher than that of hardwoods, whereas hemicelluloses
8
Figure 2.1: Diagrammatic illustration of the framework of lignocellulose (Menon and Rao, 2012)
9
Figure 2.2: Schematic structural formula cellulose (glucan), hemicelluloses (homoxylan) and lignin (core lignin) (Chundawat et al., 2011).
10
content of hardwoods is higher than the softwoods. The compositions of various
lignocellulosic materials are illustrated in Table 2.1.
Table 2.1: Composition of representative lignocellulosic materials. Feedstocks Carbohydrate composition (% dry wt) Reference
Cellulose Hemicellulose Lignin Alfalfa 21.8 12.4 9.7 Dijkerman et al.
(1997) 33 18 8 Sreenath et al. (1999) Barley straw 38.08 22.63 22.27 Kim et al. (2011) 38.0 21.9 17.3 Kim et al. (2014) Bamboo 46.68 16.4 17.66 Kuttiraja et al. (2013) 38.4 25.9 20.8 Littlewood et al.
(2013) Banana waste (pseudo stem)
54-60 8.2-16 12-21 Santa-Maria et al. (2013)
44.32 22 9.66 Gabhane et al. (2014) Banana waste (leaf)
27-34 11-19 25-26 Santa-Maria et al. (2013)
32.56 12 21.80 Gabhane et al. (2014) Banana waste (pith)
36.14 7 16.43 Gabhane et al. (2014)
Corn cob 35-39 38-42 4.5-6.6 Okeke and Obi (1994) 45 35 15 Howard et al. (2003) 36.4 34.9 14.8 Sahare et al. (2012) Corn stover 39 19.1 15.1 Lee (1997) 37.5 22.4 17.6 Mosier et al. (2005b) 34.1 20.4 11.6 Jin et al. (2011) 41.7 20.5 18 Merino and Cherry
(2007) Cotton stalk 31.1 10.7 30.1 Silverstein et al.
(2007) 30.58 16.85 29.99 Kaur et al. (2012) Coffee pulp 24.0 8.9 19.4 Dijkerman et al.
(1997) Douglas fir 50 17.8 28.3 Lee (1997) 47.3 19.5 30.3 Kumar et al. (2012) Rice straw 41 21.5 9.9 Lee (1997) 32.1 24 18 Howard et al. (2003) Rice husk 34.4 17.5 23 Yáñez et al. (2006) 42.2 18.47 19.4 Banerjee et al. (2009) 47.3 23.0 29.7 Takahashi et al.
(2014) Wheat straw 33-40 20-25 15-20 McKendry (2002) 35-39 22-30 12-16 Prasad et al. (2007) 37-41 27-32 13-15 Balat (2011)
11
Table 2.1 Continued…… Newspaper 40-55 24-39 18-30 Howard et al. (2003) 40-55 25-40 18-30 Balat (2011) 40 23 20 Lee (1997) Sugarcane bagasse
38.1 26.9 18.4 Lee (1997)
25-45 28-32 15-25 Singh et al. (2009) 34.1 29.6 19.4 Maeda et al. (2011) Sunflower stalks
38.5 33.5 17.5 Sharma et al. (2002)
32.56 20.73 13.32 Díaz et al. (2011) 33.45 21.71 14.26 Ruiz et al. (2013) Olive tree biomass
34.4 20.3 20.4 Ruiz et al. (2006)
25.0 15.8 18.8 Cara et al. (2008) 25.0 15.8 16.6 Manzanares et al.
(2011) 33.96 17.86 18.56 López-Linares et al.
(2013) Switchgrass 30-50 10-40 5-20 McKendry (2002) 45 31.4 12 Howard et al. (2003) 31 20.4 17.6 Mosier et al. (2005b) 31 22 18 Merino and Cherry
(2007) Softwood 35-40 25-30 27-30 McKendry (2002) 42 27 28 Balat (2011) Hardwood 45-50 20-25 20-25 McKendry (2002) 45 30 20 Balat (2011) Oat straw 35.0 28.2 4.1 Gomez-Tovar et al.
(2012) Nut shells 25-30 25-30 30-40 Howard et al. (2003) Sorghum straw
35.1 24.0 25.4 Vázquez et al. (2007)
44.51 38.62 6.18 Poonsrisawat et al. (2013)
12
Composition of the same lignocellulosic materials might be different substantially
depending on the source of the lignocelluloses and the specific species variety (Van
Dyk and Pletschke, 2012). On the other hand, particular sugar analysis methods
(Foyle et al., 2007), analysis procedures such as pretreatment methods as well as the
growing location and harvesting season (Agblevor et al., 2003; Kaur et al., 2012;
Van Dyk and Pletschke, 2012) for one particular crop had put the great impact on the
compositional changes or differences from the same lignocellulosic biomass. For
instance, hydrolysis of switchgrass was impacted substantially by the time of harvest
as reported by Wyman et al. (2011). Therefore, compositional analysis for one
particular lignocellulosic biomass should be analyzed independently for each
conducted experimental unit as recommended by Van Dyk and Pletschke (2012).
2.1.1.1 Cellulose
The celluloses are complex molecules consisting of homopolymer of glucose units
linked with β-1,4-glucosidic units. These complex celluloses composed of linear β-
1,4-glucan chains which form aggregation of microfibrils (3 to 5 nm in diameter) via
interaction of intra- and intermolecular hydrogen bonds and van der Waals forces
resulting from pyranose ring stacking (Chundawat et al., 2011). Glucose and
cellodextrins are the products when cellulose is hydrolyzed. Depending on the
different type of sources, the degree of polymerization of cellulose might range from
100 to 10,000 (O'Sullivan, 1997; Somerville et al., 2004; Chundawat et al., 2011)
and even might reach up to 15,000 (Béguin and Aubert, 1994; Jørgensen et al., 2007).
For instance, cellulose in nature consisted of glucopyranose chain ranging from
10,000 units in wood, whereas 15,000 glucopyranose units was found polymerized in
native cotton (Agbor et al., 2011). Laureano-Perez et al. (2005) reported that the
13
‘straightness’ of the chain was determined by the hydrogen bonding within a
cellulose microfibril. On the other hand, the crystallinity and amorphous regions of
cellulose were determined by interchain hydrogen bonds. The degree of crystallinity
contributes to the low saccharification of cellulose due to existence of steric
hindrance from the cellulose molecule itself causing resistance to microbial attack
and enzymatic hydrolysis, whereas amorphous cellulose is less resistant to
degradation (Jørgensen et al., 2007; Menon and Rao, 2012). A dense layer of water
was formed surround the hydrophobic surface of cellulose which might contribute to
the obstruction for the enzymes to interact with substrate.
2.1.1.2 Hemicellulose
Hemicelluloses are matrix polysaccharides or heteropolymers, mostly constructed
from both hexoses and pentoses such as arabinose, galactose, glucose, mannose and
xylose (Brown, 1983; Bastawde, 1992; Rubin, 2008). The cellulose fibres are linked
together with these matrix polysaccharides into microfibrils and cross-linked with
lignin to create the complex linkages that provide structural strength (van Wyk,
2001). β-1,4-xylans are the second most abundant element of hemicelluloses for
lignocellulosic biomass. They make up around 20-30% of the dry weight of tropical
hardwood and annual plants, which are accounted for one-third of the renewable
biomass available on earth (Dhiman et al., 2008).
Xylans can be derived from different plants and grasses, which generally have the
same backbone structure of β-(1-4) linked xylose residues. The degree of branching
can be attributed by the origin of the sources and the differences in the branched
residues, which might compose of D-glucuronic acid, L-arabinose and 4-O-
14
methylesters of D-glucuronic acid (Bajpai, 2009). Bastawde (1992) reported that
fewer acidic side chains were found available in softwood xylans compared to
hardwood xylans. Softwood xylans comprise one acidic group of per nine to 12 D-
xylose residues, whereas there are per five to six D-xylose units found in hardwood
xylans. The major hemicellulose components in softwood are mannan-based, and
those in hardwood are xylan-based.
The major constituents of the xylans compose of D-xylose and L-arabinose. On the
other hand, the mannans are made up of D-glucose, D-galactose and D-mannose.
Collectively, the principal sugar components of these hemicellulose
heteropolysaccharides are: D-xylose, D-glucose, L-arabinose, D-mannose, D-
galactose, D-glucoronic acid, D-galactouronic acid, 4-O-methyl- D-glucoronic acid,
and a minority of L-rhamnose, L-fucose and various O-methylated sugars. They
usually have degree of polymerization of 100 to 200 (Jørgensen et al., 2007), and
side chain can be acetylated (Bastawde, 1992; Kuhad et al., 1997). The mannan
hemicelluloses, galactoglucomannans and glucomannans, in softwoods and
hardwoods, are both branced heteropolysaccharides. Their backbones are constructed
by both β-D-mannopyranose and 1,4-linked β-D-glucopyranose units. The acetyl
group partially replaces the C-2 and C-3 position of the mannose and glucose in the
backbone.
The hemicelluloses have been described as the most thermo-chemically sensitive
components of lignocellulosic biomass (Hendriks and Zeeman, 2009), and suggested
to ‘coat’ cellulose-fibrils within plant cell walls. To enhance the effectiveness of
cellulose digestibility, not less than 50% of hemicelluloses should be eliminated has
15
been suggested. Hence, comprehensive steps should be taken into account upon what
type of pretreatment is used, so that the formation of degradation products from
hemicelluloses such as furfurals and hydroxymethyl furfurals can be avoided. These
compounds have been reported to impede the fermentation process (Palmqvist and
Hahn-Hägerdal, 2000a-b).
2.1.1.3 Lignin
Lignins are highly complex network molecules consisted of phenyl-propane-based
monomeric units linked together by different types of bonds, including alkyl-aryl,
alkyl-alkyl, and aryl-aryl ether bonds (Jørgensen et al., 2007), which provides
rigidity, support, and protection to the plants (Blanchette et al., 2004). The molecular
weight of lignins may be 100 kDa or more. The relative proportions of the three
cinamyl alcohol precursors mingled into lignin, i.e., coniferyl alcohol, p-coumaryl
and sinapyl alcohol (Arora and Sharma, 2010), vary not only with plant species
(Sanderson, 2011) but also with location of the lignins within the plant cell wall as
well as the plant tissues. Lignin is found at the highest concentration in the middle
lamella. On the other hand, it is found the most abundantly available in the secondary
walls of the vascular plants.
Two types of bonding: β-aryl ether and α-aryl ether are suggested hydrolysable in the
linkage of lignin (Adler, 1977). The predominant β-aryl ether type bond is more
resistant to cleavage. Under mild hydrolytic conditions, the cleavage of the ether
bond is exclusively restricted to the α-aryl ether type (Kirk, 1987 ). Lignin is usually
insoluble in water due to its characteristic as the ‘glue’ that combines the different
constituents of lignocellulosic biomass together. Microbial and chemical degradation
16
were found less efficient due to the embedded of polymer cellulose microfibrils
within lignin (Malherbe and Cloete, 2002; Jørgensen et al., 2007) by forming a
physical barrier that restricts accession of microbial enzymes and chemicals for
further degradation (Avgerinos and Wang, 1983; Mooney et al., 1998). Efficient
pretreatment methods need to be adopted for removal of lignin from different type of
feedstocks, so that enhancing biomass digestibility. Chang and Holtzapple (2000)
showed that lignin removal gradually able to improve the biomass digestibility
successfully. Other than this lignin as a physical barrier, toxicity effect to the
microorganisms from lignin derivatives, less efficient binding of cellulolytic
enzymes to lignin-carbohydrates complexes specifically, and non-specific adsorption
of lignocellulolytic enzymes to lignin are some of the unfavorable effects caused by
lignin components (Agbor et al., 2011).
2.2 Pretreatment of lignocellulosic biomass
Many factors influence the reactivity and digestibility of the cellulose fractions of
lignocellulose biomass. These factors include the porosity of the biomass materials,
lignin and hemicellulose content, crystallinity of cellulose, cellulose degree of
polymerization, substrates availability surface area, feedstock particle size, cell wall
thickness and change of substrate in accessibility with its conversion (Alvira et al.,
2010). For accomplishing the industrially satisfactory time frame, pretreatment is
required for destruction of the robust structure of lignocellulosic biomass prior to
utilization for the conversion to fermentable sugars by enzymatic hydrolysis, such as
for the biomass-to-ethanol conversion processes.
17
The objective of the pretreatment is to render biomass materials more accessible to
either chemical or enzymatic hydrolysis for efficient product generation. The goals of
the pretreatment are: to remove and separate hemicellulose from cellulose; to disrupt
and get rid of the lignin sheath; to collapse the hydrogen bonds that rendering the
crystallinity of cellulose; enable maximal coverage of cellulases to cellulose surface
area; and to increase the pore size of cellulose to facilitate the penetration of
hydrolysis agents (Gong et al., 1999; Haghighi Mood et al., 2013). Hence, an
efficient enzymatic hydrolysis with reduced energy consumption and a maximal
sugar recovery can be accomplished without any obstacles ( Yang and Wyman, 2008;
Zhu and Pan, 2010; Limayem and Ricke, 2012). The aims of pretreatment on
lignocellulosic biomass are depicted in Figure 2.3.
Figure 2.3: Schematic of aims of pretreatment on lignocellulosic biomass (Mosier et al., 2005b; Balat, 2011).
Lignin
Amorphous
Region
Crystalline
Region
Cellulose
Hemicellulose
Pretreatment
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An advanced pretreatment process not only to increases accessibility of one enzymes
but also to enhance the complete solubilization of the polymer to monomer sugars.
However several other factors should fulfill its criteria of feasibility: preserving
hemicellulose fractions, avoiding size reduction, reduced energy intensive, limiting
formation of inhibitors due to degradation products, and being cost-effective (Menon
and Rao, 2012). High yields for multiple crops, sites ages, and harvesting time,
minimum amount of toxic compounds derived from sugar decomposition during
pretreatment, fermentation compatibility, such as the choice of an organism
able to ferment pentoses in hemicellulose are several key properties to take into
consideration for low-cost and successful pretreatment process (Yang and Wyman,
2008; Alvira et al., 2010).
A multitude of diverse pretreatment technologies have been established during the
last decade due to the diverse nature of different biomass feedstocks (Alvira et al.,
2010; Menon and Rao, 2012), and its economic assessment and environmental
impact (Menon and Rao, 2012). A generalized classification of pretreatment groups
them into physical, chemical, biological and combinatorial pretreatments which is
termed as physico-chemical method. Some of the promising pretreatment methods
are summarized in Table 2.2, which describe the pros and cons of the selected
particular pretreatments by a few reviewed articles.
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Table 2.2: Most promising pretreatment technologies (advantages and disadvantages).
Method of pretreatment Advantages Limitation and disadvantages References Mechanical Reduce cellulose crystallinity, applicable for various
feedstocks, success at pilot scale High power consumption than inherent biomass energy, low sugar yield, high equipment cost
Menon and Rao (2012)
Increase accessible surface and pore size, decrease the crystalinity and degree of polymerization
High energy requirement Behera et al. (2014)
Mineral acids Hydrolysis of cellulose and hemicellulose, alter lignin structure, applicable for various feedstocks, high sugar yield, allow to reuse of chemicals, success at pilot scale
Hazardous, toxic and corrosive, formation of inhibitors, high equipment cost
Menon and Rao (2012)
Majorly allow to hydrolyze hemicelluloses (xylan) Formation of inhibitory compounds, such as furfural, hydroxymethyl furfural, formic acid, levulinic acid and phenolic compounds; processing cost increased for removal of these compounds
Behera et al. (2014)
High solubility of hemicellulose and lignin, high glucose yield Process for recovery of acids used is expensive, costly corrosion-resistant equipment is required, formation of high concentration of inhibitors
Badiei et al. (2014)
Alkali Removal of lignin and hemicellulose, increases accessible surface area, high sugar yield, applicable for various feedstocks, allow to reuse of chemicals, success at pilot scale
Long residence time, irrecoverable salts formed
Menon and Rao (2012)
Effective to remove acetyl groups from hemicelluloses, reduce steric hindrance of enzymes subsequently enhance cellulose digestibility
Irrecoverable salts formed Behera et al. (2014)
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Table 2.2 Continued……
Alteration of lignin structure and effective to remove lignin, applicable to various biomasses, high sugar yield, success in pilot scale, low amount of inhibitors are formed
Operational cost is high, catalysts used are expensive
Badiei et al. (2014)
Liquid hot water Removal of hemicelluloses making enzymes accessible to cellulose, high sugar yield, formation high concentration of inhibitors, success at pilot scale
Long residence time, less lignin removal, not allow to reuse of chemicals
Menon and Rao (2012)
Achieved maximum solubilization of hemicelluloses by >80%, applicable for different kinds of lignocellulosic material, low or no production of inhibitors
─ Haghighi Mood et al. (2013)
Removal of lignin, no any catalyst or chemical required, minimal of product degradation, no washing step or neutralization required, success at pilot scale
Incomplete removal of lignin Behera et al. (2014)
Organosolv Hydrolyze lignin and hemicelluloses, high sugar yield, allow to reuse of chemicals, applicable for various feedstocks, success at pilot scale
Solvents need to be drained, evaporated, condensed and reused, formation of inhibitors, high equipment cost
Menon and Rao (2012)
Remove lignin extensively and almost complete solubilization of hemicellulose, applicable for various feedstocks
Solvents need to be drained, evaporated, condensed and reused, high operational cost
Behera et al. (2014)
Removal of lignin and hydrolyze hemicelluloses, improve enzymatic degradability of cellulose, applicable to various biomasses with high potential sugar yield
Removal of solvent is needed, costly process
Badiei et al. (2014)
Wet oxidation Removal of lignin, dissolves hemicellulose and causes cellulose decrystallization
High equipment cost, not allow to reuse of chemicals
Menon and Rao (2012)
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Table 2.2 Continued……
Efficient to fractionate lignocellulosic materials by solubilization, and hydrolysis of hemicelluloses, removal of lignin, less inhibitory compounds generation
High capital cost, such as high cost of pressure equipment and the oxygen as well as catalyst used
Haghighi Mood et al. (2013)
Solubilization of hemicellulose, removal of lignin, and lignocellulosic materials are fractionated
─ Badiei et al. (2014)
Ozonolysis Reduces lignin content, no toxic residues, high sugar yield with low amount of inhibitor production
Large amount of ozone required, not allow to reuse of chemicals, high equipment cost
Menon and Rao (2012)
Reduces lignin content, applicable to various feedstocks, process is carried out at room temperature and normal pressure, no formation of any inhibitory compound
─ Behera et al. (2014)
Reducing lignin content, applicable to various feedstocks, lack of degradation product, occurring at ambient temperature and normal pressure
Large amount of ozone required, costly process and less applicable
Xu and Huang (2014)
CO2 explosion Hemicellulose removal, cellulose decrystallization, cost-effective, high sugar yield with low inhibitor formation
Does not modify lignin, not allow to reuse of chemical, high equipment cost
Menon and Rao (2012)
Effective to solubilize lignin, low cost process, no generation of toxic, use of low temperature and high solid capacity
High cost of equipment Behera et al. (2014)
Steam explosion Hemicellulose removal and alteration in lignin structure, high sugar yield, applicable to various feedstocks, success at pilot scale
Incomplete destruction of lignin-carbohydrate matrix, high equipment cost, formation of inhibitory compounds
Menon and Rao (2012)
Applicable to various feedstocks, low energy requirement, no recycling or environment cost, cost effective
Less effective to softwood, incomplete disruption of the lignin-carbohydrate matrix, generation of inhibitory compounds
Behera et al. (2014)
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Table 2.2 Continued……
Increased removal of hemicelluloses from solid fraction, limited use of chemicals, low energy input
Incomplete destruction of lignin-carbohydrate matrix, formation of fermentation inhibitors at higher temperature
Xu and Huang (2014)
Ammonia Fiber Expansion Pretreatment (AFEX)
Removal of lignin and hemicelluloses, high sugar yield with low amount of inhibitory compounds formed, allow to reuse of chemicals
Not efficient for biomass with high lignin content, high equipment cost
Menon and Rao (2012)
Changes of biomass structure and enhance the digestibility, applicable to various feedstocks, no formation of any toxic materials, reduction of particle size is not required, allow to reuse of chemicals
Not efficient for biomass with high lignin content
Behera et al. (2014)
Induces the disruption in lignin-carbohydrate linkage, lower moisture content, lower sugar degradation products, almost complete recovery of solid material
─ Xu and Huang (2014)
Ionic liquids Dissolution of cellulose, increased amenability to cellulase, applicable to various feedstocks, low amount of inhibitor formed, allow to reuse of chemicals
─ Menon and Rao (2012)
Reduction of cellulose crystallinity, less dangerous process condition and chemicals, mild operational conditions and ionic liquid are easily recycled, lower energy consumption
Lead to inactivation and unfolding of the protein, high temperature cause some negative side-effects, such as deteriorated stability of ionic liquids, and occurrence of side-reactions
Haghighi Mood et al. (2013)
Applicable to various feedstocks, allow selective extraction of unaltered lignin and cellulose, process under ambient temperature without alkaline or acidic reagents required and inhibitor compound formation
Costly solvent are required, lower biomass conversion efficiency
Badiei et al. (2014)
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2.2.1 Physical pretreatments
2.2.1.1 Mechanical comminution
Most pretreatment approaches require the collected biomass to undergo some degree
of mechanical size reduction prior to introduction into a pretreatment reactor. The
goal of the mechanical pretreatment is to increase the available surface area and
decrystallize the cellulose components. Hence, this helps to increase the enzymatic
digestibility of the comminuted biomass and further reducing the degree of
polymerization (Alvira et al., 2010). A combination of grinding, chipping or milling
can be conducted to enhance the accessibility of enzymes to substrates, which
depending on the final particle size of the material (10–30 mm after chipping and
0.2–2.0 mm after milling or grinding) (Sun and Cheng, 2002; Kumar et al., 2009).
Due to higher energy consumption in milling stage, the use of very small particles is
unfavorable, and subsequently imposes negative effect on the following pretreatment
method. The amount of energy consumption that used for lignocellulosic biomass
pretreatment by mechanical comminution depends on the final particle size and
biomass characteristics (Cadoche and López, 1989; McMillan, 1994). As shown in
Figure 2.4, the power requirements increase rapidly with decreasing particle size.
The intensive mechanical energy requirements causes comminution to be cost-
prohibitive for use as a individual pretreatment in a biomass to ethanol conversion
process ( Alvira et al., 2010; Talebnia et al., 2010; Agbor et al., 2011; Balat, 2011).
2.2.1.2 Pyrolysis
Pyrolysis is a pretreatment method, in which lignocellulosic materials treated at
temperatures greater than 300°C and cellulose rapidly decomposes to produce
residual char and gaseous products under this condition (Sun and Cheng, 2002;
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Figure 2.4: Energy requirements for ball milling municipal solid waste (McMillan, 1994).
Pasha and Rao, 2009; Balat, 2011). The decomposition is time-consuming and less
volatile products are formed during the process at lower temperatures. Fan et al.
(1987) reported that a total of 80–85% conversion of cellulose to reducing sugars
with more than 50% glucose was achieved by applying mild acid hydrolysis (1 N
H2SO4, 97°C, 2.5 h) pretreatment on biomass residues. Pyrolysis can be boosted up
with the existence of oxygen (Shafizadeh and Bradbury, 1979), which further
resulted in depolymerization, oxidation, and dehydration during the process. The
conversion of cellulose to glucose yield through enzymatic degradation was
successfully improved by pyrolysis pretreatment of ground material (Leustean, 2009).
2.2.1.3 Extrusion
In this context, extrusion is a novel and promising physical pretreatment method for
biomass conversion to ethanol production. Physical and chemical modifications were
