development of a fungal cellulolytic enzyme ...

DEVELOPMENT OF A FUNGAL CELLULOLYTIC ENZYME COMBINATION FOR USE IN BIOETHANOL PRODUCTION USING HYPARRHENIA SPP AS A SOURCE OF FERMENTABLE SUGARS By

THEMBEKILE NCUBE

Thesis submitted in fulfilment of the requirements for the degree of

DOCTOR OF PHILOSOPHY in MICROBIOLOGY in the FACULTY OF SCIENCE AND AGRICULTURE (School of Molecular and Life Sciences) at the UNIVERSITY OF LIMPOPO

SUPERVISOR: Prof I. Ncube CO-SUPERVISORS: Prof R. L. Howard Prof E. Abotsi Prof. E. L. Jansen van Rensberg

2013

DECLARATION I declare that the thesis hereby submitted to the University of Limpopo for the degree of Doctor of Philosophy in Microbiology has not been previously submitted by me for a degree at this or any other university; that it is my work in design and execution; and that any material herein contained has been duly acknowledged.

___________________________

_______________________

i

Dedication

This work is dedicated to my beloved husband, Lee Ncube and my children Tumelo Simphiwe, Thapelo Musa, and Thabo Busiso for their patience, love, prayers and encouragement during the entire time of my study.

ii

Acknowledgements

I would like to express my sincere gratitude to the following people and institutions: •

Prof. I. Ncube for his mentoring and unwavering support throughout the study.

•

Profs. Howard, Abotsi and. van. Rensburg for their contributions in various aspects of the project.

•

Dr. P. van Zyl for an opportunity to do part of the research at the CSIR fermentation laboratory.

•

The BMBT technical staff especially Mrs. B. Kekana, Mr. B. Moganedi, Mr. M. Mokgobedi and Mr. Z. Simayi for their technical assistance.

•

National Research Foundation for the the Africa doctoral scholarship.

•

Vlaammse Interuniversitaire Raad (VLIR- UOS) Own initiative project and the Interuniversity Council for funding the project.

•

The office of the Dean of the Faculty of Science and Agriculture, University of Limpopo, for assistance with tuition fees.

•

National University of Science and Technology (Zimbabwe) for providing financial supprt for tuition and subsitance.

•

Biotechnology students Manape Modiba, Reagile Maribeng, Emmanuel Rubagumya, Rumbidzai Rakumazondo, Kgomotso Mokatse for assistance and moral support.

•

My entire family for love, support and prayers throughout.

iii

Abstract

The current study investigated four fungal species namely Aspergillus niger FGSC A733, Aspergillus versicolor EF23, Penicillium citrinum AZ01 and Trichoderma harzianum NCGR 0509 for their abilities to produce cellulases and xylanases in submerged and solid state fermentations. Five different substrates (carboxymethyl cellulose, xylan, common thatch grass, wheat bran and Jatropha curcas seed cake) were examined for their potential use as low cost feedstock for fermentation by the fungal species. Aspergillus niger FGSC A733 produced the highest titres of cellulase and xylanase in solid state fermentations using wheat bran as a substrate. However, because of the need to lower the cost of enzyme production, Jatropha seed cake a relatively underutilised oilseed cake was used.

Supplementation of the Jatropha seedcake with 10% common thatch grass (Hyperrhenia sp) resulted in a fivefold increase in the levels of xylanase produced. Cellulase production was not affected by this supplementation. Addition of ammonium chloride increased production of xylanase while cellulase production was not affected nitrogen supplementation. Maximum xylanase was produced on Jatropha seed cake at 25 °C after 96 hours while cellulase was maximally produced at 40 °C after 96 hours of solid state fermentations. Peak production of xylanase was obtained at an initial pH of 3 whilst cellulase was maximally produced at an initial pH of 5. The crude xylanase was most active at pH 5 and cellulase at pH 4. The optimum temperature for cellulase activity was 65 °C and that of xylanase was 50 °C. Under optimized conditions, 6087 U/g and 3974 U/g of xylanase and cellulase per gram of substrate used were obtained respectively.

The diversity of cellulases was investigated so as to determine the most appropriate enzyme mixture for saccharification of the common thatch grass. Proteins from the four species under investigation were partially purified by affinity chromatography on swollen Avicel. The proteins were analysed using sodium dodecyl sulphate-polyacrylamide gel electrophoresis iv

SDS-PAGE and zymography. Potential cellulase bands from SDS-PAGE were sequenced by mass spectrometry. The basic logical alignment tool (BLAST) and Clustal W were used for matching and identifying the sequences with closely related ones in the databases. The identified proteins from Penicillium citrinum AZ01 and Aspergillus versicolor EF23 were found to closely resemble a catalytic domain of cellobiohydrolase from Trichoderma sp. The three proteins obtained from Aspergillus niger showed resemblance to 1,4-beta glucan cellobiohydrolase A precursor from Aspergillus niger FGSC A733 was also found to have cellobiase and endoglucanase activity was determined using cellobiase and carboxymethyl cellulose as substrates. Cellulase and xylanase zymograms of proteins from A. niger FGSC A733 demonstrated six active bands ranging from 20 kDa to 43 kDa for cellulase and a 31 kDa active band for xylanase.

The cellulase produced by Aspergillus niger FGSC A733 on Jatropha seed cake under optimised conditions was used for saccharification of 2% (w/v) common thatch grass (CTG) in combination with Celluclast™. Celluclast™ and Aspergillus niger cellulase were mixed at different ratios and the amount of glucose produced over time was monitored using high performance liquid chromatography (HPLC). A ratio of 2 volumes Celluclast™ to one volume Aspergillus niger cellulase was chosen for the saccharification process. The main enzymes in the mixture were identified using peptide mass fingerprinting as endoglucanases from the Celluclast™ and cellobiase from the Aspergillus niger cellulase. Concentration of the Celluclast™ tenfold times (164 FPU) improved the yield of glucose by 42.8 and 37.8% in acid and alkali pre-treated CTG, respectively. Concentrating Aspergillus niger cellulase (13.2 FPU) decreased the production of glucose by 4.8% in acid pre-treated CTG while in alkali pre-treated CTG, a 5% increase in glucose production was observed. Increasing the substrate loading of acid pre-treated CTG from 2% to 10% (w/v) resulted in a two and a half times increase in glucose production while an increase of 1.5 g/l glucose was obtained from 7% (w/v) alkali pre-treated CTG. Addition of xylanases from Aspergillus niger to the Celluclast™-Aspergillus niger cellulase mixture decreased glucose production by 16.3% on acid pre-treated CTG while there was an increase of 18.3% glucose in alkali pre-treated CTG. Addition of enzyme preparations from Aspergillus versicolor EF23, Penicillium citrium AZ01 and Trichoderma harzianum NCGR 0509 to the Celluclast™-Aspergillus niger cellulase mixture resulted in lower glucose production both in acid and alkali pre-treated v

CTG. Addition of Pentopan™ improved glucose production by 8 and 25% on 10% acid and 7.5% alkali loading of pre-treated CTG respectively. The optimal conditions for the production of the glucose rich hydrolysate in 10% (w/v) acid and 7% (w/v) alkali pre-treated CTG was found to be the use of Celluclast™-Aspergillus niger cellulase-Pentopan™ mixture (164 FPU Celluclast™ and 13 FPU Aspergillus niger cellulase, 7178 IU) Pentopan™ at 50 °C for 32 hours.

The fermentability of the glucose in glucose-rich CTG hydrolysates to ethanol using Saccharomyces cerevisae WBSA 1386 and Candida shehatae CSIR Y-0492 was investigated. The highest yield of ethanol produced by S. cerevisae WBSA 1386 was 9.8 g/l in the alkali pre-treated CTG hydrolysate and 8.7 g/l in acid pre-treated CTG. C. shehatae CSIR Y-0492 produced 9 g/l of ethanol in alkali pre-treated CTG within 48 hours while acid pre-treated CTG hydrolysate produced 8.8 g/l of ethanol within 24 hours of the fermentation process. Addition of the nutrient supplement boosted the ethanol yield in the acid pre-treated hydrolysates. The consumption of glucose during fermentation by S. cerevisae WBSA 1386 and C. shehatae CSIR Y-0492 on average was 97%. The C. shehatae CSIR Y-0492 was expected to produce much higher ethanol yield than the Saccharomyces because of its ability to utilize xylose for ethanol production. This however was not observed in this investigation. The conclusion of this study is that it is possible to produce bioethanol from Hyperrhenia spp. (CTG) using a combination of fungal enzymes for the production of fermentable sugars.

vi

TABLE OF CONTENTS Contents

Page

Declaration ........................................................................................................................... i Dedication ........................................................................................................................... ii Acknowledgements ............................................................................................................ iii Abstract .............................................................................................................................. iv Table of contents ............................................................................................................... vii List of tables .................................................................................................................... xxii List of figures ................................................................................................................. xxiii List of abbreviations ...................................................................................................... xxxi

1. INTRODUCTION.........................................................................................................1 1.1 Background information ................................................................................1 1.2 Motivation of study .........................................................................................4 1.3 Aims and objectives ........................................................................................5 1.3.1 Aims ...................................................................................................5 1.3.2 Objectives ..........................................................................................5 1.4 Research hypothesis ........................................................................................6 1.5 Significance of study .......................................................................................6

2. LITERATURE REVIEW ............................................................................................7 vii

2.1 Lignocellulose ..................................................................................................7 2.1.1 Lignin .................................................................................................8 2.1.2 Cellulose ............................................................................................8 2.1.3 Hemicelluloses ...................................................................................9 2.2 Lignocellulolytic enzymes .............................................................................10 2.2.1 Ligninases ........................................................................................11 2.2.1.1 Lignin peroxidase..............................................................12 2.2.1.2. Manganese peroxidase .....................................................12 2.2.1.3 Laccases ............................................................................12 2.2.1.4 Versatile peroxidases ........................................................12 2.2.1.5 Other lignases....................................................................13 2.2.1.6 Lignocellulolytic enzymes from microorganisms ............13 2.2.1.7 Applications of ligninases .................................................13 2.2.1.8 Lignin degradation ............................................................14 2.2.2 Cellulases .........................................................................................17 2.2.2.1 Mechanism of action of cellulases ....................................17 2.2.2.2 Production of cellulases ....................................................18 2.2.2.3 Cellulase sytem and control of cellulase gene expression 19 2.2.2.4 Regulation of cellulase synthesis ......................................20 2.2.2.5 Genetic engineering of cellulases .....................................20 2.2.2.6 Application of cellulases ...................................................21 2.2.2.7 Global cellulase market.....................................................22 viii

2.2.3 Hemicellulases .................................................................................22 2.2.3.1 Sources of xylanases .........................................................24 2.2.3.2 Biochemical properties of xylanases ................................25 2.2.3.3 Production of xylanases ....................................................26 2.2.3.4 Factors affecting xylanase yield........................................27 2.2.3.5 Xylanase from extremophilic regions ...............................28 2.2.3.6 Regulation of xylanase synthesis ......................................28 2.2.3.7 Application of xylanase ....................................................30 2.3 Processing of lignocellulosics to bioethanol ................................................32 2.3.1 Pre-treatment of lignocellulosics .....................................................32 2.3.1.1 Mechanical pretreatment ...................................................33 2.3.1.2 Pyrolysis ............................................................................33 2.3.1.3 Steam explosion (autohydrolysis) .....................................33 2.3.1.4 Ammonia fibre/freeze explosion (AFEX) ........................34 2.3.1.5 Ozone treatment ................................................................34 2.3.1.6 Organosolv ........................................................................35 2.3.1.7 Concentrated acid hydrolysis ............................................35 2.3.1.8 Dilute acid hydrolysis .......................................................36 2.3.1.9 Solid (super) acids.............................................................37 2.3.1.10 Alkaline hydrolysis .........................................................37 2.3.1.11 Biological treatment ........................................................38 2.3.1.12 Enzymatic hydrolysis ......................................................38 ix

2.3.2 Depolymerisation of cellulose and hemicellulose ...........................39 2.3.3 Fermentation of lignocellulosic hydrolysates ..................................39 2.2.3.1 Simultaneous saccharification and fermentation ..............40 2.2.3.2 Separate hydrolysis and fermentation ...............................41 2.2.3.3 Direct microbial conversion (DMC) .................................42 2.2.3.4 Microorganisms for ethanolic fermentations ....................43 2.3.4 Products and solids recovery ...........................................................45 2.4 Biofuels ...........................................................................................................45 2.4.1 Global demand for fuels...................................................................48 2.4.2 Energy overview in Africa ...............................................................49 2.5 Bioethanol production ...................................................................................50 2.5.1 Feedstock for bioethanol production ...............................................51 2.5.1.1 Sucrose containing feedstocks ..........................................52 2.5.1.2 Starchy materials ...............................................................52 2.5.1.3 Lignocellulosic biomass.....................................................53 2.5.1.4 High value products from lignocellulosics .......................53 2.6 Solid state fermentations ..............................................................................55 2.6.1 General aspects of solid state fermentations ....................................56 2.6.2 Microorganims used in solid state fermentations ............................56 2.6.3 Solid state fermentation bioreactors.................................................57 2.6.3.1 Tray reactors .....................................................................58 2.6.3.2 Packed bed reactors...........................................................58 x

2.6.3.3 Horizontal drum reactors ..................................................58 2.6.3.4 Fluidised bed reactors .......................................................59 2.6.4 Applications and advantages of solid state fermentations ...............59 2.7 Jatropha carcus ..............................................................................................59

3. MATERIALS AND METHODS ...............................................................................61 3.1 Materials ........................................................................................................61 3.1.1 Lignocellulosic substrates ................................................................61 3.1.2 Microbial strains ..............................................................................61 3.1.3 Reagents ...........................................................................................61 3.1.4 Equipment ........................................................................................62 3.2 Methods ..........................................................................................................62 3.2.1 Screening of fungi for cellulolytic and xylanolytic activity ............63 3.2.1.1 Maintenance of fungal cultures .........................................63 3.2.1.2 Identification of fungal strains ..........................................63 3.2.1.3 Preparation of fermentation substrates..............................63 3.2.1.4 Pre-treatment of common thatch grass .............................64 3.2.1.5 Culturing of fungal strains for production of inoculums ..64 3.2.1.6 Production of cellulolytic enzyme sin submerged fermentations.....................................................................65 3.2.1.7 Production of cellulolytic enzymes in solid state fermentations.....................................................................65 xi

3.2.1.8 Extraction of enzymes from solid state fermentation fungal cultures ...................................................................65 3.2.1.9 Assaying for cellulase and xylanase .................................66 3.2.1.10 Preparation of standard curve for glucose and xylose determination .......................................................66 3.2.2 Optimization of conditions for production of xylanase and cellulase by A. niger FGSC A733 on Jatropha seed cake in solid state fermentation ...................................................................66 3.2.2.1 Culturing of A. niger FGSC A733 for cellulase and xylanase production ..........................................................67 3.2.2.2 Effect of adding common thatch grass on the production of cellulase and xylanase by A. niger FGSC A733 ..................................................................................67 3.2.2.3 Determination of optimum temperature for cellulase and xylanase production by A. niger FGSC A733............67 3.2.2..4 Effect of intitial pH on cellulase and xylanase production by A. niger FGSC A73 ...................................67 3.2.2.5 Effect of nitrogen source on cellulase and xylanase production by A. niger FGSC A733 ................................68 3.2.3 Partial characterization of cellulase and xylanase from the fungal strains ...............................................................................................68 3.2.3.1 Determination of optimum temperature for cellulase xii

and xylanase activity .........................................................68 3.2.3.2 Optimum pH for cellulase and xylanase activity ..............68 3.2.3.3 Ammonium sulphate precipitation of the crude enzyme extract ................................................................................68 3.2.3.4 Sodium dodecyl sulphate polyacrylamide gel electrophoresis (SDS-PAGE) for A. niger FGSC A733 enzyme ..............................................................................69 3.2.3.5 Zymography for cellulase and xylanase ............................70 3.2.4 Determination of the diversity of cellulases in the enzyme extracts of the fungal strains ..........................................................................70 3.2.4.1 Filter paper assay ..............................................................70 3.2.4.2 Cellobiase assay and determination of the amount of glucose released by cellobiase ..........................................71 3.2.4.3 Caboxymethylcellulose assay for detection of endoglucanases .................................................................71 3.2.4.4 Preparation of phosphoric acid cellulose (Walseth cellulose) ...........................................................................72 3.2.4.5 Cellulose affinity chromatography ...................................72 3.2.4.6 Preparation of samples for protein sequencing .................72 3.2.4.7 Tryptic digestion of the protein bands ..............................73 3.2.4.8 Mass spectrometry for peptides from gel slices ................73 xiii

3.2.4.9 Data analysis .....................................................................74 3.2.5 Enzymatic saccharification of lignocellulosic material ...................74 3.2.5.1 Proximate analysis of lignocellulosic material .................74 3.2.5.2 Production of cellulase and xylanase for saccharification of lignocellulosic materials .....................75 3.2.5.3 Preparation of enzymes for saccharification of lignocellulosic materials ..................................................75 3.2.5.4 Sachharification of common thatch grass .........................76 3.2.5.5 Effect of substrate loading on the production of glucose from common thatch grass ................................................76 3.2.5.6 Production of glucose from common thatch grass under optimized conditions .........................................................77 3.2.5.7 Analysis of sugars from the hydrolysis mixture ...............77 3.2.6 Ethanol fermentation and quantification ..........................................78 3.2.6.1 Maintainace and propagation of yeasts .............................78 3.2.6.2 Fermentation of acid and alkaline common thatch grass hydrolysates for ethanol production..................................79 3.2.6.3 Quantification of ethanol ..................................................79 4. RESULTS .....................................................................................................................81 4.1 Utilisation of different substrates for production of cellulase and xylanase by selected fungal strain on submerged and solid state xiv

fermentations ..................................................................................................80 4.1.1 Utilisation of different substrates for production of cellulase by selected fungal strains on submerged fermentations .......................81 4.1.2 Utilisation of different substrates for production of cellulase by selected fungal strains on solid state fermentations .........................83 4.1.3 Utilisation of different substrates for production of xylanase by selected fungal strains on submerged fermentations .......................84 4.1.4 Utilisation of different substrates for production of xylanse by selected fungal strains on solid state fermentations .........................86 4.2 Production and characterization of xylanase from four fungal strains ...89 4.2.1 Time course for cellulase production by the four fungal strains .....89 4.2.2 Time course for xylanase production by the four fungal strains .....90 4.2.3 Optimum temperature for activity of cellulase from four fungal strains ...............................................................................................91 4.2.4 Optimum temperature for xylanase activity of enzymes produced by the four fungal strains ................................................................92 4.2.5 Optimum pH for the cellulase activity of the four fungal strains ....93 4.2.6 Optimum pH for activity of xylanase of the four fungal strains ......94 4.3 Production and characterization of the cellulase from Aspergillus niger FGSC A733 ........................................................................................96 4.3.1 Time course analysis for the production of cellulase and xylanase .96 xv

4.3.2 Effect of adding common thatch grass to Jatropha seed cake during the production of cellulase....................................................96 4.3.3 Effect of adding common thatch grass to Jatropha seed cake during the production of xylanase....................................................97 4.3.4 Optimum temperature for cellulase production by Aspergillus niger FGSC A733 ............................................................................98 4.3.5 Optimum temperature for xylanse production by Aspergillus niger FGSC A733 ............................................................................99 4.3.6 Effect of nitrogen source on the production of cellulase ..............100 4.3.7 Effect of nitrogen source on the production of xylanase ..............101 4.3.8 Effect of initial pH on the production of cellulase by Aspergillus niger FGSC A733 .......................................................102 4.3.9 Effect of initial pH on the production of xylanase by Aspergillus niger FGSC A733 .......................................................103 4.3.10 Optimum temperature and pH for cellulase and xylanase activity..........................................................................................104 4.4 Diversity of cellulases in fungal species.....................................................105 4.4.1 Total cellulase, endoglucanase and cellobiase activity from crude enzymes of the fungal strains .........................................................105 4.4.2 Cellulase and xylanse zymography of Aspergillus niger FGSC A733 crude enzyme preparation .....................................................106 xvi

4.4.3 Gel chromatography for crude and partially purified enzymes from the fungal strains ...................................................................107 4.4.4 Matrix assisted laser desorption ionization time of flight analysis of partially purified cellulase bands obtained from SDS-PAGE gels for the four fungal strains .......................................................111 4.5 Enzymatic hydrolysis of Hyparrhenia sp. for the production of fermentable sugars for ethanol ..................................................................116 4.5.1 Enzymatic saccharification of lignocellulose for the production of fermentable glucose ...................................................................117 4.5.2 Saccharification of Avicel using Celluclast™ and Aspergillus niger cellulase preparation ..............................................................117 4.5.3 Sacharification of untreated and pre-treated common thatch grass using Celluclast™ and Aspergillus niger cellulase preparation ....119 4.5.4 Effect of concentrating the Celluclast™-Aspergillus niger cellulase mixture on the production of glucose .............................126 4.5.5 Effect of adding xylanase on the production of glucose on acid and alkali pre-treated common thatch grass ...........................128 4.5.6 Effect of substrate loading on the production of glucose ..............133 4.5.7 Effect of enzyme preparations from other fungal strains...............135 4.6 Production of glucose under optimized conditions ..................................141 4.7 Ethanolic fermentation of saccharified common thatch grass xvii

hydrolysate...................................................................................................142 4.7.1 Ethanolic fermentation of acid and alkaline pre-treated hydrolysates ...................................................................................143 4.7.2 Yield factor for production of ethanol by Candida shehatae CSIR Y-0492 and Sacchormyces cerevisae WBSA 1386 .............151 5. DISCUSSION ............................................................................................................153 5.1 Production of cellulases and xylanases by selected fungal strains on submerged and solid state fermentations .................................................152 5.1.1 Solid state and submerged fermentations ......................................153 5.1.2 Comparison of substrates for production of cellulase and xylanase.........................................................................................154 5.1.3 Time course analysis of the fungal strains grown on Jatropha seed cake .......................................................................................156 5.1.4 Optimum temperatures for cellulase and xylanase activities for Apergillus niger FGSC A733, Aspergillus versicolor EF23, Penicillium citrinum AZ01 and Trichoderma harzianum NCGR 0509 ...................................................................................156 5.1.5 Optimum pH of activity for the fungal strains ...............................157 5.2 Optimization of cellulose and xylanase production by Aspergillus niger FGSC A733 on Jatropha seed cake ..................................................158 5.2.1 Selection of the substrate and organism for the optimization xviii

process............................................................................................159 5.2.2 Time course for production of cellulase and xylanase...................159 5.2.3 Effect of supplementation of Jatropha seed cake with common thatch grass on cellulase and xylanase production .......................160 5.2.4 Optimum temperature for production of cellulase and xylanase ...160 5.2.5 Influence of nitrogen supplementation on the production of cellulase and xylanase ....................................................................161 5.2.6 Effect of initial pH on the production of cellulase and xylanase ..162 5.2.7 Optimum pH and temperature for cellulase and xylanase activity 163 5.2.8 Production of cellulase and xylanase under optimized conditions 164 5.2.9 Cellulase and xylanase zymograms ...............................................164 5.2.10 Diversity of cellulases and xylanases from the fungal strains .....164 5.3 Proximate analysis of common thatch grass and Jatropha seed cake ..165 5.4 Production of glucose from lignocellulolytic materials by different cellulase and xylanase combinations .........................................................166 5.4.1 Enzyme cocktails ...........................................................................166 5.4.2 Production of glucose from Avicel ...............................................166 5.4.3 Production of glucose by cellulase mixtures on untreated, acid and alkaline pre-treated common thatch grass ..............................167 5.4.4 Effect of Celluclast™ concentration on the saccharification of common thatch grass......................................................................169 xix

5.4.5 Effect of Aspergillus niger cellulase concentration on the saccharification of common thatch grass .......................................167 5.4.6 Effect of Celluclast™ and Aspergillus niger cellulase concentration on the production of glucose ...................................168 5.4.7 Effect of adding Pentopan™ on the saccharification of common thatch grass.....................................................................................170 5.4.8 Effect of adding other fungal cellulases on the saccharification of common thatch grass......................................................................170 5.4.9 Effect of adding Aspergillus niger cellulase on the saccharification of common thatch grass .......................................171 5.4.10 Effect of increasing substrate concentration on the saccharification of common thatch grass ....................................171 5.5 Production of ethanol from common thatch grass hydrolysate by Candida shehatae WBSA 1386 and Saccharomyces cerevisae CSIR Y0492 ............................................................................................................172 6. CONCLUSION ..........................................................................................................174 Recommendations ..............................................................................................175 7. REFERENCES ..........................................................................................................176 Appendix 1 ..........................................................................................................193 Appendix 2 ..........................................................................................................200

xx

LIST OF TABLES

Table 2.1: Comparative production of bioethanol by different feedstock ......................51 Table 4.1: Proximate analysis of Jatropha seed cake .....................................................88 Table. 4.2: Activity of different cellulases found in Celluclast™ and four fungal strains ...........................................................................................................106 Table 4.3. Proximate analysis of Hyparhhenia sp ........................................................116 Table 4.4

Yield factor for Candida shehatae CSIR Y-0492 based on glucose as the substrate and ethanol as the product ..................................................151

Table 4.5: Yield factor for Saccharomyces cerevisae WBSA 1386 based on glucose as the substrate and ethanol as the product ..................................................152

xxi

LIST OF FIGURES

Figure 2.1: Structure of linear anhydropyranose units .......................................................9 Figure 2.2: Lignin degradation by white rot fungi............................................................16 Figure 2.3: Separate hydrolysis and fermentation with separate pentose and hexose and combined sugar fermentation .................................................................42 Figure 2.4: Generalised process stages in lignocellulose waste bioconversion ................55 Figure 4.1: Production of cellulase by A. niger FGSC A733, A. versicolor EF23, P. citrinum AZ01 and T. harzianum NCGR 0509 during submerged fermentations on CTG, xylan, Jatropha seed cake and wheat bran .............82 Figure 4.2: Production of cellulase by A. niger FGSC A733, A. versicolor EF23, P. citrinum AZ01 and T. harzianum NCGR 0509 during solid state fermentations on CTG, xylan, Jatropha seed cake and wheat bran ............ 84 Figure 4.3: Production of xylanase by A. niger FGSC A733, A. versicolor EF23, P. citrinum AZ01 and T. harzianum NCGR 0509 during submerged fermentations on common thatch grass, xylan, Jatropha seed cake and wheat bran ......................................................................................................85 Figure 4.4: Production of xylanase by A. niger FGSC A733, A. versicolor EF23, P. citrinum AZ01 and T. harzianum NCGR 0509 during solid state fermentations of grass, Jatropha seed cake and wheat bran ..........................87 Figure 4.5: Time course of cellulase production by A. niger FGSC A733, A. versicolor EF23, P. citrinum AZ01 and T. harzianum NCGR 0509 during solid xxii

state fermentation of Jatropha seed cake.......................................................90 Figure 4.6: Time course of xylanase production by A. niger FGSC A733, A. versicolor EF23, P. citrinum AZ01 and T. harzianum NCGR 0509 during solid state fermentation of Jatropha seed cake.......................................................91 Figure 4.7: Optimum temperature for cellulase activity of A. niger FGSC A733, A. versicolor EF23, P. citrinum AZ01 and T. harzianum NCGR 0509grown on Jatropha seed cake in SSF......................................................92 Figure 4.8: Optimum temperature for activity of xylanase from A. niger FGSC A733, A. versicolor EF23, P. citrinum AZ01 and T. harzianum NCGR 0509 grown on Jatropha seed cake in SSF.............................................................93 Figure 4.9: Optimum pH for activity of crude cellulases obtained from A. niger FGSC A733, A. versicolor EF23, P. citrinum AZ01 and T. harzianum NCGR 0509 cultures during solid state fermentation of Jatropha seed cake ..............................................................................................................94 Figure 4.10: Optimum pH for activity of crude xylanase obtained from A. niger FGSC A733, A. versicolor EF23, P. citrinum AZ01 and T. harzianum NCGR 0509 enzymes produced in SSF of Jatropha seed cake ...............................95 Figure 4.11: Effect of supplementing Jatropha seed cake with common thatch grass on production of cellulase by A. niger FGSC A733 during solid state fermentation on Jatropha seed cake ....................................................97 Figure 4.12: Effect of supplementing Jatropha seed cake with common thatch grass on production of xylanase by A. niger FGSC A733 during solid xxiii

state fermentation on Jatropha seed cake ....................................................98 Figure 4.13: Optimum temperature for production of cellulase by A. niger FGSC A733 during SSF on Jatropha seed cake and 10% (w/w) common thatch grass...................................................................................................99 Figure 4.14: Optimum incubation temperature for xylanase production by A. niger FGSC A733 during SSF on Jatropha and 10% (w/w) common thatch grass ...........................................................................................................100 Figure 4.15: Influence of supplementation of Jatropha seed cake with nitrogen on the production of cellulase by A. niger FGSC A733during solid state fermentation on Jatropha seed cake ..................................................101 Figure 4.16: Influence of supplementation of Jatropha seed cake with nitrogen on the production of xylanase by A. niger FGSC A733during solid state fermentation on Jatropha seed cake ..................................................102 Figure 4.17: Effect of initial pH on production of cellulase during SSF of Jatropha seed cake by A. niger FGSC A733 ...........................................................103 Figure 4.18: Effect of initial pH on production of xylanase during SSF of Jatropha seed cake by A. niger FGSC A733 ..........................................................104 Figure 4.19 Zymograms for crude cellulase and xylanase produced by A. niger FGSC A733...............................................................................................107 Figure 4.20: SDS-page gel for crude enzyme and partially purified cellulases from fungal isolates ...........................................................................................108 Figure 4.21: Cellulase zymogram for fungal strains obtained from the partially xxiv

purified cellulase versus the crude extract ................................................109 Figure 4.22: SDS-page gel of fungal cellulase obtained from partially purified cellulases versus the crude extract fungal isolates ...................................110 Figure 4.23: Clustal W sequence alignment of peptides obtained from a 106 kDa protein from Penicillium citrium AZO1 ..................................................111 Figure 4.24: Clustal W sequence alignment of peptides obtained from a 73 kDa protein

from Aspergillus versicolor EF23 ............................................112

Figure 4.25: Clustal W sequence alignment of peptides obtained from a 66 kDa protein from Aspergillus niger FGSC A 733 ...........................................113 Figure 4.26: Clustal W sequence alignment of peptides obtained from a 58 kDa protein from Aspergillus niger FGSC A 733 ...........................................114 Figure 4.27: Clustal W sequence alignment of peptides obtained from a 40 kDa protein from Aspergillus niger FGSC A 733 ............................................115 Figure 4.28: Enzymatic saccharification of Avicel to glucose using different Celluclast™ and Aspergillus niger cellulase mixtures .............................118 Figure 4.29: Enzymatic saccharification of 2% (w/v) acid pre-treated CTG suspension using different Celluclast™ preparation and Aspergillus niger cellulase mixtures......................................................................................119 Figure 4.30: Enzymatic saccharification of 2% (w/v) alkali pre-treated CTG suspension using different Celluclast™ and Aspergillus niger cellulase mixtures....................................................................................................120 Figure 4.31: Enzymatic saccharification of 2% (w/v) untreated CTG suspension xxv

using different Celluclast™ and Aspergillus niger cellulase mixtures .....121 Figure 4.32: Effect of varying the concentration Celluclast™ on the saccharification of 2% acid pre-treated CTG suspension to glucose ..................................122 Figure 4.33: Effect of varying the concentration of Celluclast™ on the saccharification of 2% alkali pre-treated CTG to produce glucose ..........123 Figure 4.34

Effect of varying the concentration of Aspergillus niger cellulase on the saccharification of 2% acid pre-treated CTG to glucose ...............124

Figure 4.35: Effect of varying the concentration of Aspergillus niger cellulase on the saccharification of 2% acid pre-treated CTG to glucose. .................................................................................................125 Figure 4.36: Effect of varying the concentration of Celluclast™ and Aspergillus niger cellulase on the saccharification of 2% acid pre-treated CTG to glucose ..................................................................................................126 Figure 4.37: Effect of Celluclast™ and Aspergillus niger cellulase concentration on the saccharification of 2% alkali pre-treated CTG to glucose ..................127 Figure 4.38: Effect of adding Pentopan™ to Celluclast™-Aspergillus niger cellulase on the production of glucose on 2% acid pre-treated CTG ......................129 Figure 4.39: Effect of adding Pentopan to a Celluclast™-Aspergillus niger cellulase mixture, on the production of glucose on 2% alkali pre-treated CTG ......130 Figure 4.40: Effect of adding Aspergillus xylanase to a Celluclast™-Aspergillus cellulase mixture on the production of glucose on 2% acid pre-treated CTG...........................................................................................................131 xxvi

Figure 4.41: Effect of adding Pentopan to a Celluclast™-Aspergillus cellulase mixture on the production of glucose on 2% alkali pre-treated CTG ......132 Figure 4.42: Effect of substrate loading on the saccharification of acid pre-treated CTG to glucose .........................................................................................133 Figure 4.43: Effect of substrate loading on the saccharification of alkali pre-treated CTG to glucose on the production of glucose on acid pre-treated CTG ..134 Figure 4.44: Effect of adding A. versicolor EF23 enzyme preparation to the Celluclast™- Aspergillus niger cellulase mixture on the production of glucose on acid pre-treated CTG ..............................................................135 Figure 4.45: Effect of A. versicolor EF23 enzyme preparation to the Celluclast™ -Aspergillus niger cellulase mixture .............................136 Figure 4.46: Effect of adding P. citrinum AZ01 enzyme preparation to the Celluclast™-Aspergillus niger cellulase mixture on the production of glucose on acid pre-treated CTG ..............................................................137 Figure 4.47: Effect of adding P. citrinum AZ01 enzyme preparation to the Celluclast™-Aspergillus niger cellulase mixture on the production of glucose on alkali pre-treated CTG ............................................................138 Figure 4.48: Effect of adding T. harzianum NCGR 0509 enzyme preparation to the Celluclast™-Aspergillus niger cellulase mixture on the production of glucose ..................................................................................................139 Figure 4.49: Effect of adding T. harzianum NGCR 0509 enzyme preparation to the Celluclast™-Aspergillus niger cellulase mixture on the production xxvii

of glucose ..................................................................................................140 Figure 4.50: Production of glucose under optimized conditions ...................................141 Figure 4.51: Ethanol production and glucose consumption from an alkaline CTG treated hydrolysate without nutritional supplementation by S. cerevisae WBSA 1386 .............................................................................................143 Figure 4.52: Ethanol production and glucose consumption from an alkaline CTG treated hydrolysate with nutritional supplementation by S. cerevisae WBSA 1386 .............................................................................................144 Figure 4.53: Ethanol production and glucose consumption from acid CTG treated hydrolysate without nutritional supplementation by S. cerevisae WBSA 1386 .............................................................................................145 Figure 4.54: Ethanol production and glucose consumption from acid pre-treated CTG hydrolysate with nutritional supplementation by S. cerevisae WBSA 1386 ..............................................................................................146 Figure 4.55: Ethanol production and glucose consumption from an alkaline pre-treated CTG hydrolysate without nutritional supplementation by C. shehatae CSIR Y-0492....................................................................147 Figure 4.56: Ethanol production (○) and glucose consumption from an alkaline pre-treated CTG hydrolysate with nutritional supplementation by C .shehatae CSIR Y-0492.........................................................................148 Figure 4.57: Ethanol production and glucose consumption from acid pre-treated CTG hydrolysate without nutritional supplementation by C. shehatae xxviii

CSIR Y-0492 ............................................................................................149 Figure 4.58: Ethanol production and glucose consumption from acid pre-CTG hydrolysate with mineral supplementation by C. shehatae CSIR Y-0492 ......................................................................................................150

xxix

LIST OF ABBREVIATIONS

APS

ammonium persulphate

α

alpha

β

Beta

BCA

Bicinchoninic acid

EDTA

ethylenediamine tetra acetic acid

CaCl 2

Calcium chloride

CMC

carboxymethylcellulose

CMCase

carboxymethycellulase

CTG

common thatch grass

°C

degrees centigrade

DNS

3,5-Dinotrosalicyclic acic

Exo

outer

Endo

inner

Fig.

Figure

FGSC

Fungal Genetic Stock Centre

GC

Gas chromatograph

HCl

hydrochloric acid

HPLC

High performance liquid chromatography

KH 2 PO 4

potassium dihydrogen phosphate

K 2 HPO 4

di-potassium dihydrogen phosphate xxx

kDal

kilo Dalton

L

litre

M

molar

mM

millimolar

mL

milliliter

mg

milligram

min

minute

MgSO 4

magnesium sulphate

nkat

nano kat

NaCl

Sodium chloride

NaOH

Sodium hydroxide

NaNO 3

sodium nitrate

NaNO 3

Sodium azide

%

percentage

PAGE

polyacrylamide gel electrophoresis

rpm

revolutions per minute

RID

refractive index detector

SDS

Sodium dodecyl sulphate

sp.

species

SmF

Submerged fermentation

SSF

Solid state fermentation

TEMED

N,N,N,N Tetramethyl-ethylenediamine xxxi

µl

microliter

µg

microgram

w/v

weight per volume

v/v

volume per volume

V

volts

YMP

yeast, malt and peptone

YPD

yeast, peptone, dextrose medium

xxxii

CHAPTER 1

INTRODUCTION 1.1

Background information

The world’s present economy is highly dependent on fossil fuels which are still the primary energy source as they contribute more than 80% of the world’s energy. Among the various fossil fuels, oil is the most highly consumed (38% of the world consumption), followed by coal (26%) and natural gas (23%) for industrial, commercial, household and transportation purposes (Sriroth et al., 2010; Sarkar et al., 2012). Although there are reports of oil reserves worldwide, the world’s demand for fossil fuels exceeds the supply, hence the increase in oil prices (Sriroth et al., 2010). The drastic increase in oil prices, finite nature of fossil fuels, increasing concerns regarding the environmental impact (especially the green house emissions) as well as health and safety considerations have sparked interest in alternative cheap renewable energy sources (Malherbe and Cloete, 2002; Balat, 2011). An alternative fuel must be technically feasible, economically competitive, environmentally acceptable and readily available (Balat, 2011). Numerous potential alternative fuels have been proposed including bioethanol, biodiesel, methanol, hydrogen, boron, natural gas, liquefied petroleum gas (LPG), and solar energy (Sarkar et al., 2012). Biomass based fuels also known as biofuels are among the alternative fuels that are being considered. Biofuels, mainly from lignocellulosic materials, have been considered in that; lignocellulosics are abundant and evenly distributed geographically as compared to the fossil fuels, generate low net greenhouse gas emissions, minimize the conflict between land use for food and feed production as well as energy feedstock production and may provide employment for rural areas (Hahn-Hägerdal et al., 2006; Balat, 2011; Sarkar et al., 2012). The global phenomenon has been to burn this waste as the cheapest way of disposing of it. This, however, leads to increased pollution. The increased concern over the security of the non-renewable fuel and the negative impact of fossil fuels on the environment have recently stimulated an interest in optimizing the fermentation process for large scale production of biofuels such as ethanol. Current industrial 1

processes for ethanol production use sugarcane (Southern hemisphere) and corn (Northern hemisphere) (Chandrakant and Bisara, 1998; Howard et al., 2003). The world bioethanol production was 31 billion litres in 2001, 39 billion litres in 2006 and shot up to 85 billion litres in 2010. It is expected to reach 100 billion litres by 2015 (Sarkar et al., 2012, www.globalrfa.com). It has been estimated that 442 billion litres can be produced from lignocellulosic material and 491 billion litres can be obtained from total crop residues and wasted crops (Sarkar et al., 2012). The biofuel growth scenario aims to replace 15% of gasoline production with biofuels by 2015 and 20% by 2020 throughout most of the world (Chen and Qiu, 2010). Brazil has utilized ethanol since 1925 while the USA used gasohol or oxygenated fuels since the 1980’s. These gasoline fuels use 10% ethanol by volume. The demand for ethanol is likely to rise significantly with the plans by automobile industries to manufacture flexi-fuelled engines which use 85% ethanol and 15% gasoline by volume (Balat, 2011). The current bioethanol production based on corn, starch and sugar crops compete directly with the food and feed sectors. Moreover, production of bioethanol from foods and feed products is not economical thus making bioethanol more expensive than the fossil fuels. Materials such as corn stover, crop straws, sugar bagasse, switchgrass, forestry residues, short rotation woody crops, waste paper and other municipal and industrial wastes are potential feedstock for ethanol production (Dawson and Boopathy, 2007; Dermibas, 2007; Prasad et al., 2007). The common barrier to lignocellulose degradation is the chemical composition of the lignocellulosic structure. Generally, the materials contain cellulose, hemicellulose and lignin. Pre-treatment of the lignocellulosic material to delignify the material is necessary for the ease of access to cellulose and hemicellulose which have to be depolymerized by enzymes. A lot of work has been done on lignification and methods include hydrolyzing the substance by treating with either acid or alkali (Martín et al., 2002; Dawson and Boopathy, 2007; Prasad et al., 2007). The method of choice for pre-treatment may depend on the hardness of the material as well as the target end product. It is, however, important to keep the pre-treatment costs low in order to ensure a financially viable process. Microorganisms and their enzymes have been studied for the bioconversion of the lignocellulosic material into fermentable sugars at maximum production and the lowest 2

possible cost. Studies have been conducted into lignocellulases from microbial sources such as bacteria and fungi (Lopez et al., 2007; Valavoska et al., 2007; Maciel et al., 2008). The three main classes of enzymes; ligninases, hemicellulases and cellulases which have varying biochemical properties have been identified under different research conditions for hydrolysing different lignocellulosic to fermentable sugars (Tengerdy and Szakacs, 2003). Enzyme selection has to balance the efficient conversion of the lignocellulosic material while keeping the entire production process at minimal cost. Currently, the economics of ethanol production from lignocellulosics shows that the cost of cellulolytic enzymes is the major contributor (30-49%) of the net production costs (Chen and Qiu, 2010). According to an estimate, cellulase alone contributes 22.5-43% of the total cost of cellulosic ethanol production when enzymes are procured from commercial sources (Singhania et al., 2010). However, there are reports that the cost for a litre of cellulosic ethanol has come down by 72% between 2008 and 2012 (www.about.bnef.com). The demand for more stable, highly active and specific enzymes is growing rapidly. The global world market for industrial enzymes was valued at US$3.1 billion in 2009 and reached about US$ 3.6 billion in 2010. The estimated market for 2011 was US$3.9 billion and the market is projected to grow by an annual compounded growth rate of 9.1% to reach US$6 billion by 2016 (www.marketresearch.com). The majority of the world’s supply of enzymes is from Europe, USA and Japan. About 75% of enzymes produced are hydrolases with the cellulases accounting for approximately 20% of the world enzyme market in the last decade. The cost of production and low yields of the enzymes are the major problems for industrial application of the enzymes in industrial processes (Chen and Qiu, 2010; Singhania et al., 2010). There is a need for cost effective production of enzymes that can be used for important processes such as production of biofuels from lignocellulosics. Therefore, the ability of microbial strains to utilize inexpensive substrates to produce cellulase and hemicellulase needs investigation. The reduction in cost of enzyme production may lead to a breakthrough in the commercialisation of lignocellulosic ethanol. This study essentially aims to investigate the use of locally isolated fungal strains to utilize Jatropha curcas seed cake (an agricultural waste product) in the production of cellulases and xylanases for saccharification of abundant Hyparrhenia sp. (ordinary thatch grass) in the production of bioethanol.

3

1.2

Motivation for the study

The production of ethanol from lignocellulosics has received much attention due to the immense potential for the conversion of this renewable biomaterial into biofuels and other useful chemicals. Currently, the major producers of bioethanol are Brazil and the US which account for 62% of the world production. In this regard, biofuel production is mainly from sugarcane, sugar beet and other food crops such as maize. The 2007/2008 food crisis across the world was mainly caused by farmers switching to production of cash food crops for biofuels. Utilisation of food crops is undesirable and unsustainable, hence the need to investigate the use of other non-food materials. Also, there is a need for clean alternative energy sources since the primary energy from fossil fuels is not sustainable and the overuse of fossil fuels is the main contributor to global warming. Biofuel technologies are relevant to both the industrialized and developing countries for energy security, protection of the environment, foreign exchange savings and socioeconomic benefits related to the rural sector. Developing countries have a comparative advantage for biofuel production because of greater availability of land, favourable climatic conditions for agriculture and lower labour costs (Guerra-Perez et al., 2003). The Hyparrhenia sp. used in this study has advantages in that it is widely abundant in South Africa and grows naturally in the veld. Except for low level usage as thatching and as an animal feed in the early growing season and after fires, it is not used much by local communities. The mature grass is unpalatable to livestock and is often burnt as a way of getting rid of it. This contributes to air pollution. This investigation aims to develop technologies around the use of this grass for the production of bioethanol using suitable enzymes from selected fungal strains and yeasts for ethanolic fermentation. The Jatropha curcas seed cake is a by-product of biodiesel production and could be a suitable substrate for the production of the cellulolytic enzymes. The Jatropha seed cake is rich in proteins but has toxic compounds such as phorbol esters which pose a threat to the environment. Utilization of the Jatropha seed cake will produce value added products in the form of lignocellulolytic enzymes and may help reduce the toxicity of the cake for safer disposal.

4

The fungal strains used for enzyme production have been isolated at locations from the natural South African and Zimbabwean environments. This may allow for development of unique lignocellulosic degrading enzyme combinations.

1.3

AIMS AND OBJECTIVES

1.3.1 Aim The aim of the project is to develop Hyparrhenia spp into a bioethanol feedstock through efficient saccharification using a locally developed lignocellulolytic enzyme combination consisting of commercial cellulases and cellulases from fungal strains isolated in southern African environments.

1.3.2 Objectives The specific objectives are to: •

Screen fungal strains in order to select hyper-degraders (based on cellulase and hemicellulase activities) of different lignocellulosic materials.

•

Establish an inexpensive substrate for cultivation of fungi and production of lignocellulosic enzymes.

•

Develop a system for production of lignocellulolytic enzymes on an inexpensive substrate

•

Determine the chemical composition of the Hyparrhenia spp. (common thatch grass) and Jatropha being.

•

Optimise conditions (temperature, pH, cofactors) for cellulase and xylanase production.

•

Investigate the enzyme (culture supernatant) cocktail with respect to degradation of cellulose and hemicellulose as indicated by production of reducing sugars. 5

•

Carry out combinatorial enzyme experiments so as to determine a combination that gives the most extensive degradation of Hyparrhenia spp to monomeric sugars.

•

Characterize the enzyme combination involved in the saccharification of the common thatch grass.

•

Evaluate the effectiveness of enzyme combination in a two-stage process consisting of saccharification of grass and fermentation of the resulting sugars to bioethanol.

1.4

Research hypothesis

Selected fungal strains have the capability to produce lignocellulolytic enzymes (ligninases, hemicellulases and cellulases) that can degrade Hyparrhenia to different extents to produce sugars necessary for bioethanol production singularly or in combination.

1.5

Significance of the study

Results from this study may lead to production of enzymes from local strains by a simple inexpensive fermentation process. Optimization of enzyme production may lead to the production of high enzyme titres. The use of enzyme cocktails may lead to efficient utilization of waste lignocellulosic material for the production of fermentable sugars that can be used for production of bioethanol. The study will also contribute to scientific knowledge about the production and characteristics of cellulase and xylanase as well as production of biofuels from grass.

6

CHAPTER 2 LITERATURE REVIEW 2.1

Lignocellulose

Lignocellulose is the major component of biomass making up about 50% of the matter produced by photosynthesis. Lignocellulose consists of three types of polymers which are cellulose, hemicellulose and lignin and these components make up 90% of lignocelluloses. (Howard et al., 2003; Martínez-Herrera et al., 2006; Balat, 2011). The lignocellulose polymers are strongly intermeshed and chemically bonded by non-covalent and covalent cross linkages. Lignocelluloses are a major component of wood, grass, agricultural residues and municipal solid wastes. Large amounts of lignocellulose waste are generated by the agricultural and forestry practices as well as the paper and pulp industries (Howard et al., 2003; Mussatto and Teixeira, 2010) Agricultural crops such as wheat, rice, and corn produce straw, cobs, stalks and husk as residues. The residues are normally used as animal feed, compost, soil conditioner and sometimes are burnt as a fuel. Processing of corn, wheat, rice and soyabean produces wastewater and bran. Most of this waste is used as animal feed. Fruit and vegetable processing results in the production of seeds, peels, wastewater, husks, shells, stones, rejected whole fruit and juice. The waste generated from the fruit and vegetable processing is used as animal and fish food and the seeds are used for oil extraction (Howard et al., 2003). Sugarcane produces bagasse which is burnt as a fuel. Saw and plywood waste results in woodchips, shavings and saw dust which are utilised by the pulp and paper industry and also for production of chip and fibre board. The pulp and paper industry produces sulphite liquor and this is re-used in the pulp and board industry as a fuel. Lignocellulose wastes from the community include paper, cardboard, disused furniture and animal wastes. A small percentage of the paper, cardboard and disused furniture is recycled, the rest is burnt. Animal wastes are used as soil conditioners (Howard et al., 2003).

7

2.1.1

Lignin

Lignin (15-25% of total dry matter) is the most abundant aromatic polymer in nature and it confers structural support, impermeability and resistance against microbial attack and oxidative stress (Crawford and Crawford, 1984; Hammel, 1997; Mussatto and Teixeira, 2010). Lignin is a recalcitrant amorphous heteropolymer which is non-water soluble, optically inactive heterogeneous phenylpropanoid polymer that contains a diverse range of stable C-C bonds and aryl ether linkages. The lignins from most hardwoods and softwoods contain a predictable complement of phenylpropanone units derived from coniferyl, sinapyl and p-coumaryl alcohol monomers (Hammel, 1997; Brunow, 2001; Dashtan et al., 2010). The lignin contents on a dry basis in softwoods and in hardwoods range from 20 % to 40 % by weight and 10 % to 40 % in various herbaceous species such as bagasse, corncobs, peanut shells, rice hulls and straws (Balat et al., 2008). Grass lignins are complexed with significant quantities of esterified phenylpropanoid acids (Crawford and Crawford, 1984).

2.1.2

Cellulose

Cellulose is the most common organic polymer representing about 1.5 x 1012 tons of the total biomass production through photosynthesis. Cellulose is considered an inexhaustible source of raw material for different products and is the most abundant biopolymer dominating waste material from agriculture (Sukumaran et al., 2005; Mathew et al., 2008). Cellulose is the major component of lignocellulose and makes up about 35-50% of the dry weight of wood (Mussatto and Teixeira, 2010). Cellulose is a linear polymer isomeric with starch and is composed of D-glucose sub-units linked by β-1,4 glycosidic bonds forming the cellobiose molecules. The glucose units are in six-membered rings called pyranoses that are joined by single oxygen atoms between C-1 of the pyranose ring and the C-4 of the next ring (Figure 2.1) (Martínez-Herrera et al., 2006; Balat, 2011).

8

Figure 2.1: Structure of linear anhydropyranose units (Martínez-Herrera et al., 2006).

Loss of water as the glycosidic bond is formed leads to the glucose units in the polymer being referred to as the anhydroglucose units. The terminal glucose residues forming the chain differ from each other. One contains a reducing hemiacetal group and is thus known as the reducing end group. The other contains an extra secondary hydroxyl group and is known as the non-reducing end group. These ends are present in native cellulose and after hydrolytic cleavage of the glycosidic bond. Determination of the end groups is used as a means of measuring the molecular weight of cellulose as well as following the course of the hydrolytic degradation (Dutta et al., 2008). The polymeric molecules of cellulose form long chains (elemental fibrils) linked together by hydrogen bonds and van der Waals forces. Hemicellulose and lignin cover the cellulose microfibrils. Cellulose can appear in a crystalline form which is called crystalline cellulose. There is also a small percentage of non-organized cellulose chains which form the amorphous cellulose.

2.1.3

Hemicelluloses

Hemicelluloses are a heterogeneous class of polymers which makes up 15-35% of the total dry weight of wood (Gírio et al., 2010). The amount of hemicellulose in a plant depends on type of tissue, stage of growth, growth environment, physiological conditions, and storage of the material as well the method of extraction. Differences also exist in the hemicellulose content and composition between stem, branches, roots and bark. Hemicellulose has a lower 9

molecular weight than cellulose. Hemicellulose is made up of branched-chain heteropolysaccharides containing hexosans and pentosans which are easily hydrolysed chemically and enzymatically to give simple sugars and some acetic acid (Martinez et al., 2005). Hemicelluloses include xylan, mannan, galactan and arabinan. The classification of the hemicellulose fractions depends on the sugar moieties present. The principal monomers present in most of the hemicelluloses are D-xylose and D-arabinose which are the pentosans, D-mannose, D-glucose, and D-galactose, the hexosans as well as 4-O-methyl-D-glucoronic acid and D-galactoronic acid (Beg et al., 2001; Gírio et al., 2010). The sugars are linked by the β-1,4 and occasionally β-1,3-glucosidic bonds. The principal component of hardwood hemicellulose is glucoronoxylan while glucomannan predominates in softwoods (Martinez et al., 2005). Xylan a pentosan made up of β-1,4-linked D-xylopyranosyl residues is the most abundant of all the hemicelluloses representing up to 30-35% of the total dry weight (Kulkarni et al., 1999; Knob et al., 2010). The xylopyranosyl backbone is substituted at positions C-2, C-3 and C-5 to varying degrees depending upon the plant and the developmental stage it is at . The modifications which can be found on xylan yield different types of xylan such as glucoroxylans (substituted at the C-2 postion by α-D-glucoronic acid), glucomannan, and galactoglucomannans (Tseng et al., 2002; Knob et al., 2010). Xylans seem to be interspersed, intertwined, and covalently linked at various points with the overlying sheath of lignin while producing a coat around underlying strands of cellulose via hydrogen bonding. The linkages of xylan to lignin and cellulose may be important in maintaining the integrity of cellulose in situ and help in protecting fibres against degradation by cellulases (Beg et al., 2001).

2.2

Lignocellulolytic enzymes

Lignocellulolytic enzymes are enzymes which degrade the lignin and cellulosic structures in plants. These include both cellulolytic and ligninolytic enzymes. The ability of the enzymes to biodegrade carbohydrates into value added products is the basis of current interest into biomass conversion and utilisation. While plant biomass is the single largest source of carbohydrates which are fermentable by microorganisms, the carbohydrates are not available 10

in free form. The monomers are present in an extensively polymerized nature. Each of these polymers is degraded by a variety of enzymes which work synergistically to achieve the breakdown. The enzymes are collectively known as the lignocellulolytic enzymes (Gray et al., 2006). Lignocellulolytic enzymes have numerous applications and biotechnological potential for industries such as chemical, fuel, brewery and wineries, animal feed, textiles and laundry as well as pulp and paper. (Howard et al., 2003). About 20% of the world’s sale of approximately US$3.9 billion industrial enzymes consists of cellulases, hemicellulases and pectinases. The world market for industrial enzymes is projected to grow by an annual compounded growth rate of 9.1% to reach US$6 billion by 2016 (www.bbcresearch.com).

2.2.1

Ligninases

2.2.1.1

Lignin peroxidases

Lignin peroxidases (LiPs) are enzyme that contain ferric heme glycoproteins and play a role in the biodegradation of lignin (Dashtan et al., 2010) The LiPs operate via a typical peroxidise catalytic cycle. The enzyme lignin peroxidase is oxidized by hydrogen peroxide to a two-electron deficient intermediate, which returns to its resting state by performing two electron oxidations of donor substrates (Grande and Domínguez de María, 2012). Lignin peroxidases are more powerful oxidants than typical peroxidases and therefore are capable of oxidizing substrates such as phenols, anilines, non-phenolic lignin structures and other aromatic ethers that resemble the basic structural unit of lignin (Hammel, 1997).

2.2.1.2

Manganese peroxidase

Manganese peroxidases (MnP) are extracellular glycoproteins and are secreted in multiple isoforms which contain one molecule of heme as iron protoporhyrin IX (Dashtban et al., 2010). Manganese peroxidases have Mn(II) as an obligatory electron donor for reduction of one electron deficient enzyme to its resting state and Mn(III) is produced as a result. The reaction requires the presence of bidentate organic acid chelators such as glycolate or oxalate 11

which stabilize Mn(III) and promote its release from the enzyme. The resulting Mn(III) chelates are small diffusible oxidants that can act at a distance from the MnP active site. The chelates are not strongly oxidizing thus they are not able to attack the recalcitrant nonphenolic structures that make up approximately 10% of lignin. The MnP reactions are limited in their degree of ligninolysis but the Mn(III) that these generate facilitates later attack by the bulkier more powerful oxidant LiP (Hammel, 1997; Dashtban et al., 2010).

2.2.1.3

Laccases

Laccases are glycosylated blue copper oxidases which use molecular oxygen to oxidize various

phenolics and other electron-rich substrates (Sanchez, 2009). Laccases contain

multiple copper atoms which are reduced as the substrates are oxidized (Sigoillot et al., 2012). After four electrons have been received by the laccase molecules and molecular oxygen is reduced to water, the laccase returns to the native state. In the presence of artificial auxiliary substrates such as 2,2'-azino-bis(3-ethylbenzothiazoline-6-sulphonic acid (ABTS), the effect of laccase can be enhanced so that it oxidizes non-phenolic compounds that otherwise would not be attacked (Call and Mücke, 1997; Hammel, 1997; Dashtan et al., 2010).

2.2.1.4

Versatile peroxidases

Versatile peroxidases (VPs) are glycoproteins with hybrid properties capable of oxidizing typical substrates of other basidiomycetes peroxidases including Mn(II) and also veratryl alcohol, MnP and typical LiP substrate respectively. The versatile peroxidases form an attractive lignocellulolytic enzyme group due to their dual oxidative ability to oxidize Mn(II) and also phenolic and non-phenolic aromatic compounds (Dashtban et al., 2010; Sigoillot et al., 2012).

12

2.2.1.5

Other ligninases

Other enzymes involved in lignin degradation are known as peroxide producing-enzymes. Aryl alcohol oxidases (AAO) provide a route for hydrogen peroxide production in some white rot fungi (Sigoillot et al., 2012). Glyoxal oxidase (GLOX) accepts a variety of 1-3 carbon aldehydes as electron donors. Some GLOX substrates e.g. glyoxal and methylglyoxal are natural extracellular metabolites of organisms such as Phanerochaete chrysosporium. Aryl alcohol dehydrogenases (AAD) provide another route for hydrogen peroxide, production in some white rot fungi. Quinine reductases (QR) are also involved in lignin degradation (Hammel, 1997; Sanchez, 2009).

2.2.1.6

Lignocelluloytic enzymes from microorganisms

There is a diverse spectrum of lignocellulolytic microorganisms, mainly fungi and bacteria that have been isolated and identified over the years. Despite the huge collection of lignocellulolytic microorganisms only a few have been studied extensively (Sigoillot et al., 2012). Trichoderma reesei and its mutants are widely employed for the commercial production of cellulases and hemicelluloses. T. reesei does not however produce ligninases. White-rot fungi belonging to basidiomyecetes are the most efficient and extensive lignin degraders. P. chrysosporium is the best studied lignin degrading fungus producing a cocktail of lignocelluloytic enzymes. The less known fungi such as Daedalea flavida, Phlebia fascicularia, P. floridensis and P. radiate have been found to selectively degrade lignin in wheat straw while leaving the other components intact. Less prolific degraders among the bacteria are those belonging to the genera Celluomonas, Pseudomonas, Clostridium thermocellum and that of the actinomyces Thermomonospora and Microbiospora are receiving attention as they might be representing a gene pool with possibly unique genes for genetic engineering of lignocellulose producers (Dashtan et al., 2010).

2.2.1.7

Application of ligninases

Ligninases from the white rot fungi can be used for degradation of persistent aromatic pollutants such as dichlorophenol, dinitrotoluene and anthracene. In the animal feed industry, 13

fibre degrading enzymes are used to improve feed utilisation, milk yield and body weight gain (Howard et al., 2003). As lacasses work efficiently on a broad range of substrates without cofactors. They may have significant value in many biotechnological applications such as pulp bio-bleaching, biosensors, food industries, textile industries, soil remediation and in the production of complex synthetic polymers in synthetic chemistry. Laccases are also suitable for wastewater treatment in the textile industry (Mayer and Staples, 2002; Arantes and Millagres, 2007; Dashtban et al., 2010). Manganese peroxidases increase the degree of dye decolourization and are important in the biobleaching of kraft pulps. The main drawback in the utilization of manganese peroxidases is the unavailability of the enzyme in large quantities. This can however be resolved by the use of deoxyribonucleic acid (DNA) recombinant technology (Job-Cei et al., 1996; Qian and Goodell, 2005). The commercial application of laccases is hampered by lack of sufficient enzyme stocks and the cost of the redox mediators. Heterologous expression of the enzymes with protein engineering allows for the cost effective creation of more robust and active enzymes. In addition, the improvement in immobilization methods would result in greater stability of laccases with long life times (Dashtan et al., 2010).

2.2.1.8

Lignin degradation

LiP and MnP oxidise the substrate by two consecutive one electron oxidation steps with intermediate cation radical formation. LiP degrades the non-phenolic lignin units (up to 90 %) of the polymer. The oxidation begins by abstraction of one electron from the donor substrates aromatic ring (a) (Figure 2.2). The resulting species, an aryl cation radical then undergoes a variety of post-enzymatic reactions. MnP generates Mn3+ which acts as a diffusible oxidizer on phenolic or non-phenolic lignin units via lipid peroxidation reactions (Sanchez, 2009). The aromatic radicals evolve in different non-enzymatic reactions including the C-4-ether breakdown (b) (Figure 2.2), aromatic ring cleavage (c) (Figure 2.2 Cα-Cβ breakdown (d) 14

(Figure 2.2) and demethoxylation. The aromatic aldehydes released from Cα-Cβ breakdown of lignin or synthesized de novo by the fungus (f, g) (Figure 2.2) are substrates for hydrogen peroxide generation by AAO in cyclic reactions involving AAD (Hammel, 1997; Sánchez, 2009). Phenoxy radicals from C4 ether breakdown (b) (Figure 2.2) can repolymerize on the lignin polymer (h) if they are not first reduced by oxidases to phenolic compounds (i). The phenolic compounds formed can again be reoxidized by laccases or peroxidase (j). Phenoxy radicals can also be subjected to a Cα-Cβ breakdown (k)( (Figure 2.2) yielding p-quinones. Quinones from g and/or k contribute to oxygen activation in recycling reactions involving oxygen activation with QR, laccases and peroxidases (l,m), resulting in reduction of the ferric iron present in wood (n), either by the superoxide cation radical or directly by the semiquinone radicals, and its oxidation with concomitant reduction of hydrogen peroxide to a hydroxyl free radical. The latter is a very mobile and strong oxidizer that can initiate the attack on lignin (p) in the initial stages of wood decay, when the small size of pores in the still intact cell wall prevents the penetration of lignocelluloytic enzymes. Then, lignin degradation proceeds by oxidative attack of the enzymes described above (Sanchez, 2009).

15

Figure 2.2:

Lignin degradation by white rot fungi (Sanchez, 2009).

16

The enzymes involved in lignin degradation lead to the formation of humic and fulvic acids which are subsequently rearranged, cleaved and partially mineralized (Valavoska et al., 2007). Other enzymes whose roles have not been fully elucidated include hydrogen peroxide producing microorganisms, glyoxal oxidase, glucose oxidase, vetratryl alcohol oxidases, methanol oxidase and oxidoreductase. The ligninases are too large to penetrate the unaltered cell wall of plants. There are suggestions that ligninases employ low-molecular weight diffusable reactive compounds to effect the initial changes to the lignin substrate (Howard et al., 2003).

2.2.2

Cellulases

Cellulase is a multi-component system generally composed of three major components. These are the endo-1,4 β-glucanase (endo-1,4-β-glucan 4 glucanohydrolase EC 3.2.1.4), cellobiohydrolase, (1.4-β-D-glucan cellobiohydrolase EC 3.2.1.91) and cellobiose (βglucosidase EC 3.2.1.21). Glucohydrolase (1.4-β-D-glucan 4- glucohydrolase E.C 3.2.1.71) is sometimes present as part of the minor components. The complete cellulase system acts synergistically to convert crystalline cellulose to glucose (Sukumaran et al., 2005).

2.2.2.1

Mechanism of action for cellulases

All cellulases are glycoside hydrolase (GH) enzymes that utilize the acid-base catalysis mechanism of hydolysis, with inversion or retention of the glucose anomeric configuration. There are two common types of cellulase active sites. GHs with open (groove, cleft) active sites that typically exhibit endocellulolysis (endocellulases), binding anywhere along the length of the cellulose molecule and hydrolysing the β-1,4 glycosidic linkage. The other types are those with tunnel like active sites exhibiting exocellulolytic activity (cellobiohydrolses) binding ends of the cellulose molecule and producing short-chain oligosaccharides (Gray et al., 2006; Prasad et al., 2007; Balat et al., 2008; Sukharnikov et al., 2011). Exocellulases are processive enzymes, i.e. they are attached to a cellulose chain until it is completely hydrolysed. Endocellulases can be processive or non-processive. Efficiency of the processive enzymes contributes to the rate limiting step of cellulose hydrolysis (Sukharnikov et al., 2011). Endocellulases and exocellulases work co-operatively and 17

sometimes synergistically to hydrolyse the crystalline cellulosic substrate (Balat et al., 2008). It has been further discovered that more than one type of each of the cellulases are responsible for the attack of the cellulose crystalline structure (Everleigh, 1987; Dutta et al., 2008). The synergistic effect of these two enzymes results in soluble oligomers, cellobiose and glucose. Cellulases with endo-mode of action appear to be represented by a larger number of protein folds. This indicates that endocellulases are either more evolutionary diverse or many novel exocellulases are yet to be found. Many cellulases are multidomain proteins and in addition to the catalytic domain, have accessory domains such as carbohydrate binding module (CBMs) connected by a flexible linker. The main role of the CBMs is to help cellulases bind cellulose, although they might also participate in the initial disruption of cellulose fibres. Cellulases preferentially bind to the amorphous regions of the cellulose crystalline fibre. Endocellulases (sometimes along CBMs) help disrupt the cellulose fibres and create accessible ends, whereas cellobiohydrolases continue the degradation by removing di- and oligosaccharides (usually 2-4 residues) from the ends of the disrupted cellulose fibres (Sukharnikov et al., 2011). The multiplicity of the cellulase forms is a result of multiple genes, macro heterogeneity due to complexing of cellulases with proteins, glycoproteins and polysaccharides, synthesis of variants of a single gene product via infidelity of translation, proteolysis and variable glycosylation or interaction with components of the broth (Everleigh, 1987).

2.2.2.2

Production of cellulases

Cellulolytic microbes are largely carbohydrate degraders and are generally unable to use proteins or lipids as energy sources for growth. Notable cellulolytic bacteria include Bacilli, Pseudomonas, Cellulomonas and Cytophaga (Sukumaran et al., 2005). The cellulolytic actinomycetes include Streptomyces and Actinomucor. Filamentous fungi are the major sources of cellulases and hemicellulases. Wild type and mutant strains of Trichoderma sp. (T. viridae, T. reesei, T. longibranhiatum) have been considered as the most productive and powerful cellulases (Balat et al., 2008). T. reesei has received much attention due to the ability of the genetically engineered strains to produce extremely large amounts (110 g/L) of 18

crude cellulase, the relatively high specific activity of the crude cellulase on crystalline cellulose and the ability to genetically modify the strains to tailor the set of enzymes to produce so as to give optimal activity for specific uses (Wilson, 2009). Other fungi such as Penicillium and Aspergillus also have the ability to yield high levels of extracellular cellulases (Sukumaran et al., 2005). The majority of reports on microbial production of cellulases utilize submerged fermentation technologies. In nature however, growth and cellulase utilization in aerobic organisms resemble solid state fermentations than liquid cultures. Nevertheless, better monitoring and handling is still associated with submerged fermentations (Sukumaran et al., 2005). Cellulase production in growth cultures is influenced by various factors and their interactions can affect cellulase productivity. Lactose is a known economically feasible additive that induces cellulase genes in industrial fermentation media. Carbon sources in the majority of the commercial cellulase fermentations are cellulosic biomass including straw, spent hull of cereals and pulses, rice, wheat bran, bagasse, paper industry waste and other lignocellulosic residues. The majority of the processes are batch processes. Solid state fermentations for production of cellulases are gaining interest as a cost effective technology not only for cellulase production but also for the bioconversion of lignocellulosic biomass (Sukumaran et al., 2005).

2.2.2.3

Cellulase system and control of cellulase gene expression

Cellulase systems in microorganisms can be generally regarded as complexed or noncomplexed. Utilization of insoluble cellulose requires the production of extracellular cellulases by the organism. The cellulase system consists of either secreted or cell associated enzymes belonging to different classes categorized based on their mode of action and structural properties. (Bhat and Bhat, 1997; Sukumaran et al., 2005). Non-complexed cellulase systems from fungi or bacteria have components of cellulase system free and are mostly secreted. Typical examples include the cellulase system from T. reesei. The fungus produces two exoglucanases and about eight endoglucanases and seven βglucosidases. Complexed systems (cellulosomes) on the other hand are native to anaerobic bacteria. Cellulosomes are protuberances on the cell wall of the organism which harbour 19

stable enzyme complexes. In Clostridia, the cellulosome consists of a non-catalytic cipA protein which has different catalytic modules responsible for exo- and endoglucanase activities. Individual composition of the cellulosome varies with respect to the organism (Sukumaran et al., 2005; Mathew et al., 2008).

2.2.2.4

Regulation of cellulase synthesis

Cellulases are inducible and the regulation of cellulase production is finely controlled by activation and repression mechanisms. In T. reesei, the production of cellulolytic enzymes is induced only in the presence of the substrate, cellulose, and is repressed when easily utilisable sugars are available. The disaccharide sophorose inducer of cellulolytic enzymes is generated by trans-glycosylation activity of the basally expressed β-glucosidase. Cellobiose, δ-cellobiose-1-5 lactone and other oxidized products of cellulose hydrolysis can also act as inducers of cellulose (Bhat and Bhat, 1997; Sukumaran et al., 2005). Glucose repression of the cellulose system overrides induction and de-repression is believed to occur by an induction mechanism mediated by trans-glycosylation of glucose (Sukumaran et al., 2005; Mathew et al., 2008). The promoter region of cellulases harbours binding sites for the carbon catabolite repressor protein (CRE1) catabolite repressor protein as well as sites for transcriptional activators including activator of cellulase expression protein II (ACEII). Glucose expression of cellulose is mediated through the CRE1 in T. reesei (Sukumaran et al., 2005).

2.2.2.5

Genetic engineering of cellulases

Several approaches have been employed to improve cellulase performance and decrease the amount of enzyme needed to saccharify biomass substrates. The primary target for cellulase engineering has been cellobiohydrolase as it naturally constitutes 60-80% of the natural cellulase systems (Gray et al., 2006). Methods used in the engineering of cellulases include: sited directed mutagenesis, site saturation mutagenesis, error prone PCR and DNA shuffling to generate variants of the organisms of interest. Another approach is to introduce

20

heterologous enzymes into an existing system such as T. reesei so that the overall performance of the system is enhanced (Bhat and Bhat, 1997; Gray et al., 2006).

2.2.2.6

Application of cellulases

Cellulases are employed in the food (Mandels, 1985; Hamer, 1991; Béguin and Aubert, 1994), wine and brewery industry (Galante et al., 1993; Godfrey and West, 1996), animal feed (Chesson, 1987; Graham and Balnave, 1995), textile and laundry (Kumar et al., 1994), pulp and paper industries (Suurnakki et al., 1996) as well as in agriculture (Viikari et al., 1993). In the food industry cellulases are used for partial or complete hydrolysis of cell wall polysaccharides and substituted celluloses. The hydrolysis is applied in improving the soaking efficiency and homogenous absorption of water by cereal, to improve the nutritive quality of fermented food, for ease of rehydration of dried vegetables and soups and for production of oligosaccharides as functional food ingredients and low calorie food substituents (Mandels, 1985; Béguin and Aubert, 1994; Bhat and Bhat, 1997). In the beer making industry, cellulases are used in improvement of skin maceration, quality, stability, filtration and clarification of wines (Galante et al., 1993). In feed biotechnology, cellulases improve the nutritional quality of feeds by partial hydrolysis of the lignocellulosic materials, dehulling of cereal grains, decrease in internal viscosity and producing flexibility in the feed materials (Bhat, 2000). Cellulases have achieved great success in the textile and laundry industry because of their ability to modify cellulosic fibres in a controlled and desired manner so as to improve the quality of fabrics (Kumar et al., 1994). Cellulases have been used for removal of excess dye from denim fabrics thus softening the cotton without damaging the fabric. The softening of the fabric has led to production of high quality environmentally friendly washing powders. Cellulases have also been applied in removal of excess microfibrils on the surface of cotton and non-denim fabrics, a process known as biopolishing. Another application has been to restore the brightness and softness of cotton fabrics leading to high quality fabrics (Kumar et al., 1994; Bhat, 2000). In the pulp and paper industry cellulases are used for modification of coarse mechanical pulp and handsheet strength properties, partial hydrolysis of the carbohydrate molecules, the 21

release of ink from the fibre surfaces and hydrolysis of colloidal materials in paper mill drainage (Suurnakki et al., 1996; Bhat, 2000). Cellulases are also employed in the partial or complete hydrolysis of the pulp fibres thus a better bio-characterisation of pulp fibres (Bhat, 2000). 2.2.2.7

Global cellulase market

Cellulases are currently the third largest industrial enzyme by dollar volume. The demand for cellulases is increasing due to its diverse applications in industry (Wilson, 2009). Several companies are involved in cellulase production for textile, detergent, pulp and paper industries. Globally, the two major players are Genencor and Novozymes. Genencor has launched Accelerase 150, a cellulase complex intended specifically for lignocellulosic biomass. Accelerase 150 is claimed to be cost effective and efficient for the bioethanol industry. Accelerase XY has also been launched and this enhances both xylan and glucan conversion (Singhania et al., 2010).

Novozymes also has a diverse range of cellulase preparations available based on application as Cellulusoft AP and Cellulusoft CR for bioblasting in textile mills, Carezyme and Celluclean for laundry in detergents and Denimax for the stonewash industry (Singhania et al., 2010).. There has been a release in a cellulase preparation for biomass hydrolysis from Novozymes known as Cellic CTec3. This is a highly robust enzyme that optimizes biomass conversion with a reduction of the total cost of ethanol production (www.biotimes.com). Though most enzyme producing companies world-wide are involved in production and marketing of cellulase for diverse applications, very few of them develop cellulase for biomass conversion (Singhania et al., 2010; Sarrouh et al., 2012).

2.2.3

Hemicellulases

Hemicellulases are a diverse group of enzymes that hydrolyse hemicelluloses to give simple sugars, organic acids, solvents, animal feeds, methane and ethanol. Hemicellulases

are

mainly glycoside hydrolases that hydrolyse glycosidic bonds between two or more carbohydrates or between a carbohydrate and non-carbohydrate moiety (Polizeli et al., 2005). 22

The glycosidic hydrolases are key in the degradation of plant biomass and carbon flow in nature. A lot of attention has been given to hemicellulases because of the complex branched chain can result in much value added products (Shallom and Shoham, 2003). The degradation of hemicellulose is carried out by microorganisms that can be found free in nature as part of the digestive flora in the digestive tract of higher animals. A wide range of microorganisms produce different types of hemicellulases in response to the different types of hemicelluloses in their environments. The enzymes usually work synergistically to degrade the polymers into monosaccharides or disaccharides (Shallom and Shoham, 2003). The classification of the hemicellulases largely depends on the substrates they work on, the bonds they break and by the patterns of product formation. Catalytic modules of the hemicellulases are either glycoside hydrolases that hydrolyse glycosidic bonds or carbohydrate esterases which hydrolyse ester linkages of acetate or ferulic acid chains (Shallom and Shoham, 2003). Hemicellulases comprise of a set of multiple enzymes mainly the endoxylanase and xylosidase that degrade mainly xylans, mannans and other heteropolysaccharides. Much attention has been placed on xylanases since xylan is the main carbohydrate found in hemicellulose. Xylanases (EC 3.2.1.8) hydrolyze the β-1,4 bonds in the xylan backbone yielding short xylooligomers. The complete hydrolysis of xylan requires synergistic action of enzymes such as endoxylanase (EC 3.2.1.8), exoxylanase (β-D-1,4xylan xylohydrolase), β-Dxyloxidase along with a variety of de-branching enzymes such as α-L-arabinofuranosidase, αglucoonidases and acetyl esterases (Tseng et al., 2002; Shallom and Shoham, 2003; Maciel et al., 2008). Many of the xylanase producing microorganisms express multiple isoforms that have been ascribed a variety of reasons such as heterogeneity and complexity of the xylan structure, genetic redundancy and post translational modifications (Ghotora et al., 2006). The endo-1,4-β-xylanase produce oligosaccharides that are eventually degraded by the xylan 1,4-β xylosidase to produce xylose units. The endoxylanases appear to have a greater specificity for straight chain substrates while the other xylanases accommodate more frequent side chains or branching (Gírio et al., 2010). There is however a variation in xylanases produced in terms of molecular mass, optimal pH and specificity towards the xylan polymer. Some enzymes will cut randomly between unsubstituted xylose residues whereas other xylanases strongly depend on substituents on the xylose residues neighbouring the attacked 23

residues. In addition, hemicellulose degradation needs accessory enzymes such as xylan esterases, ferulic and p-coumaric esterases, α-1-arabinofuranosidases, and α-4-O-methyl glucoronosidases acting synergistically to efficiently hydrolyze wood xylans and mannans (Sanchez, 2009). Xylanases are preferred catalysts for xylan hydrolysis because of their high specificity, mild reaction conditions, negligible substrate loss and side product generation (Kulkarni et al., 1999). The concept of substrate recognition and induction i.e. the control of transcription by the co-operative actions of an activator and a repressor have generally been accepted for xylanases (Tseng et al., 2002). For induction to take place there has to be physical contact between part of the regulatory machinery of the cell and the inducer, which suggests a recognition site on the cell surface. Constitutive xylanases at relatively low levels are supposed to initiate xylan hydrolysis producing small β-D-xylopyranosyl oligosaccharides such as xylobiose and xylotriose among others. The corresponding oligosaccharides are transported into the cell with the help of β-xylosidase permeases where they trigger endoxylanase synthesis (Shallom and Shoham, 2003; Polizeli et al., 2005). The permease activity of the induced cells diminishes in the presence of glucose (Polizeli et al., 2005). This catabolite repression is mediated by the repressor CreA (catabolite repressible entities A). Regulation is mediated by the transcriptional activator XlnR. X1nR regulates the number of genes involved in xylan degradation such as those encoding for β-xylosidase, αglucororonidase, xylanase, arabinoxylan arabinofuranohydrolase and D-xylose reductase (Knob et al., 2010).

2.2.3.1

Sources of xylanases

Xylanases are produced mainly by microorganisms. The hemicellulases have been produced from a number of microorganisms including bacteria, yeasts, and filamentous fungi such as Trichoderma, Bacillus, Cryptococcus, Aspergillus, Penicillium. Aureobasidium, Fusarium, Chaetomium, Phanerochaete, Rhizomucor, Humicola, Talaromyces and many others. These fungi produce enzymes extracellularly with a wide range of activities. Various substrates such as corn stover (Kheng and Omar, 2004; Kumar and Wyman, 2009) and wheat bran, corn cobs, lentil bran and groundnut shells (Kang et al., 2004) have been used in both submerged and solid state fermentation processes for the production of xylanase. 24

Naturally, xylanases take part in the breakdown of plant cell walls along with other enzymes. Plant xylanases digest xylan during the germination of some seeds (e.g. the malting of barley). Xylanolytic enzymes can also be found in marine algae, protozoans, crustaceans, insects, snails and seeds of land plants. Among the microbial sources, filamentous fungi are especially interesting because they secrete these enzymes into the medium and their xylanase levels are much higher than those found in yeasts and bacteria. In bacteria, not only are xylanases produced at lower activity levels than in fungi, but they are also restricted to the intracellular or periplasmic fractions (Knob et al., 2010). Furthermore, enzymes expressed in bacteria are not subjected to post-translational modifications such as glycosylation. Glycosylation affects stability of protein conformation, protects proteins from proteolysis and improves protein solubility (Polizeli et al., 2005).

2.2.3.2

Biochemical properties of xylanases

Fungi and bacteria with the ability to degrade xylans produce a multiplicity of enzymes, some belonging to the same functional class and polymer specificity. Generally filamentous fungi produce a multiplicity of β-xylosidases. Multiplicity of xylanases has been explained by the heterogeneous nature of the xylan polymer. These enzymes exhibit a diversity of physicochemical properties, structures, specific activities and yields (Wong et al., 1988; Wu and Lee, 1997; Yang et al., 2006; Sanchez, 2009). Microbial xylanases are single subunit proteins with molecular masses in the range of 8-145 kDa. The optimum temperature for endoxylanase activity is between 40 and 60 °C (Kulkarni et al., 1999; Knob et al., 2010). Fungal xylanases are less thermostable than bacterial xylanases. Ceratocystis paradoxa, a fungus of mesophilc origin has been found to produce thermostable xylanase able to withstand 80 °C for 1 hour (Dekker and Richards, 1975). A low temperature active enzyme having both carboxymethyl cellulase and xylanase activities from Acremonium alcaliphilum JCM 7366 has been reported. The xylanase and cellulase activities at 0 °C were 25 and 48.4% respectively of their activities at an optimum temperature of 40 °C ((Kazuya et al., 1997; Kulkarni et al., 1999). Xylanases from different organisms are usually stable over a wide pH range (3-10) with an optimum range of 4-7. The isoelectric points of the endoxylanases from various sources range between 3 and 10 (Kulkarni et al., 1999). Most fungal xylanases exhibit isoelectric points in 25

the range of 4-5 though xylanases from Talaromyces emersonni have an unusual pI of 8.9 (Knob et al., 2010). Generally bacteria are known to produce two xylanases; high molecular mass acidic xylanases and low molecular weight basic xylanases. This relationship is however not exhibited in fungi. The amino acid compositions of the xylanases from various sources indicate predominantly aspartic acid, glutamic acid, glycine, serine and threonine (Kulkarni et al., 1999). The occurrence of glycosylated enzymes is a common phenomenon among many eukaryotic xylanases. The carbohydrate groups are covalently linked with protein or are present as dissociable complexes with xylanases. Glycosylation is implicated in promoting stability of the xylanases against extreme environments (Kulkarni et al., 1999; Knob et al., 2010). The carbohydrate content of most xylanases is estimated to be 10-30%. Higher carbohydrate content has been verified from Aspergillus versicolor (47%), Paecilomyces thermophila (61.5%) and Aspergillus phoenicis (43.5%) (Kulkarni et al., 1999).

2.2.3.3

Production of xylanases

The basic factors for efficient production of xylanases are the use of an appropriate inducing substrate and an optimum media composition. The production of cellulase-free xylanase systems is of major importance in the pulp and paper industry. Cellulase free xylanase offers selective removal of xylan from dissolving pulps which ensures minimal damage to the pulp fibres, generate rayon grade or superior quality dissolving pulps, and minimize the use of chemical substances, thus decreasing the amount of waste released during bleaching processes (Coman and Bahrim, 2011). Filamentous fungi are capable of excreting large amounts of xylanases extracellularly. However fungal xylanases are generally associated with cellulases (Kulkarni et al., 1999). Selective production of xylanase is possible in the case of Trichoderma and Aspergillus species using only xylan as a carbon source. On cellulose the strains produce both cellulase and xylanase which may be due to traces of hemicellulose present in cellulosic substrates (Biely, 1993). The mechanisms that govern the formation of extracellular enzymes with reference to carbon sources present in the medium are influenced by the availability of precursors for protein synthesis. Therefore in some fungi, growing cells 26

in xylan not contaminated by cellulose under a lower nitrogen/carbon ratio in the medium may be one of the strategies for producing xylonolytic systems that are free of cellulases (Biely, 1991) . However cellulosic substrates were also found to be essential in medium for maximum xylanase production by Clostridium stercorarium (Berenger et al., 1985), Thermomonospora curvata (Stutzenberger and Bodine, 1992)) and Neurospora crassa (Deshpande et al., 1986; Kulkarni et al., 1999; Gírio et al., 2010). Cheaper hemicellulosic substrates like corn cob, wheat bran, rice bran, rice straw, corn stalk and bagasse have been found to be suitable for production of xylanases in the case of Aspergillus awamori and Penicillium purpurogenum (Dey et al., 1992).

2.2.3.4

Factors affecting xylanase yield

The yield of xylanases in fermentation is governed by a few factors in addition to the standard parameters. When xylanase fermentation is carried out on complex heterogeneous substrates various factors have a combined effect on the level of xylanase expression. These include substrate accessibility, rate and amount of release of the xylo-oligosaccharides and their chemical nature and quantity of xylose released which acts as the carbon source and as an inhibitor of xylanase synthesis in most cases. Generally the slow release of the inducer molecules and the possibility of the culture filtrate converting the inducer to its nonmetabolizable derivative are believed to boost the level of xylanase activity (Kulkarni et al., 1999). Xylanases bind tightly to the substrate; therefore part

of the enzyme produced during

fermentation is often lost or discarded as bound enzyme along with the insoluble substrate. The metabolic enzymes of the xylanase producer such as proteases and transglycosides also affect the yield of xylanases (Chauthaiwale and Rao, 1994). These enzymes are optimally expressed at the end of the exponential phase and thus harvesting of the xylanases must be correlated to the production of these enzymes. Other bioprocess parameters that can affect the activity and productivity of xylanase attained in a fermentation process include pH, temperature and agitation (Kulkarni et al., 1999).

27

2.2.3.5

Xylanases from extremophilic regions

Commercial applications of xylanases demand highly stable enzymes active under routine handling conditions. Advantages such as reduced contamination risk and faster reaction rates have been proposed for the use of thermophiles in biotechnology processes. Thermostability of the enzyme in the presence of the substrate and residual activities of enzymes in harsh processing environments are important (Kulkarni et al., 1999; Knob et al., 2010). Alkaline xylanases have gained importance due to their application for the development of ecofriendly technologies used in the pulp and paper industries. The enzymes are able to hydrolyze xylan in alkaline solutions. Many xylanases produced by alkaliphilic microorganisms such as Bacillus sp. (Okazaki et al., 1999) and Aeromonas sp. with an optimum growth at pH 10.0 show stability at pH 9-10 but have lowered activity above pH 8. Xylanase from Cephalosporium an alkaliphilic fungus has been reported as having a broad pH range of 6.5-9.0 (Chandra and Chandra, 1996). Xylanases from thermophilic bacteria such as Thermonospora fusca, thermophilic Bacillus sp. and Bacillus stearothermophilus show an optimum temperature in the range of 65-80 °C (Khasin et al., 1993). The thermostable xylanases produced by a thermotolerant Aspergillus strain at 37 °C showed maximum activity at 80 °C. Xylanase from Clostridium stercorarium has a temperature optimum of 70 °C and a half -life of 90 minutes at 80 °C. Thermogata sp. produced a xylanse with a temperature optimum of 105 °C at pH 5.5 with a half-life of 90 minutes at 95 °C (Simpson et al., 1991). Among the fungal species, Thermoascus aurantiacus xylanase has been reported to be stable at 70 °C for 24 hours with a half-life of 88 minutes at 80 °C (Yu et al., 1987). Paecilomyces variota and T. byssochlamydoides show an optimum temperature of 65-75 °C at pH 5-6.5 (Kulkarni et al., 1999).

2.2.3.6

Regulation of xylanase synthesis

Xylanase production by various bacteria and fungi has been shown to be inducible (Kulkarni et al., 1999). Rare examples of constitutive xylanases have been reported. Regulation of xylan synthesis is a complex phenomenon and the response to an individual inducer varies with organisms. An inducer producing maximum xylanase activity in one species may be an 28

inhibitor of activity in another species. The substrate derivatives and the enzymatic end products may often play a key positive role in the induction of xylanases; they can also act as end product inhibitors, possibly at much higher concentrations (Kulkarni et al., 1999; Knob et al., 2010). Xylan being a high molecular mass polymer cannot penetrate the cell wall. The low molecular mass fragments of xylan play a key role in the regulation of xylanase biosynthesis. The low molecular weight fragments include xylose, xylobiose, xylooligosaccharides, heterodisaccharides of xylose and glucose and their positional isomers. These molecules are liberated from xylan by the action of small amounts of constitutively produced enzyme. Cellulose has also been shown to be an inducer of xylanase in a few cases though it is still not clear whether the inducing effect lies with cellulose or the contaminating xylan fraction (Kulkarni et al., 1999). Xylose produced higher xylanase yields when used as an inducer in Bacillus pumilus, (Paul and Varma, 1990) and Streptomyces lividans (Kluepfel et al., 1990). However, in Cryptococcus albidus xylose repressed xylanase production (Biely and Petrakova, 1984). Xylobiose was found to be a specific inducer of xylanase in T. reesei (Hrmova and Vrsanska, 1986). D-xylan fragments as well as methy-β-D-xylopyranoside induce xylanse in Streptomyces sp. and in yeast of the genera Cryptococcus and Trichosporon (Kulkarni et al., 1999; Knob et al., 2010). At molecular level, regulation of xylanase synthesis is through catabolite repression. Catabolite repression by glucose is a common phenomenon. Relatively few reports on the relation of cAMP to xylanase induction are available. In the case of C. albidus, when xylan or methyl-β-xylopyranoside were used as inducers, cAMP caused two-fold increase in xylanase production (Morosoli et al., 1989). However cAMP had no effect on the repression caused by D-xylose. It has been suggested that a 15 base pair nucleotide sequence upstream from the βxylanase gene may be part of the cAMP regulatory sequence. In Aspergillus tubigensis, a 158 base pair region upstream of the xylanase gene was shown to be involved in xylan specific induction (De Graai et al., 1994). Catabolite repression of the xylanase gene was found to be controlled directly through repression of gene transcription and indirectly by repression of the transcriptional activator (Kulkarni et al., 1999). Deletion and functional analysis of xylanase genes from A. niger and A. tubigensis led to the discovery of a triplicated sequence that

appears to control enzyme induction. Enzyme 29

synthesis was also found to be repressed when easily metabolizable carbon sources were present in the growth medium suggesting that the synthesis of the enzyme is controlled by transitional state regulators and catabolite repression (Knob et al., 2010). In order to be competitive with lignocellulose derived ethanol, the enzymes used for saccharification processes must be more efficient, produced at high yields and far less expensive (Gray et al., 2006). Cost reduction has been achieved by a combination of enzyme engineering and fermentation process development (Wong et al., 1988). Genetic modifications have been performed to meet the industrial requirements of activity, enantioselectivity and sterospecificity. Stability, tolerance to toxic reagents and extreme conditions (Wu and Lee, 1997). Random and site directed mutagenesis have been applied for strain improvement. Mutagenesis has been applied successfully to hyperproducing xylanase organisms such as Aspergillus usamii, Penicillium oxalicum and Thermomyces lanuginosus and Pseudomonas sp (Gírio et al., 2010).

2.2.3.7

Application of xylanases

Xylanases are employed in bread-making together with α-amylase, malting amylase, glucose oxidase and proteases. Xylanases break down the hemicelluloses in wheat flour, helping in the re-distribution of water, leaving the dough softer and easier. During the baking process, the effect of the xylanases is to delay crumb formation thus allowing the dough to grow thus an increase of bread volume, improvement of bread texture and extension of shelf life (Comacho and Aguilar, 2003; Polizeli et al., 2005). In biscuit making, xylanase is recommended for making cream crackers lighter and improving the texture, palatability and uniformity of the wafers (Beg et al., 2001; Polizeli et al., 2005). The production of fruit and vegetable juices requires that the juices be extracted, cleared and stabilized. Xylanases used in conjunction with cellulases, amylases and pectinases lead to improved yield of juice has been attained by means of liquefaction of the juices, stabilization of the fruit pulp, increased recovery of aromas, essential oils, vitamins, mineral salts, edible dyes and pigments, reduction of viscosity, hydrolysis of substances that hinder the physical or chemical clearing of the juice or that may cause cloudiness in the concentrate (Wong et al., 1988; Polizeli et al., 2005). Xylanases are applied in the preparation of dextrans for use as 30

food thickeners. Furthermore, xylanase in combination with endoglucanase takes part in the hydrolysis of arabinoxylan and starch, separating and isolating the gluten from the starch in the wheat flour (Polizeli et al., 2005). Combinations of cellulases and xylanases are used in partial hydrolysis of animal feeds to improve the quality of silage and green feed (Muthezhilan et al., 2007). The anti-nutrient factors in the feed raise the viscosity of the ingested feed interfering with the mobility and absorption of other components. Xylanases are used in animal feed along with glucanases, pectinases, cellulases, proteases, amylases, phytase, galactosidase and lipases (Twomey et al., 2003). Xylanases break down arabinoxylans in the ingredients of the feed, reducing viscosity of the raw material. The arabinoxylan found in the cell walls of grains has an anti-nutrient effect on poultry, fowl and swine (Wong et al., 1988; Wong and Saddler, 1993; Beg et al., 2001) which produce xylanases in smaller quantities than adults, so food supplements containing these exogenous enzymes improve their energy utilization (Wong et al., 1988; Polizeli et al., 2005). Cellulase free xylanolytic complex is used in the textile industry to process plant fibres such as hessian or linen. This step eliminates the need for use of strong bleach in the production of textiles (Coman and Bahrim, 2011). Xylanases from different families have been proved to be a serious candidate in substitution of conventional chemical soap treatment for wool and speciality hair fibres (Paës et al., 2012). In the pulp and paper industry, xylanases are also used for bleaching the cellulose pulp. Xylanases hydrolyse the xylan precipitated on the lignin due to lowering of the pH at the end of the cooking process (Beg et al., 2001). Xylanases may also be used to prepare materials for scientific research. Well characterised xylanases may be useful for the characterisation of polysaccharides and plant cell walls (Paës et al., 2012). Selected xylanases may be suitable for hydrolysing branched and unbranched, short and long or labelled xylooligosaccharides, which are model compounds used for studying mechanisms of xylanase action. Some xylanases may be used to improve cell wall maceration for production of plant protoplasts (Wong et al., 1988; Paës et al., 2012).

31

2.3

Processing of lignocellulosics to bioethanol

Lignins are often bound to adjacent cellulose fibres to form a lignocellulosic complex (Balat et al., 2008). The lignocellulosic complex is highly resistant to chemical and biological degradation (Martínez-Herrera et al., 2006). The factors to be considered in the use of lignocellulosics for ethanol production in comparison to starch based ethanol production include the efficient depolymerisation of cellulose and hemicelluloses to soluble sugars, efficient fermentation of a mixed sugar hydrolysate (hexoses and pentoses) as well as inhibitory compounds from fermentation, advanced process integration to minimize energy demand and lowering the lignin content of the feedstock thus decreasing the cost of bioethanol (Balat et al., 2008). Bioconversion of lignocellulosics to sugars has the advantage of creating a biorefinery and producing value added products plus fuel bioethanol. The processing of ligocellulosics consists of four major unit operations which are pre-treatment, hydrolysis, fermentation and product separation/distillation and effluent treatment (Cardona and Sánchez, 2007; Balat et al., 2008).

2.3.1

Pre-treatment of lignocellulosics

The initial step in the use of lignocellulosic biomass is size reduction and pre-treatment. Raw untreated biomass is recalcitrant to enzymatic digestion. There seems to be a direct correlation between the removal of the lignin and the hemicelluloses and the digestibility of the cellulose (Gray et al., 2006). Pre-treatment alters or removes the structural and compositional impediments to hydrolysis, ensuring the disruption of the plant cell wall, increase in pore size and reduction of crystallinity and improves enzymatic access to the polysaccharide (Gray et al., 2006; Hahn-Hägerdal et al., 2006; Balat et al., 2008). An effective pre-treatment process must be able to improve the hydrolysis of sugars or the ability to subsequently form sugars by hydrolysis, avoid degradation or loss of carbohydrate, avoid formation of by-products inhibitory to subsequent hydrolysis and fermentation process and be cost effective (Balat et al., 2008). Pre-treatment defines the cost and extent to which the carbohydrate can be converted to ethanol. Currently pre-treatment is the major challenge of cellulose bioethanol technology research and development (Balat et al., 2008).

32

Pre-treatment can be carried out in different ways such as mechanical pre-treatment, steam explosion, ammonia fibre explosion, supercritical carbon dioxide treatment, alkali or acid pre-treatment, ozone and biological treatments (Balat et al., 2008).

2.3.1.1

Mechanical pre-treatment

Lignocellulosics can be comminuted by a combination of chipping, grinding and milling to reduce the cellulose crystallinity. Vibratory ball milling has been found to be more effective in breaking down the cellulose crystallinity of aspen and spruce chips and improving digestibility of the biomass than ordinary ball milling (Prasad et al., 2007).

2.3.1.2

Pyrolysis

This is a process in which materials are treated at temperatures greater than 300 °C. Cellulose rapidly decomposes at this temperature to produce gaseous products and residual char. Decomposition is much slower and less volatile products are formed at lower temperatures (Prasad et al., 2007). Analysis of the hydrolysates from this pre-treatment method shows the presence of xylan oligomers and polymers with large chains (Cardona et al., 2010).

2.3.1.3

Steam explosion (autohydrolysis)

In this method, chipped biomass is treated with high-pressure saturated steam. The pressure is swiftly reduced making the material to undergo an explosive decomposition. The process causes hemicellulosic degradation and lignin transformation due to high temperature. Addition of sulphur dioxide or carbon dioxide in steam explosion improves the hydrolysis, decreases the production of inhibitory compounds and leads to more complete removal of hemicelluloses (Prasad et al., 2007; Balat, 2011). Steam explosion uses 70 % less energy as compared to mechanical communition to achieve the same size reduction. Limitations of steam explosion include disruption of the lignin-carbohydrate matrix and generation of compounds that are inhibitory to microorganisms in downstream processes. To remove the

33

inhibitory compounds, washing with water is necessary. Washing will however remove soluble hemicelluloses (Prasad et al., 2007). A relatively high hemicellulose recovery in the range of 55-84% together with low levels of inhibitory products by-products has been obtained through steam explosion. Cellulose and lignin are not significantly affected, yielding a cellulose and lignin rich solid phase together with a liquid that has a relatively low concentration of potential fermentation inhibitors. Corrosion problems are reduced owing to the mild pH conditions. Capital and operational costs are reduced with beneficial consequences on the environment compared to other hydrolytic technologies (Gírio et al., 2010).

2.3.1.4

Ammonia fibre/freeze explosion (AFEX)

This is another type of physico-chemical pre-treatment in which lignocellulosic materials are exposed to liquid ammonia at high temperature and pressure for a period of time after which the pressure is swiftly reduced. AFEX significantly improves the saccharification rates of various herbaceous crops and grasses. The AFEX treatment is however not very effective for biomass with high lignin contents. AFEX has the advantage of not producing inhibitors for the downstream biological processing and does not require small particle size for efficacy. (Prasad et al., 2007; Gírio et al., 2010; Balat, 2011).

2.3.1.5

Ozone treatment

Ozone can be used to degrade lignin and hemicelluloses in many lignocellulosic materials such as wheat straw, bagasse, green hay, peanut pine, cotton straw and poplar sawdust. The degradation is limited to lignin though hemicellulose is slightly affected. Ozolysis has no effect on cellulose. The ozolytic reaction effectively removes lignin and does not produce toxic residues for the downstream processing. Reactions are carried out at room temperature and atmospheric pressure thus making the process easy to carry out. The disadvantage with this process is that large amount of ozone is required making the treatment to be expensive (Sun and Cheng, 2002).

34

2.3.1.6

Organosolv

Organoslov is a treatment process which employs the direct action of water dissolved organic solvents such as ethanol, methanol and acetone. The dissolved organic solvents usually act in combination with an acid and solubilize the lignin whilst hydrolysing the hemicellulose fraction (Chen and Qiu, 2010; Gírio et al., 2010). The process temperatures vary from room temperature up to 295 °C depending mainly on the solvent used. Among the most effective processes are the water:ethanol blends catalysed by sulphuric acid with an operational temperature between 180 and 200 °C. Ethanol has the advantage of being recovered by distillation. The process has high cellulose digestibility. The disadvantage of using organoslov is that there is a significant amount of furfural produced implying low hemicellulose recovery. Peracid based processes are more specific for lignin but are more expensive and the technology is demanding for safety reasons (Gírio et al., 2010).

2.3.1.7

Concentrated acid hydrolysis

Concentrated sulphuric and hydrochloric acids are used in the treatment of lignocellulosic materials and the treatment provides rapid and complete conversion of cellulose to glucose and hemicelluloses to 5-carbon sugars with little degradation (Balat et al., 2008). Relatively mild temperature is used for this process and the only pressure involved is that of pumping materials from vessel to vessel. Although the acids are powerful oxidizing agents they are toxic, corrosive, and hazardous and require reactors that are highly resistant to corrosion. It is critical to optimize sugar recovery and to recover the acid for recycling in order to keep the process economically viable (Dermibas, 2007; Prasad et al., 2007). Acidic pre-treatments may result in high concentrations of furfurals in the liquid phase (Gray et al., 2006). Even at low concentrations (3-15 mM), furfural can severely affect the rate of ethanol production and final conversion, thus limiting the ethanol production processes. The advantage of this process is the quick rapid conversion of cellulose to glucose and hemicelluloses and to 5carbon sugars with little degradation (Prasad et al., 2007).

35

2.3.1.8

Dilute acid hydrolysis

Dilute acid hydrolysis has been successfully developed for treatment of lignocellulosic materials. Dilute sulphuric acid pre-treatment can achieve high reaction rates and significantly improve cellulose hydrolysis. The dilute acid mainly attacks the hemicelluloses, leaving the cellulose and lignin fractions almost unaltered (Cardona et al., 2010). The mechanism of hydrolysis includes the diffusion of protons through the wet lignocellulosic materials, protonation of the oxygen of a heterocyclic ether bond between the sugar monomers, breaking the ether bond, generation of a carbocation as an intermediate, salvation of the carbocation with water, regeneration of the proton cogeneration of the sugar monomer, oligomer or polymer depending on the position of the ether bond and diffusion of the reaction products in the liquid phase (Aguilar et al., 2002). There are two types of dilute acid treatment processes, the high temperature (>160 °C), continuous flow process for low solid loading (5-10%) and low temperature (