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Production of Biobutanol from Inulin-Rich Biomass and Industrial

Production of Biobutanol from Inulin-Rich Biomass and Industrial

Production of Biobutanol from inulin-rich biomass and industrial

food processing wastes

Thesis

Presented in Partial Fulfillment of the Requirements for the Degree Master of Science in the Graduate School of The Ohio State University

By

Ashok Kumar Bharathidasan

Graduate Program in Food, Agricultural and Biological Engineering

The Ohio State University

2013

Master's Examination Committee:

Dr. Katrina Cornish, Advisor

Dr. Thaddeus C. Ezeji

Dr. Frederick C. Michel

Copyrighted by

Ashok Kumar Bharathidasan

2013

ABSTRACT

Inflation of crude oil prices, diminishing oil resources and increasing environmental concerns have accelerated the search for renewable alternatives for gasoline. In recent years, biobutanol has gained enormous attention as a potential gasoline substitute due to its high energy density, low vapor pressure, low heat of vaporization and high hydrophobicity. These physical and chemical properties make butanol suitable for blending with or direct substitution of gasoline. Biobutanol can be produced through acetone-butanol- (ABE) from diverse feedstocks.

Butanol could occupy a significant portion of advanced markets if the economics of ABE fermentation process improve. Although, butanol toxicity, low yield, and high butanol recovery costs are some of the challenges of ABE fermentation, high substrate cost still makes up least 50% of the total production cost. The objectives of this study were to utilize locally available waste biomass for butanol production using selected strains of Clostridia.

Different food processing wastes were obtained from major food processing industries throughout Ohio and screened for their suitability for ABE fermentation.

Among 48 different sample wastes, four substrates, namely, milk dust powder, breading, inedible dough and batter liquid were selected for direct-utilization of these substrates for butanol production. The ability of C. beijerinckii NCIMB 8052 and C. acetobutylicum

ATCC 824 to ferment food processing wastes was tested in batch-fermentation mode. C. ii

acetobutylicum ATCC 824 gave the highest ABE yields on the media with milk dust powder (10.25 g/L), inedible dough (16.30 g/L) and batter liquid (17.41 g/L).

C.beijerinckii NCIMB 8052 gave the highest ABE yields on the breading fermentation medium (14.80 g/L).

Besides food processing wastes, inulin extract was tested for its potential to produce butanol. This is a co-product obtained during rubber extraction from alternate rubber producing crop, Taraxacum Kok-saghyz, also known as TKS (Kazak dandelion,

Russian dandelion or Buckeye Gold). Four different strains, namely, C. beijerinckii

NCIMB 8052, C. acetobutylicum ATCC 824, C. saccharobutylicum P262, and C. beijerinckii NRRL B592 were investigated for their ability to use raw (unhydrolyzed) and enzymatically-hydrolyzed inulin medium. Chicory inulin, which has similar molecular characteristics to TKS inulin, also was tested. C. saccharobutylicum P262 fermented the raw inulin media best (TKS, 8.48 g/L ABE; chicory, 12.50 g/L ABE), whereas C. beijerinckii NCIMB 8052 did best in the enzymatically-hydrolyzed inulin medium.

From, C. beijerinckii NCIMB 8052 gave maximum ABE of 10.00 g/L and 12.60 g/L from the enzymatically-hydrolyzed TKS and chicory inulin media, respectively.

Fermentation of food processing wastes and/or inulin derived from TKS could be scaled up into an industrial fermentation process that would improve economics, help meet local energy demands, provide easy value-added disposal of these wastes, and provide needed by other industries. In addition, a valuable co-product from TKS will help commercialization of TKS as a viable natural rubber producing crop in USA.

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Dedicated to my family and my friends

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ACKNOWLEDGMENTS

I record my profound gratitude and indebtedness to my advisor Prof. Katrina

Cornish for her unfailing support, inspiration and intellectual stimulation during the period of my research. Working with her taught me the value of hard work, dedication perseverance and time management not only in research projects but also in personal life.

I also owe special thanks to my advisor for her patience, encouragement and help during my most difficult times.

This thesis would not have been possible without valuable inputs and timely guidance from my co-advisor Dr. Thaddeus Ezeji who provided all the necessary lab facilities for successful completion of my research work. I would also like to express my deep appreciation to my committee member Dr. Frederick C. Michel and his research team for their generous support and immense help to accomplish my HPLC analysis.

Without them, it would have taken much long to produce these results. Special thanks are due to all my committee members for their scholarly suggestions, constructive criticism, and prompt help in reviewing this document.

Thanks are extended to all my lab members in Williams and Gerlaugh Hall for their timely help and support, which kept me moving forward during my stay in OARDC,

Wooster, OH. Dr. Sukhbir Grewal, Mrs. Candy McBride, and Mrs. Peggy Christman, deserve special mention for their administrative support.

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I express my heartfelt thanks to my beloved parents, Aarthi and my friends whose love, blessings and encouragement have always strengthened my motivation and boosted my morale. Last but not least, I want to thank the Department of Food, Agricultural and

Biological Engineering of The Ohio State University for providing me an opportunity to pursue my Master’s in the United States. The intellectual ambiance and the time I spent with The Ohio State University will be etched in my memory forever.

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VITA

October 30, 1986 ...... Born, Arantangi, India

2007 ...... B. Tech., Agricultural Engineering, Tamil

Nadu Agricultural University, India

2007-2009 ...... Junior Research Fellow, ICAR, India

2010 to present ...... Graduate Research Assistant, Department of

Food, Agricultural and Biological

Engineering, The Ohio State University

FIELDS OF STUDY

Major Field: Food, Agricultural, and Biological Engineering

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TABLE OF CONTENTS

Abstract ...... ii

Acknowledgments ...... v

Vita……...... vii

Fields of Study ...... vii

Table of Contents ...... viii

List of Tables...... xiv

List of Figures ...... xvii

Chapter 1: Introduction ...... 1

1.1 Background ...... 1

1.2 Rationale and Significance ...... 2

1.3 Current butanol production scenario ...... 6

1.4 Limitations of butanol fermentation ...... 8

1.5 Research objectives ...... 8

Chapter 2: Literature Review ...... 13

2.1 ...... 13

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2.1.1 Current scenario of biofuels ...... 14

2.1.2 Classification of biofuels...... 15

2.1.3 Liquid biofuels ...... 17

2.2 Production of Butanol ...... 20

2.2.1 Chemical Synthesis of Butanol ...... 20

2.2.2 Acetone-Butanol-Ethanol (ABE) fermentation ...... 22

2.2.3 General description of species and biobutanol production ...... 25

2.2.4 Fermentative pathways of Clostridia ...... 26

2.2.5 Strain development ...... 27

2.3 Alternate feedstocks ...... 30

2.3.1 ...... 30

2.3.2 utilization ...... 35

2.3.3 Microalgae ...... 36

2.3.4 Food processing wastes ...... 37

2.3.5 Inulin as substrate ...... 42

2.4 Classification of fructans ...... 43

2.4.2 Role of fructans in plants ...... 49

2.4.3 Normal occurrence of fructan ...... 49

2.5 Inulin ...... 50

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2.5.1 Nomenclature ...... 51

2.5.2 Physicochemical properties of Inulin ...... 52

2.5.3 Applications and uses of inulin ...... 52

2.5.4 Inulin Hydrolysis ...... 54

2.6 Production of exo- and endo-inulinases ...... 60

2.6.1 Optimum pH and temperature for inulinase production ...... 62

2.6.2 Factors affecting inulinase production ...... 63

2.6.3 Substrates for inulinase production ...... 64

2.7 Inulin characterization and estimation ...... 65

Chapter 3: Biobutanol production from industrial food processing wastes...... 92

3.1 Introduction ...... 92

3.2 Materials and Methods ...... 97

3.2.1. Determination of total solids and moisture ...... 97

3.2.2. Determination of ash content ...... 98

3.2.3. Estimation of Total Organic (TOC) ...... 99

3.2.4. Estimation of Total Nitrogen (TN) ...... 99

3.2.5. Determination of major, minor and trace elements ...... 100

3.2.5. Measurement of pH ...... 101

3.2.6. Determination of energy content ...... 102

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3.2.7 Selection of an ideal substrate for butanol production ...... 102

3.2.8 Selection of microorganism, culture maintenance and inoculum preparation 106

3.2.9 Medium preparation and ABE fermentation ...... 107

3.2.10 Analytical methods ...... 109

3.3 Carbohydrate metabolism ...... 111

3.3.1 Lactose metabolism in solventogenic Clostridium species ...... 111

3.3.2 Starch metabolism in solventogenic Clostridium species ...... 112

3.4 Results and discussion ...... 113

3.4.1 ABE production from milk dust powder...... 113

3.4.2 ABE production from starchy food processing wastes ...... 118

3.4.3 Feasibility of using other starchy wastes ...... 125

3.5 Conclusions ...... 125

3.5.1 Milk dust powder ...... 125

3.5.2 Starchy food processing wastes ...... 126

Chapter 4: Biobutanol production from inulin-rich biomass ...... 166

4.1 Introduction ...... 166

4.2 Materials and Methods ...... 170

4.2.1 Microorganisms and culture maintenance ...... 170

4.2.2 Inulin extracts from different sources ...... 171

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4.2.3 Media preparation ...... 173

4.2.4 Calcium carbonate as a component of the fermentation medium...... 174

4.2.5 Enzymatic hydrolysis of Inulin ...... 176

4.2.6 Analytical methods ...... 178

4.2.7 Production of endo-and exo-inulinase enzymes ...... 181

4.3 Results and discussion ...... 184

4.3.2 Production of exo- and endo-inulinases ...... 188

4.3.3 Butanol production from unhydrolyzed inulin media...... 191

4.3.4 Sugar consumption pattern of C. saccharobutylicum P262 from unhydrolyzed

inulin extract media ...... 196

4.3.6 Applications of residual biomass of TKS and guayule ...... 205

4.4 Conclusion ...... 207

References ...... 237

Appendix A: Description of food processing wastes and modes of utilization by food

processing industries ...... 283

Appendix B: Calculations on amount of substrate required to make 50 g/L starch medium

...... 292

Appendix C: Cell growth (colony forming unit per ml) of Clostdria in food processing

wastes fermentation media ...... 294

Appendix D: HPLC chromatograms of unhydrolyzed inulin medium before fermentation

...... 296 xii

Appendix E: HPLC chromatograms of unhydrolyzed inulin medium after fermentation

...... 298

Appendix F: HPLC chromatograms of 1 h enzymatically-hydrolyzed pure inulin medium

before and after fermentation ...... 300

Appendix G: HPLC chromatograms of 48 h enzymatically-hydrolyzed pure inulin

medium before and after fermentation ...... 302

Appendix H HPLC chromatograms of enzymatically hydrolyzed inulin extracts...... 304

Appendix I HPLC Chromatogram calibration standards ...... 306

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LIST OF TABLES

Table 2.1 Physical and chemical properties of n-butanol compared with other ...... 71

Table 2.2 Comparison of properties of n-butanol with its ...... 72

Table 2.3 Major applications of butanol isomers ...... 73

Table 2.4 Composition of guayule biomass ...... 74

Table 2.5 Elemental composition of guayule biomass ...... 74

Table 2.6 Types of food processing wastes and its modes of utilization ...... 75

Table 2.7 General nomenclature used in Inulin studies ...... 77

Table 2.8 Physicochemical properties of inulin ...... 78

Table 2.9 Inulinase producing microorganisms and their maximum yield ...... 79

Table 3.1 . Amount of starch present in selected food wastes ...... 128

Table 3.2 .Amount of sugars present in dairy wastes ...... 128

Table 3.3 Performance and kinetic parameters of ABE production from starchy-food

processing wastes using C. beijerinckii NCIMB 8052 after 72 h fermentation ..129

Table 3.4 Performance and kinetic parameters of ABE production from starchy-food

processing wastes using C. acetobutylicum ATCC 824 after 72 h fermentation

...... 130

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Table 3.5 Performance and kinetic parameters of ABE production from milk dust powder

after 72 h fermentation ...... 131

Table 3.6 Different types of wastes and their possible applications ...... 132

Table 4.1 Amount of sugars present in raw inulin extract from different sources ...... 208

Table 4.2 Energy density of residual TKS and guayule biomass...... 208

Table 4.3 Performance and kinetic parameters of ABE production from unhydrolyzed

inulin media by C. saccharobutylicum P262 ...... 209

Table 4.4 Performance and kinetic parameters of ABE production from 48 h

enzymatically-hydrolyzed inulin extract media after 72 h fermentation by C.

beijerinckii NCIMB 8052 ...... 210

Table 4.5 Amount of sugars present in 1 h enzymatically-hydrolyzed pure inulin medium

before and after fermentation by Clostridium beijerinckii NCIMB 8052 ...... 211

Table 4.6 Amount of sugars present in 1 h enzymatically-hydrolyzed pure inulin medium

before and after fermentation by Clostridium acetobutylicum ATCC 824 ...... 211

Table 4.7 Amount of sugars present in 48 h enzymatically-hydrolyzed pure inulin

medium before and after fermentation by Clostridium beijerinckii NCIMB 8052

...... 212

Table 4.8 Amount of sugars present in 48 h enzymatically-hydrolyzed pure inulin

medium before and after fermentation by Clostridium acetobutylicum ATCC 824

...... 212

Table A1 Types of food processing wastes and modes of utilization by food processing

industries ...... 284

Table A2 Description of food processing wastes ...... 288 xv

Table B1 Calculations on amount of substrate required to make 50 g/L starch medium.

...... 293

Table C1Growth of C. beijerinckii NCIMB 8052 (CFU/ml) in food processing wastes

media ...... 295

Table C2 Growth of C. acetobutylicum ATCC 824 (CFU/ml) in food processing wastes

media ...... 295

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LIST OF FIGURES

Figure 2.1 World’s total energy supply and consumption in 2009 ...... 83

Figure 2.2 Major classifications of biofuels ...... 84

Figure 2.3 U.S. Primary energy consumption by energy sources, 2011 ...... 85

Figure 2.4 U.S. Primary energy consumption by energy sources, 2011 ...... 85

Figure 2.5 Acidogenic and solventogenic phase of Clostridia ...... 86

Figure 2.6 Metabolic pathway of Clostridium acetobutylicum ...... 87

Figure 2.7 Pretreatment of lignocellulosic structure ...... 88

Figure 2.8 GF2 Fructan ...... 89

Figure 2.9 F3 Fructan ...... 89

Figure 2.10 Structures of fructo-oligosaccharides ...... 90

Figure 2.11 Enzymes involved in fructan biosynthesis ...... 91

Figure 3.1 Percent total solids, ash and pH (a) vegetable wastes, (b) fat-rich industrial

wastes, (c) industrial sludge wastes, (d) starchy and other industrial wastes ...... 142

Figure 3.2 Mean calorific value, percent carbon, nitrogen and C/N ratio (a) vegetable

wastes, (b) fat-rich industrial wastes, (c) industrial sludge wastes, (d) starchy and

other industrial wastes ...... 146

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Figure 3.3 Concentration of major elements (a) vegetable wastes, (b) fat-rich industrial

wastes, (c) industrial sludge wastes, (d) starchy and other industrial wastes ...... 150

Figure 3.4 Concentration of minor elements (a) vegetable wastes, (b) fat-rich industrial

wastes, (c) industrial sludge wastes, (d) starchy and other industrial wastes ...... 154

Figure 3.5 Fermentation of food processing wastes using C. beijerinckii NCIMB 8052 (a)

butanol production, (b) total ABE production ...... 158

Figure 3.6 Fermentation of food processing wastes using C. beijerinckii NCIMB 8052 (a)

Acetone production, (b) Change in pH ...... 159

Figure 3.7 Fermentation of food processing wastes using C. beijerinckii NCIMB 8052 (a)

acetic acid production, (b) butyric acid production ...... 160

Figure 3.8 Fermentation of food processing wastes using C. acetobutylicum ATCC 824

(a) butanol production, (b) total ABE production...... 161

Figure 3.9 Fermentation of food processing wastes using C. acetobutylicum ATCC 824

(a) acetone production, (b) change in pH ...... 162

Figure 3.10 Fermentation of food processing wastes using C. acetobutylicum ATCC 824

(a) acetic acid production, (b) butyric acid production ...... 163

Figure 3.11 Milk dust powder consumption by C. beijerinckii NCIMB 8052 and C.

acetobutylicum ATCC 824 ...... 164

Figure 3.12 Cell growth of C. beijerinckii NCIMB 8052 in food processing waste media

...... 164

Figure 3.13 Cell growth of C. acetobutylicum ATCC 824 in food processing waste media

...... 165

Figure 3.14 Structural changes of milk dust medium ...... 165 xviii

Figure 4.1 Flow schemes of TKS processing and rubber extraction…………………...213

Figure 4.2 Overview of inulin fermentation ...... 214

Figure 4.3 Appearance of chicory and TKS extract… ...... 215

Figure 4.4 Effect of pH on endoinulinase, Novozyme 960 ...... 216

Figure 4.5 Overview of inulinase production ...... 217

Figure 4.6 Kestose hydrolysis pattern of C. beijerinckii NCIMB 8052 ...... ……….218

Figure 4.7 Inulin hydrolysis pattern at different enzyme concentrations using endo-

inulinase (Novozyme 960) (a) TKS Eskew inulin, (b) Chicory inulin, (c)

Commercial standard inulin ...... 219

Figure 4.8 Inulin hydrolysis and sugar production pattern at maximum enzyme

concentration (150 IU/g of inulin) from different inulin source ...... 221

Figure 4.9 Change in pH of (a) TKS and chicory inulin extract, (b) standard commercial

inulin during hydrolysis ...... 222

Figure 4.10 Effect of substrate concentration on biomass concentration of (a)

Kluyveromyces marxianus ATCC 16045, (b) Kluyveromyces marxianus ATCC

52466 ...... 223

Figure 4.11 Effect of substrate concentration on production of (a) endoinulinase from

Kluyveromyces marxianus ATCC 16045, (b) exo-inulinase from Kluyveromyces

marxianus ATCC 52466 ...... 224

Figure 4.12 Pure inulin fermentation using different strains of Clostridia species ...... 225

Figure 4.13 Fermentation of unhydrolyzed inulin extract media using C.

saccharobutylicum P262 (a) butanol, (b) total ABE production ...... 226

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Figure 4.14 Fermentation of unhydrolyzed inulin extract media using C.

saccharobutylicum P262 (a) acetic acid, (b) butyric acid ...... 227

Figure 4.15 Fermentation of unhydrolyzed inulin extract media using C.

saccharobutylicum P262 (a) acetone production, (b) change in pH ...... 228

Figure 4.16 Inulin consumption pattern by C. saccharobutylicum P262 (a) pure inulin P2

medium, (b) unhydrolyzed chicory inulin medium, (c) unhydrolyzed TKS Eskew

inulin medium ...... 229

Figure 4.17: Pure fructose fermentation using different strains of Clostridia species .... 231

Figure 4.18 Fermentation of 48 h enzymatically-hydrolyzed inulin extract media using C.

beijerinckii NCIMB 8052 (a) butanol production, (b) total ABE production ..... 232

Figure 4.19 Fermentation of 48 h enzymatically-hydrolyzed inulin extract media using C.

beijerinckii NCIMB 8052 (a) acetic acid, (b) butyric acid, (c) acetone production

...... 233

Figure 4.20 Cell growth of C. saccharobutylicum P262 in unhydrolyzed inulin media .235

Figure 4.21 Cell growth of C. beijerinckii NCIMB 8052 in 48 h enzymatically-

hydrolyzed inulin extract media ...... 235

Figure 4.22 Fermentation of 48 h and 1 h enzymatically-hydrolyzed pure inulin media.

...... 236

Figure D1 Raw TKS Extract before fermentation...... 297

Figure D2 Raw chicory extract before fermentation ...... 297

Figure D3 Pure inulin medium before fermentation by C. saccharobutylicum P262 ..... 297

Figure E1 Raw TKS extract after 72 h fermentation by C. saccharobutylicum P262 .... 299

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Figure E2 Raw Chicory extract after 72 h fermentation by C. saccharobutylicum P262

...... 299

Figure E3 Pure inulin medium after fermentation by C. saccharobutylicum P262 ...... 299

Figure F1 1 h enzymatically-hydrolyzed pure inulin medium before fermentation by C.

beijerinckii NCIMB 8052 ...... 301

Figure F2 1 h enzymatically-hydrolyzed pure inulin medium after 72 h fermentation by

C. beijerinckii NCIMB 8052 ...... 301

Figure G1 48 h enzymatically-hydrolyzed pure inulin medium before fermentation by C.

beijerinckii NCIMB 8052 ...... 303

Figure G2 48 h enzymatically-hydrolyzed pure inulin medium after 72 h fermentation by

C. beijerinckii NCIMB 8052 ...... 303

Figure H1 48 h enzymatically-hydrolyzed TKS Eskew Extract using endo-inulinase

(Novozyme 960) ...... 305

Figure H2 48 h enzymactially-hydrolyzed chicory extract using endo-inulinase

(Novozyme 960) ...... 305

Figure I1 HPLC chromatogram of standard sample containing inulin, kestose, sucrose,

glucose and fructose ...... 307

Figure I2 HPLC calibration standard curves (a) inulin, (b) kestose, (c) sucrose, (d)

glucose, (e) fructose ...... 307

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CHAPTER 1: INTRODUCTION

1.1 Background

Butanol (n-butanol or 1-butanol or butyl ) is a four carbon straight chain which has the molecular formula C4H9OH (MW 74.12) and a boiling point of 118°C. Acetone-butanol-ethanol (ABE) fermentation by Clostridia is a very well-known and long established industrial fermentation process, standing second in scale only to yeast-based ethanol fermentation (Green, 2011). After earlier attempts by

L. Pasteur and others, the fermentation of starch into acetone, butanol and ethanol was achieved through the discovery of a new bacterial species by C. Weizmann (Manchester

University, UK, 1912). The new species was named Clostridium acetobutylicum which naturally produces acetone, butanol and ethanol in a 3:6:1 ratio. The first production plant, which marked the beginning of industrial microbial fermentation of butanol, was based on the production of acetone from starch, rather than butanol. Acetone was the major compound of interest due to its use in the production of cordite (a smokeless explosive powder) during the First World War and butanol was the least desired fermentation byproduct. Owing to the strategic need for enormous amounts of acetone, and persistent problems with substrate delivery during the war, large-scale industrial plants were erected in the USA and Canada (Lutke-Eversloh & Bahl, 2011; Zverlov et al,

2006). After the war, however, butanol gained significance as a for quick-drying

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lacquer for the automobile industry. At the same time there was a shift in fermentation substrate from starch to molasses (Durre, 2008; Mravec et al., 2009).

During the 1950s and 1960s, the industrial ABE fermentation completely ceased in North America and Europe because of economic competition from petrochemical synthesis of butanol, recurrence of bacteriophage infections, poor molasses quality

(through improved sugar processing), and increased cost of molasses due to its increased use as animal feed. However, until the early 1980s a number of plants continued to operate in China, the Soviet Union and South Africa (Qureshi, 2011).

1.2 Rationale and Significance

After the oil crisis in the 1970s, there was a renewed interest in the production of biofuels. During that time, the primary focus for biofuel was on ethanol because of the familiarity with its production and its relatively high yield. Though ethanol production yield was higher than butanol, distribution of ethanol was, and is, difficult since ethanol cannot be transferred through existing pipeline infrastructures in any practical concentrations without corrosion and damage to rubber seals (Jin et al., 2011).

Nonetheless, in 2008, 70% of gasoline at the pump in the US had E10 gasoline (90 % gasoline blend with 10% ethanol) and a tiny portion of ethanol was being used to prepare

E85 (85% ethanol) to use in specially designed vehicles. However, because of lower calorific value, blending of ethanol reduces economy considerably. The reported mileage losses for ethanol blend gasoline fuels of E10, E15 and E20 are 3.88, 5.30 and

2

7.72%, respectively (NREL, 2008)1. This reduced mileage of ethanol-blend fuels are compensated by reduction in fuel cost per gallon and that is why ethanol fuel costs less than gasoline at the pump.

Over the last decade, rising crude oil prices, exhausting oil resources, and growing apprehensions over environmental problems (such as global warming, attributed to the use of fossil fuels), have brought considerable attention to the development of sustainable biofuels from biomass (Tashiro & Sonomoto, 2010). The U.S. Department of

Energy and USDA define sustainable biofuels as production of biofuels which are economically competitive, preserve the natural reserves, reduce greenhouse emissions and secure social well-being (U.S.DOE & USDA, 2009). The term biomass refers to all the organic matter existing on the earth produced by photosynthesis. Biomass is renewable, abundant and practically limitless, and its use is often regarded as carbon neutral. The combustion of biofuels derived from biomass releases fewer greenhouse gases, such as , than does combustion of fossil fuels. In the U.S. based on present yield of domestically harvested switchgrass, hybrid poplar, corn stover and wheat straw, Swana et al (2011) estimated that these materials can produce 8.27 billion gallons of biobutanol annually displacing 7.55 billion gallons of gasoline from the market. The merits of Biobutanol have generated renewed interest and its potential of being a sustainable alternate fuel is recognized in recent times.

Butanol is a fuel superior to biodiesel and bioethanol for many reasons. Butanol’s energy content is 30% more than ethanol (29.2 MJ/dm3 vs 19.6 MJ/dm3) and is close to

1 http://www.nrel.gov/analysis/pdfs/44517.pdf

3

gasoline (32 MJ/dm3). Also, its low vapor pressure facilitates its application in existing gasoline supply channels, it is less hydrophilic and it is less volatile, less hazardous to handle, and less flammable than ethanol. In addition, butanol can be used in unmodified internal combustion engines blended with gasoline at any concentration (up to 100 % v/v), instead of only 10% for ethanol2,3,4,5 (R.Szulczyk, 2010). Despite its superior fuel properties, in order to capitalize the large biofuel market, Biobutanol needs to compete on cost (based on energy basis) with ethanol (Green, 2011).

Currently, the commercial production of bioethanol begins with crops especially cultivated for this purpose such as corn in the United States, wheat and sugar beet in the

Europe, and sugar cane in Brazil. Virtually all commercially-produced ethanol is derived from starch or sucrose. Until lignocellulosic fermentation becomes widespread, utilizing these crops for biofuel production imposes immense pressure on arable land as well as on food and feed supplies. A life cycle assessment of corn to ethanol or 1-butanol processes shows that the net energy stored in 1-butanol is 6.53 MJ/L compared to a mere 0.40 MJ/L stored in ethanol (Swana et al., 2011).

Furthermore, butanol-producing solventogenic Clostridia like C. acetobutylicum or C.beijerinckii have the capability to consume both hexose and pentose sugars unlike conventional ethanol-producing yeast strains which can only use hexose sugars (Qureshi

& Ezeji, 2008). However, substrate cost still makes up at least 50% of the total

2 http://bioenergy.illinois.edu/pdf/Mr%20Butanol%20for%20CABER%20page.pdf 3 http://www.mypbic.org/butanol.html 4 http://www.ethanol.org/pdf/contentmgmt/March_07_ET_secondary.pdf 5 Ramey, David E., “Butanol: The Other Alternative Fuel,” ButylFuel, LLC.

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production cost in ABE fermentation, and the process economics, feasibility and sustainability are enormously contingent upon the availability of cheaper raw materials

(García et al., 2011; Qureshi & Ezeji, 2008). In the quest for low-cost raw materials over usual substrates like corn and cane molasses, many isolates and improved strains of

Clostridia have been developed over the years which significantly improve the range of sugars utilized. It has been demonstrated that several strains of Clostridia can consume a wide range of carbohydrate sources including starch, sucrose, glucose, fructose, galactose, cellobiose, xylose, arabinose, syngas and lower-cost carbon sources like glycerol, lactose, inulin and pectin, as fermentation substrates (Jang et al., 2012; Lee et al., 2008; Patakova et al, 2012; Qureshi, 2011; Qureshi & Ezeji, 2008; Zverlov et al.,

2006).

Recent findings have revealed that solventogenic Clostridia sp. are capable of converting furan aldehyde inhibitors, such as furfural and 5-hydroxymethyl furfural

(HMF), into less toxic furfuryl alcohol and HMF-alcohol (2,5-bishydroxymethylfuran) respectively. Furfural and HMF are well known microbial inhibitory compounds produced during degradation of lignocellulosic sugars, especially during acid hydrolysis.

But interestingly, HMF and furfural were found to have a stimulatory effect on Clostridia sp. growth and solvent production, rather than being inhibitory, at the lower concentrations of 1-3 g/L (Ezeji & Blaschek, 2008; Ezeji et al., 2007a; Qureshi et al.,

2012; Zhang et al., 2012).

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1.3 Current butanol production scenario

The current market price of 1-butanol is $ 4.0 – 4.57/ US gallon ($ 1446.15–

1492.31/t) (as per 16th July, 2012; obtained from www.alibaba.com). Though gasoline is a bit cheaper than butanol per gallon at present, if we compare the cost per thousand

BTUs, butanol is actually cheaper than gasoline. Presently, butanol costs around 2.96 cents per thousand BTU (same as ethanol), and gasoline costs 3.3 cents per thousand

BTU6. The worldwide production capacity of n-butanol is around 4.5 million tonnes/year and is worth over $10 billion. The butanol market is predicted to grow at a rate of 3.25% per year until 20257(Yuan & Hui-feng, 2012). According to the N-butanol Market

Research Report 2012, the worldwide total demand for n-butanol in 2011 was above 3 million tonnes which was 60,000 tonnes, a 2.1% increase from 20108. Gasoline can be replaced by 100 % butanol with the existing fuel infrastructure and Biobutanol has the potential to substitute for both ethanol and bio-diesel in the biofuel market, which is estimated to be worth $247 billion by 2020 (Green, 2011).

Currently, the average price of gasoline per gallon hovers around $3.0-3.2 (as of

12/20/12). However, prices are expected to increase worldwide due to unstable oil supplies from Middle Eastern countries and as increasing energy needs in developing countries, like China and India, intensify the competition for oil supplies. As per the US

EIA report (dated 10th July 2012), the total crude oil consumption by the US is 18.68 million barrels/day (22% of world oil consumption) whereas US production is only 6.31

6 webberenergyblog.wordpress.com 7 http://www.biofuelsdigest.com/bdigest/2012/01/20/2012-merger-mania-gets-underway-green-biologics- butylfuel-merge/ 8 N-Butanol (CAS 71-36-3) Market Research Report 2012, published by Business Analytic Center

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million barrels/day9. Transportation and industry consume a major portion (62 %) of all the petroleum used in the USA. In 2011, total biofuels production contributed only about

8% of U.S. transportation fuel consumption (gasoline and diesel combined) on a volume basis and only 6% on gasoline-equivalent energy basis (Schnepf, 2012). Notably, currently only supplant 2.5% of the world’s total oil consumption of 88 million barrels/day (Demain, 2009).

With the advent of engineered microorganisms capable of utilizing cheap, available lignocellulosic biomass, and developments in fermentation processes and improvements in downstream processing, it should be possible to produce biobutanol at less than one dollar per gallon in the near future10,11. In summary, biobutanol production is pivotal in reducing dependence, ensuring fuel security, preserving depleted fossil fuel reserves, and lessening pollution and environmental impacts. Biobutanol outweighs other biomass-derived liquid fuels in many aspects and promises to be a significant alternate fuel provided of the world’s energy needs. Also, owing to its ability for direct use or synergy with other hydro-carbon fuels for automotive fuel applications,

Biobutanol will facilitate the biofuels markets to thrive in future. This will directly influence the markets for agricultural substrates and could offer financial benefits for farmers (Nigam & Singh, 2011).

9 U.S. Energy Information Administration, Short Term Energy Outlook, Release date: 10 July, 2012. 10 Peswiki.com/index.php/Directory:Butanol 11 www.consumerenergyreport.com/2006/05/01/Biobutanol/

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1.4 Limitations of butanol fermentation

Although butanol possesses some interesting characteristics as a biofuel compared to the other low-carbon , the industrial production of butanol by ABE fermentation has several limiting factors. One such major limitation that largely affects the economy of ABE fermentation is butanol toxicity to the microorganisms employed resulting in low concentration of butanol in the fermentation broth which causes high product recovery costs (Garcia et al., 2011). Other problems associated with ABE fermentation are increased capital costs, sluggish , degeneration of microorganism, and possible phage infections (Garcia et al., 2011; Pfromm et al., 2010).

Some of the limitations of butanol as an alternate fuel are lower heating value compared to gasoline or diesel fuel (necessitates increased fuel-flow), lower octane number

(restricts the use of higher compression ratio and higher efficiency) and potential corrosiveness due to higher viscosity (Jin et al., 2011).

1.5 Research objectives

The goal of this study is to produce butanol from inexpensive and locally available substrates using selected strains of Clostridia. The study comprises two sections such as, 1) butanol production from industrial food processing wastes and 2) butanol production from inulin-rich biomass. A complete overview of this study is presented in Figure 1.1. The specific objectives and related tasks are described below.

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Objective 1: Estimate energy content and chemical composition of industrial food processing wastes and analyze their suitability for butanol production

Ohio is home to many food processing companies which are pioneers in production of specialty foods in the USA. Food and agriculture related sectors are the backbone of Ohio’s economy. Currently, Ohio ranks 7th in the nation in food processing

(4.5% of US total) with at least 1100 food processing plants.12 All these plants generate enormous amounts of wastes rich in carbohydrates, which are potential sources for butanol production feedstocks. We especially targeted those wastes that cause unwanted expenses to the industry either to treat or for their disposal.

We generated a list of food processing companies based on revenues in Ohio and approached only those companies which have annual revenue of more than $ 1 million.

Samples were collected from the volunteering food processing plants. The collected wastes were analyzed for their energy content, pH and chemical composition. It was proposed that the appropriate sample wastes for butanol fermentation would be chosen after categorizing the wastes based on the type of carbohydrate, energy value and the minerals present.

Objective 2: Investigate substrate preferences of selected Clostridium species for fermentation of food processing wastes and inulin-rich biomass

Several strains of Clostridia were screened for their substrate specificity by testing with the chosen food processing wastes, hydrolyzed inulin extract and pure monomers of the respective wastes. The strain which adapted to the substrates and

12 Ohio’s share of National Food Processing and Beverage production, US Census Bureau,2010.

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produced higher amounts of ABE in the screening was chosen as the best strain for butanol fermentation.

Objective 3: Optimize fermentation conditions of food processing wastes

Among the collected food processing wastes, those which have high carbohydrate content, minimal salt load, and can ferment without the need for hydrolysis, were chosen as the suitable substrates since the extra hydrolysis step incurs additional cost in the overall production cost of butanol. These wastes are generated in excessive amounts by the industries and the feasibility of using them directly without further processing or treatment for butanol fermentation gives the extra edge for their more productive and profitable use. Batch fermentation of these wastes was studied in detail.

Objective 4: Determine the optimum conditions for hydrolysis and enhanced butanol production from enzymatically-hydrolyzed inulin-rich biomass, a rubber crop co-product.

Natural rubber is one of the world’s most important commodities extensively used in various automotive, industrial, medical and other applications. The current global natural rubber production is around 10.82 million tons13. In 2011, the USA imported nearly 1 million tonnes of natural rubber (a year-on-year growth of 11%) at a cost of 4.7 billion dollars, which is 69% increase from 201014,15 (Data obtained from U.S

Department of commerce). The largest suppliers of natural rubber to the United States are Indonesia, Thailand and Malaysia, respectively. Natural rubber prices have increased

13 http://rubbermarketnews.net/2012/07/global-natural-rubber-production-up-4-9-may-weigh-over-prices/ 14 http://rubbermarketnews.net/2012/03/u-s-natural-rubber-imports-to-1-05-million-tons-in-2011/ 15 http://www.census.gov/foreign-trade/statistics/product/enduse/imports/c0000.html

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many folds over the past ten years and it was predicted that demand will exceed supply in

2020 by approximately 15%16. Imported finished rubber good costs ten times more than this amount. Currently, the major source of natural rubber is from the well-known Hevea brasiliensis rubber tree. In an attempt to develop commercially-viable natural rubber producing crops alternative to Hevea that are suitable for the climatic conditions of USA, coupled with the search for hypoallergenic rubber latex, research has focused on

Parthenium argentatum (guayule ) and Tarazacum kok-saghyz (TKS, Buckeye Gold, or

Russian dandelion). Of these two species, guayule is more suitable for hot and semi-arid climatic conditions (Southern USA) whereas TKS is suitable for moist and colder conditions (Northern USA). Ohio’s geographical location offers an excellent prospect of cultivating TKS.

TKS is a fast growing annual crop can contain rubber in the range of 2-20%17,18

(Kupzow, 1980; Whaley & Bowen, 1947) and 25-40 % inulin per dry root weight

(Buranov & Elmuradov, 2010; Schutz et al., 2006; van Beilen & Poirier, 2007). The ability of TKS to provide good quality rubber with high molecular weight similar to

Hevea, coupled with its co-product inulin-rich bagasse has increased renewed interest in commercialization of this crop (van Beilen & Poirier, 2007). Inulin belongs to a class of naturally-occurring reserve carbohydrates known as fructans which are polysaccharides of fructose molecules (β 2→1 linkages) with or without glucose as the terminal moiety.

In this study, we evaluated the amount of inulin extracted from the TKS roots during

16 www.oardc.osu.edu/penra/history.html 17 Kleinhenz et al., Abstract for 2008 annual meetings of the American society for horticultural science. www.oardc.osu.edu/penra/2008_meeting_ASHS.pdf 18 www.oardc.ohio-state.edu/images/E_Rubber.pdf

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latex extraction (water extraction at room temperature 25-28°C) and Eskew extraction

(hot water extraction at 95°C). In addition, a hot water inulin extract from chicory

(Cichorium intybus), which resembles TKS inulin in molecular weight, was used as a model substrate for inulin fermentation and the results were compared with TKS inulin.

Though acid hydrolysis is faster and cheaper than enzyme based hydrolysis, it does produce potent microbial fermentation inhibitors such as salts, hydroxymethyl furfural (HMF), furfural, and acetic, ferulic, glucuronic, ρ-coumaric acids, etc. For successful butanol production, these inhibitors have to be removed prior to fermentation

(Ezeji et al., 2007b; Qureshi et al., 2008; Varga et al., 2004; Zaldivar et al., 1999).

Considering the disadvantages of acid hydrolysis and the low pH of the hydrolyzate, we opted for enzymatic hydrolysis of inulin and estimated the amount of endo-inulinase enzyme (Novozyme) required for complete hydrolysis of inulin. We have attempted to produce exo and endo-inulinase enzyme from the strains of yeast Kluyveromyces marxianus and compared its activity against commercially available inulinase before choose the later one for enzymatic hydrolysis of inulin. We also developed a method for estimation of inulin and its derivative sugars in reversed phase chromatography (RP-

HPLC).

The materials and methods, results and discussion, findings and conclusion of butanol production from industrial food processing wastes and inulin-rich biomass are presented separately in chapter 3 and chapter 4, respectively.

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CHAPTER 2: LITERATURE REVIEW

2.1 Biofuels

The term ‘Biofuel’ has been ambiguously used in the literature in different parts of the world. Some refer only to liquid fuels (bioethanol & biodiesel) as biofuels while some include gaseous fuels and direct combustion of woody biomass in this category. In this paper, the term Biofuels refers to any liquid, solid or gaseous fuel produced from biomass, which can be rapidly renewed compared to fossil fuels (Bessou et al., 2010).

Biofuels refers to three forms of energy,

1. Liquid (Ethanol, Butanol, Biodiesel, Bio-oil)

2. Solid (Fuelwood, Charcoal, Agroresidues)

3. Biogases (Methane, )

Among the categories, liquid biofuels are regarded as the most marketable commodity of biofuels (Bessou et al., 2010). Biofuels are highly regarded as an alternate for fossil fuels. These are of major interest because of the renewability, biodegradability and production of acceptable quality exhaust gases upon combustion (Bhatti et al., 2008).

The contribution of biofuels in the automotive fuel market is promised to thrive in the next decade. The environmental protection agency renewable fuel standard 2 (EPA-

RFS2) requires the production of 36 billion gallons (136 billion liter) of renewable fuels in US market by 2022. The European Union mandates use of biofuels to replace 10%

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transportation fuel by 2020, while the USA mandate is 30% by 2030. Apart from that, biofuels have great potential in countries like Brazil, China, India, etc. (C. Jin et al.,

2011). The apparent advantage in the production of biofuels is the utilization of natural bioresources (Biomass is geographically more evenly distributed than fossil fuels) and the feasibility of generating bioenergy that could provide energy independence and energy security. The potential conflict between food and fuel can be mitigated by utilizing extensively available agricultural residues and waste substrates as the raw materials for fuel production. It was reported that biofuels produced from lignocellulosic materials produce lower net greenhouse gas (GHG) emissions than fossil fuels, thus helping reduce environmental impact (Nigam & Singh, 2011).

2.1.1 Current scenario of biofuels

According to International Energy Agency report, biofuel’s contribution to the world’s total primary energy supply was 10.2 % (1239.30 Mtoe) in 2009 whereas the biofuel share in total energy consumption was 12.9 % (1077.53 Mtoe) (IEA, key world energy statics, 2011)19 (Figure 2.1). Biofuels (liquid and gaseous fuels) play an important role in reducing CO2 emissions in the transportation sector. Currently, biofuels contribute only 2 % of energy share of transportation, however, due to rapid growth in the biofuels industry, it was predicted that biofuels would supply 27 % of the total transport fuel need by 2050. This anticipated use of biofuels could avoid generation of

2.1 giga tonnes (Gt) of CO2 emissions per year (IEA, 2011).

19 http://www.iea.org/textbase/nppdf/free/2011/key_world_energy_stats.pdf

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2.1.2 Classification of biofuels

There is certain amount of confusion on how to classify biofuels, but the common accepted classification has two divisions, namely, primary and secondary biofuels.

Primary biofuel refers to the energy obtained from unprocessed raw materials such as fuelwood, wood chips and pellets, etc., mostly for heating, and cooking, or for electricity generation in small and large industrial applications. On the other hand, secondary biofuels are derived from processed biomass, and have wider application in transportation and several industrial processes. Secondary biofuels can be produced in the form of solids (e.g. charcoal), or liquids (e.g. ethanol, biodiesel, butanol, and bio-oil), or gases

(e.g. biogas, synthesis gas and hydrogen) (Nigam & Singh, 2011). The secondary biofuels are further classified into first, second and third-generation biofuels based on the substrate availability, technology maturity and GHG gas emission balance (IEA, 2011;

Larson, 2008; Nigam & Singh, 2011). The major classifications of biofuels are shown in

Figure 2.2.

2.1.2.1 First generation biofuels

First generation biofuels are often termed conventional or traditional fuels whose processes are well-established and are already in commercial scale production. This generation of fuels is being produced through the least developed technologies. These fuels utilize well known feedstocks like starch from corn and wheat, sugars from sugarcane and sugar beet, oil crops such as soybean oil palm and rape (canola), and wastes like animal fats and used cooking oils. The most widely used first generation biofuels are ethanol from corn starch and cane sugar, and biodiesel produced through

15

processes from vegetable oils. Though these fuels are produced in enormous quantities in various parts of the world, they exhibit significant economic and environmental restrictions.

Since first generation fuels are produced from edible crops, it engenders competition with agriculture, food supply, water resources, and conflicts with land protection, thereby increasing the cost of feedstock and fuels produced. Also, vigorous use of land with rich fertilizer and pesticide applications and water use can cause ecological problems (Bessou et al., 2010; Carriquiry et al., 2011; Dragone et al., 2010;

IEA, 2011; Nigam & Singh, 2011). The feedstock for these fuels comprises 50 to 70 % of the production cost of ethanol, and 60 to 80 % for biodiesel (Bessou et al., 2010).

2.1.2.2 Second generation biofuels

These fuels are primarily derived from non-edible lignocellulosic biomass such as agricultural crop residues, and whole plant biomass (e.g. fast growing trees and grasses cultivated especially for energy production). Biofuels derived from vegetable oils that do not directly compete with food crops and agricultural land, are also considered second generation biofuels (e.g. jatropha, microalgae) (Carriquiry et al., 2011). Basically two different approaches are employed to produce second-generation biofuels i.e. biochemical (hydrolysis and fermentation) and thermochemical (pyrolysis or gasification) treatments of biomass (Bessou et al., 2010; Nigam & Singh, 2011; R. Sims et al., 2008).

Feedstocks grown especially for generation of these fuels enables higher biomass production per unit land area, and much of above-ground plant material can be used for biofuels. However, converting the lignocellulosic biomass into fermentable sugars

16

necessitates expensive technologies involving pre-treatment with specific enzymes.

Second generation fuels have already reached pilot and demonstration plants but have yet to be commercialized on a large scale level (Dragone et al., 2010; Eisentraut, 2010).

Though second generation biofuels promise benefits such as low-cost feedstock and better use of wastelands, they still need further improved conversion technologies

(Eisentraut, 2010; R. Sims et al., 2008).

2.1.2.3 Third generation biofuels

To overcome the challenges of first and second generation biofuels, researchers have focused their attention beyond agricultural substrates and waste vegetable oils to microscopic organisms. Third generation biofuels derived from microbes and microalgae are considered viable alternative energy resources that are devoid of the major drawbacks associated with first and second generation biofuels. The one distinct advantage of these fuels is the ability to grow the biomass in multiple or continuous harvests that could significantly increase yields. Microalgae have a very short harvesting cycle (≈1– 10 days depending on the process) compared with conventional crop plants which are usually harvested once or twice a year. Several technological breakthroughs and integrated conversion processes are essential to make this generation fuels more viable in future

(Bessou et al., 2010; Chisti, 2007; Dragone et al., 2010; Nigam & Singh, 2011; Schenk et al., 2008).

2.1.3 Liquid biofuels

Liquid biofuels include relatively familiar ones, such as biodiesel, bioethanol, biobutanol and less familiar fuels such as biomethanol, dimedimethyl ether (DME) or

17

Fischer-Tropsch liquids (FTL) made from lignocellulosic biomass. Bioethanol and biodiesel production increased tremendously in number of developed and developing countries. Between 1980 and 2005, global liquid biofuels production increased from mere

4.4 to 50.1 billion litres (Nigam & Singh, 2011). Alcohol fuels such as bioethanol, biobutanol and biomethanol can be a substitute for gasoline in spark-ignition engines, while biodiesel, green diesel (transformation of vegetable oil into fuel either by hydrogenation or hydrocracking) and dimethyl ether (DME) are appropriate to use in compression ignition engines.

In 2010, the global production of liquid biofuels (ethanol and biodiesel) was 105 billion litres (28 billion gallons US), that is 17 % increase from 2009. The world’s ethanol production reached 86 billion litres (23 billion gallons US) in 2010, 18% hike from 2009, whereas world’ biodiesel production was 19 billion litres (5 billion gallons

US), that is 12 % more than in 2009. The United States is the largest producer of ethanol in 2010 by generating 49 billion litres (13 billion gallons US), that amounts to 57% of global production and Brazil stands second with 28 billion liters (7 billion gallons

US)20,21. According to U.S. Energy Information Administration statistics, ethanol and biodiesel produced in the USA during 2011 had supplied 2.04 quadrillion BTU (QBTU) which is 22% of total renewable energy produced and 2.6 % of USA’s total energy production (78.16 quadrillion BTU)22,23 (EIA 2012, Table 1.2 Primary energy production

20 www.greencarcongress.com/2011/08/wwi-20110831.html 21 www.worldwatch.org/vitalsigns2012 22 www.eia.gov/totalenergy/data/monthly/pdf/sec1_5.pdf 23 Ww.eia.gov/totalenergy/data/monthly/pdf/sec10_3.pdf

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by source). The total primary energy consumption and production for the year 2011 in

USA are presented in Figure 2.3 and Figure 2.4.

While ethanol and biodiesel production account for nearly all of the total global liquid biofuels production and the technologies of these fuels are far more established than butanol, the ability of these fuels to contribute hugely to the rising global energy needs is still questionable. Ethanol or butanol production from corn and biodiesel production from soybeans all increase food prices, food scarcity, and several important environmental factors like soil erosion, loss of biodiversity, high volatile organic compound and NOx pollution. The net energy balance of ethanol production was debated for decades, while there were some reports discouraging expanded ethanol use due to negative net energy balance (Giampietro et al., 1997; Solomon et al., 2007), scores of recent reports demystified those claims and showed positive energy balance

(Goldemberg, 2007; Shapouri et al., 2002). Further improvement of energy balance of ethanol production warrants utilization of by-products generated from ethanol plants such as stillage for another biofuel production (aqueous by-product from the distillation of ethanol) (Murphy & Power, 2008; Wilkie et al., 200).

Biodiesel production is hugely affected by the availability and cost of raw materials (fats and oils), the value of the fuel produced, costs of processing (cost of raw materials accounts for 60 to 75% of the total cost of biodiesel fuel), expensive catalysts, poor quality fuel, and immature technologies to convert byproducts such as glycerol to useful products such as and ethanol (Almeida et al., 2012; Lim & Teong, 2010;

F. Ma & Hanna, 1999; Melero et al., 2009; Sheedlo, 2008; Vasudevan & Briggs, 2008).

The other notable disadvantages of biodiesel are: 1.5 times more expensive than fossil 19

fuels, degrades automotive rubber hoses, and emits 10% more NOx emissions than petro diesel fuels24,25,26. 1-Butanol is a four-carbon alcohol with excellent fuel properties closer to gasoline, has higher energy density than ethanol and possesses octane rating closer to gasoline and can be produced from more sustainable feedstocks than ethanol and biodiesel. Butanol is more environment friendly than gasoline because of its low vapor pressure (2.3 kPa vs 60-90 kPa for gasoline). Furthermore, butanol can be used directly in conventional internal combustion engines without modification and it is less hygroscopic which avoids corrosion problem when transferring through existing pipelines. All these attributes make butanol, a superior fuel to other biofuels

2.2 Production of Butanol

Though the industrial production of butanol began in the 1920s through fermentation of starch into butanol and acetone by the bacterium C. acetobutylicum, increasing demand for butanol as an industrial solvent and the dramatic growth of the petrochemical industry gave way to produce butanol through chemical process (S. Y. Lee et al., 2008).

2.2.1 Chemical Synthesis of Butanol

Butanol is conventionally produced through three major chemical processes namely, Oxo synthesis, Reppe Synthesis and crotonaldehyde hydrogenation from propylene (CH3CHCH2), carbon monoxide (CO), and hydrogen (H2). In oxo synthesis

() carbon monoxide and hydrogen are added to a double bonded carbon

24 http://www.berkeleybiodiesel.org/advantages-and-disadvantages-of-biodiesel.html 25 http://www.biodiesel-energy-revolution.com/disadvantages-of-biodiesel.html 26 http://howtopowertheworld.com/disadvantages-of-biofuels.shtml

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of propylene using metal catalysts such as Co, Rh, or Ru substituted hydrocarbonyls27

(García et al., 2011).

Oxo synthesis process

In the first reaction step, aldehyde mixtures are obtained, followed by hydrogenation to produce butanol. Different isomeric ratios of butanol can be obtained by varying the reaction conditions such as pressure, temperature and catalyst.

In the Reppe process, carbon monoxide, propylene and water are treated together with a catalyst that generates a mixture of n-butaraldehyde and isobutaraldehyde in which the n-butaraldehyde is reduced to n-butanol (Wackett, 2008). This process directly produces 85% - 88% n-butanol under air free conditions at 100°C and 0.5-2 MPa

(Chauvel & Lefebvre, 1989; Karl, 2008; Weissermel & Arpe, 2007). However, this process was not commercially successful because of the expensive technologies involved.

Reppe Process

27 Falbe, J. (1970). Carbon Monoxide in Organic Synthesis. Berlin-Heidelberg-New York; Springer Verlag.

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Until a few decades ago, the most commonly used route for butanol synthesis was from acetaldehyde using crotonaldehyde hydrogenation. This process comprises aldol condensation, dehydration, and hydrogenation.

Crotonaldehyde hydrogenation

Crotonaldehyde hydrogenation can produce butanol both in liquid and vapor phases. Though 100% crotonaldehyde conversion and butanol selectivity can be achieved through this process, selectivity of unsaturated alcohol is highly catalyst dependent. Lack of cheap, efficient catalyst and self-poisoning of the catalyst are the disadvantages of this process (Campo et al., 2008; Kun et al., 2001). While other methods of chemical synthesis of butanol rely completely on petroleum derived products, the crotonaldehyde hydrogenation can use ethanol instead, which can be produced from biomass. Ethanol is dehydrogenated into acetaldehyde from which the synthesis of butanol can proceed (S. Y. Lee et al., 2008).

2.2.2 Acetone-Butanol-Ethanol (ABE) fermentation

The first account of butanol production through microbial fermentation was reported by Louis Pasteur in 1861. He observed butanol along with butyrate while he was working on a newly isolated butyric acid producing strain. Thereafter, many researchers like Albert Fitz, Martinus Beijerinck, Bredemann, Schardinger and

22

Pringsheim carried out investigations on butanol-producing microorganisms and isolated several strains (Dürre, 1998; Gabriel & Crawford, 1930; García et al., 2011). However, it was only in 1905 that Schardinger reported the production of acetone by fermentation

(Jones & Woods, 1986).

At the commencement of the 20th century, there were huge demands for acetone and butanol for the manufacture of synthetic rubber, which led to intensive research efforts on production of butanol through fermentation. In 1911 Fernbach isolated and patented a culture that enabled the production of butanol from potato starch. In 1912, C.

Weizmann isolated a new bacterium (which was later named C. acetobutylicum) that was capable of utilizing starch and which gave higher yields of acetone and butanol then seen previously and eventually replaced Fernbach’s process. The first reported use of the name C. acetobutylicum was in 1926 by McCoy et al. and it became the officially recognized and accepted butanol-producing organism (Dürre, 1998; García et al., 2011;

Jones & Woods, 1986; McCoy et al., 1926). Several countries produced butanol through microbial fermentation at industrial scale during 1920-1980 using locally-isolated

Clostridial strains. However, many plants were forced to close during the 1960s due to increased cost of substrates, high product recovery costs, low solvent yields and competition from cheaper petrochemical synthesis of butanol from crude oil (Ezeji et al.,

2004; García et al., 2011; Kumar & Gayen, 2011).

The interest in butanol as a biofuel has regained importance in the last decade with a wide spectrum of research focused on genetic manipulation and development of strains, alternate feedstocks, and advancement in downstream processing of butanol

(Demain, 2009; Dürre, 1998; Dürre, 2008; S. Y. Lee et al., 2008; Ni & Sun, 2009; Nigam 23

& Singh, 2011; Qureshi & Ezeji, 2008; Swana et al., 2011; Y. N. Zheng et al., 2009).

Butanol contains 4- in its structure and is a more complex alcohol than methanol and ethanol, which have 1 and 2-carbon structures respectively. Furthermore, butanol has higher content than biodiesel, leading to further reduction of soot pollution.

Butanol emits lower NOx emissions, as well, because of its higher heat of evaporation, which results in a lower combustion temperature (C. Jin et al., 2011; D. Rakopoulos et al., 2010). The properties of butanol as a biofuel compared to ethanol, methanol, biodiesel and traditional fuels like gasoline and diesel are presented in Table 2.1.

Butanol exists as different isomers based on the location of the hydroxyl group (-

OH) on the carbon structure. The 4 carbon structure of butanol can form either a linear chain or branched structure that results in different fuel properties. It is reported that butanol produced through fermentation is normally a straight-chained n-butanol, also known as 1-butanol, which has the OH group attached to the terminal carbon. The other straight chained butanol is 2-butanol (also known as sec-butanol) which has the hydroxyl group (-OH) attached to an internal carbon.

Iso-butanol is a branched with the OH group at a terminal carbon and tert- butanol refers to the branched isomer with the OH group at an internal carbon. All these butanol isomers can be produced through chemical synthesis from fossil fuels by the different methods described earlier in the Chemical Synthesis of Butanol Section. The properties and applications of different isomers of butanol are compared and presented in

Table 2.2 and 2.3 respectively.

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2.2.3 General description of Clostridium species and biobutanol production

Clostridia are rod shaped, obligate anaerobic, spore-forming, gram positive organisms, which are motile and heterofermentative in nature. The characteristic

Clostridial fermentation has biphasic fermentation i.e. an acidogenic phase and a solventogenic phase. The acidogenic phase takes place during the exponential growth phase of the organism and, during this phase, acid-forming pathways are activated and form carboxylic acids, mostly acetate and butyrate (Gholizadeh, 2010). These acids lower the external pH and serve as inducers for the biosynthesis of the solventogenic enzymes. The high cell growth during the acidogenic phase is because of production of high amounts of ATP. During the solventogenic phase, these acids reassimilate and function as co-substrates for the production of solvents i.e. acetone, butanol and ethanol

(isopropanol instead of acetone in some Clostridium beijerinckii strains). The production of acids and cell growth cease during solventogenic phase and the pH of the fermentation medium increases marginally because of the acid uptake. This sudden metabolic shift during the solventogenic phase is attributed to the dramatic change in gene expression pattern which is believed to be an adaptive response of the cells to the low external pH resulting from acid production. The exponential growth of Clostridia during acidogenic phase and spore formation during solventogenic phase is depicted in Figure 2.5.

Once the cells are shifted to solventogenesis, more carbon and electrons are directed to the formation of solvents where butanol is the major fermentation product.

Apparently, enzyme synthesis during acidogenic and solventogenic phases and control of electron flow are critical with respect to regulation of acetate, butyrate and butanol formation. Since the electron flow can be reversed, this substantiates the hypothesis that 25

butanol yield could respond to factors that influence the direction of electron flow. Based on this theory, many researchers have worked on electron carriers, like addition of carbon monoxide, methyl viologen, and neutral red to the fermentation medium, and butanol formation was stimulated at the expense of acetone synthesis (Dürre, 2011; Gheshlaghi et al., 2009; Gholizadeh, 2010; Green, 2011; Y. S. Jang et al., 2011; C. Jin et al., 2011; M.

Kumar & Gayen, 2011; S. Y. Lee et al., 2008; Lütke-Eversloh & Bahl, 2011; Qureshi &

Ezeji, 2008; Yu et al., 2011).

2.2.4 Fermentative pathways of Clostridia

In general Clostridium sp. consume hexose (glucose, fructose and galactose) and catabolize these sugars by the Embden–Meyerhof–Parnas (EMP) pathway, whereas pentoses (xylose and arabinose) are catabolized by the pentose phosphate pathway to finally produce pyruvate, ATP and NADH. Pyruvate is subsequently converted into acetyl coenzyme A (acetyl-CoA) by pyruvate-ferredoxin oxidoreductase (PFOR).

Oxidative decarboxylation of pyruvate by PFOR produces one reduced ferredoxin molecule, which has a more negative redox potential (E0’ ≤ –400 mV) than that of

NADH (E0’ = –340 mV). Later, this reduced ferredoxin acts as an electron donor either to reduce nicotinamide adenine dinucleotide (NAD+) to NADH by NADH ferredoxin oxidoreductase or to produce H2 by transferring electrons to the hydrogenase complex.

Acetate is synthesized via phosphotransacetylase and acetate kinase reactions with the latter reaction providing ATP.

For the biosynthesis of butyrate, two molecules of acetyl- CoA are condensed to acetoacetyl-CoA, followed by a reduction to butyryl-CoA, which is then converted to butyrate via phosphotransbutyrylase and butyrate kinase reactions with ATP generation. 26

Acetate and butyrate are reassimilated to their corresponding CoA derivatives catalyzed by the acetoacetyl-CoA:acyl-CoA transferase, with acetoacetyl-CoA as the CoA donor.

When reducing equivalents availability is limited, acetoacetate is decarboxylated to acetone in order to drive the transferase reaction by acetoacetate removal. and butanol dehyrdogenase activities, which can be provided by different dehydrogenases, convert butyryl-CoA to butyraldehyde and finally to butanol (Fig. 2.6)

(Dürre, 2011; Herrmann et al., 2008; Y. S. Jang et al., 2011; C. Jin et al., 2011; M.

Kumar & Gayen, 2011; F. Li et al., 2008; Lütke-Eversloh & Bahl, 2011; Qureshi &

Ezeji, 2008; Tashiro & Sonomoto, 2010). Schematic representation of metabolic pathway of C. acetobutylicum is presented in Figure 2.6.

2.2.5 Strain development

Several solventogenic Clostridia have been investigated on the molecular level over the years and various strains of industrial solvent-producing Clostridia belong to cluster I of the Clostridia and were classified into four species by similarity of their 16S rDNA sequences and DNA–DNA homology. The four distinct species are: C. acetobutylicum, C. beijerinckii, C. saccharobutylicum and C. saccharoperbutylacetonicum. The members of the four species differed considerably in solvent-producing ability (between10 and 24 g l-1) and solvent yield (between 6.8 and

33.2%).

Most of the studies undertaken over the last two decades have focused on strains belonging to the species C. acetobutylicum and C. beijerinckii. Till now, C. acetobutylicum ATCC 824 remains the best studied and manipulated strain, although this species group is quite distinct, both genetically and physiologically, from the three other 27

main solvent producing-species: C. saccharobutylicum, C. beijerinckii and C. saccharoper-butylacetonicum. These solventogenic Clostridial strains are characterized based on the type of substrate utilization, solvent productivity, butanol tolerance, and the ratio and type of solvent production (Ezeji et al., 2004; Green, 2011; Lütke-Eversloh &

Bahl, 2011; Shaheen et al., 2000; Zverlov et al., 2006). The other notable commercial strains include C. saccharobutylicum P262 (formerly known as acetobutylicum 262), C. beijerinckii P260, and C. beijerinckii NRRL 292 (Green, 2011; Kumar & Gayen, 2011).

The mutant strains are usually generated by using chemical/physical mutagenesis strategies applied to solventogenic Clostridia. A novel mutated strain (MEMS-7), reported to be the best strain for fermenting molasses, was developed from C. acetobutylicum. This strain was obtained by treating the parent organism with N-methyl-

N-nitro-N-nitrosoguanidine and ethyl methane sulphonate, under UV exposure. MEMS-

7 strain was reported to produce 20% more butanol than the parental strain (Kumar &

Gayen, 2011; Syed et al., 2008). Another mutant strain EA2018 was also developed from

C. acetobutylicum through chemical mutagenesis and found to produce higher butanol:solvent ratio (Butanol:Acetone:Ethanol = 7:3:1) and a total solvent concentration of up to 18–22 g/L (Ni & Sun, 2009).

Further increase in butanol:solvent ratio (8:3:1) was observed in EA2019 when the acetone pathway was knocked out (Green, 2011; L. M. Harris, Desai, Welker, &

Papoutsakis, 2000; L. Harris, Blank, Desai, Welker, & Papoutsakis, 2001; Y. Jiang et al.,

2009; Ni & Sun, 2009). Transcriptomic characterization of the strain revealed higher transcription level of genes responsible for butanol production and reduced expression level of acetone formation gene (Kumar & Gayen, 2011; Ni & Sun, 2009). 28

Another notable mutant strain is C.beijerinckii BA 101 generated from

C.beijerinckii NCIMB 8052 (formerly called as C. acetobutylicum) by applying N- methyl-N9-nitro-N-nitrosoguanidine (NTG) together with non-metabolizable glucose analog 2-deoxyglucose (2-DOG). The NTG treated cells resuspended in a specially made

P2 medium with 5 g/L starch and 1 g/L of 2-DOG were found to have increased the amylolytic activity to1.8 to 2.5 fold in C. beijerinckii BA 101 than the parent strain and also exhibited higher butanol producing capability (19-20 g/l) (Annous & Blaschek,

1991; Ezeji et al., 2004; Formanek et al., 1997; Kumar & Gayen, 2011; Qureshi &

Blaschek, 2000; Qureshi & Blaschek, 2001; Qureshi, et al 2008).

Apart from strain improvement by mutation, elucidation of butanol and acetone producing genes have paved the way for genetic manipulations, such as gene knockout or overexpression of genes to give superior performance and increase solvent production.

To date, the whole genome of C. acetobutylicum ATCC 824 and C.beijerinckii NCIMB

8052 were sequenced in 2001 and 2007, respectively (Green, 2011; Tashiro & Sonomoto,

2010; Zverlov et al., 2006). The solvent-producing genes are located on the chromosome in C. beijerinckii NCIMB 8052 whereas these genes are found on a megaplasmid in C. acetobutylicum ATCC 824. Degeneration of bacterial cells in C. acetobutylicum during extended fermentations is attributed to the loss of plasmid whereas the cause for degeneration in C. beijerinckii is not yet found (Y. Wang, Li, Mao, & Blaschek, 2012;

Zverlov et al., 2006).

In Clostridial fermentation, during the solventogenic phase, cell growth slows down and formation of spores occurs, and the cells eventually cease butanol production.

In C.acetobutylicum ATCC 824, the formation of spores hampers solvent production 29

capability of the organism when the cells are used in continuous culture or are subjected to repeated sub-culturing (Assobhei et al., 1998; Kumar & Gayen, 2011; Sillers et al.,

2008). This phenomenon is because the transcription factor responsible for initiation of sporulation (Spo0A) also initiates solvent production. Though development of non- sporulating strains such as C. acetobutylicum DSM 1731, M5 and DG1 were attempted and they produced butanol at lower levels than their parental strains, while no acetone was produced because these strains do not carry the genes necessary for acetone formation (Cornillot et al., 1997; Nair & Papoutsakis, 1994; Papoutsakis, 2008). Mutants without Spo0A are severely deficient in solvent production while strains with amplified

Spo0A overexpress solventogenic genes but fail to produce more solvent due to an accelerated sporulation process. If the ability to use Spo0A to activate the solventogenic genes without activating sporulation function is developed, then it can effectively be used to enhance solvent production (C. Jin et al., 2011; S. Y. Lee et al., 2008; Lütke-Eversloh

& Bahl, 2011).

2.3 Alternate feedstocks

2.3.1 Lignocellulosic biomass

Cellulosic biomass, also called as lignocellulosic biomass, is a major component of biomass that makes up almost half of the matter produced by photosynthesis (Pérez et al., 2002). Lignocellulose is a naturally available heterogeneous complex carbohydrate consisting of microfibrils embedded in lignin, hemicellulose and pectin, with different amounts of each component in different plant species and among plant parts of a single species (Figure 2.7). Cellulose is a crystalline linear polymer composed of 1, 4 β-

30

glucosidic linkages of anhydro-D- glucose. The linear chains of glucose units are aligned parallel to each other (called elemental fibrils) and linked to each other by a large number of strong hydrogen bonds and van der Waals forces. The microfibrils are made up of aggregates of long bundles of elementary fibrils. Both hemicellulose and lignin are considered to be in microfibrils (Nigam & Singh, 2011; Pérez et al., 2002; Srinivasan,

2010). Although cellulose normally exists in crystalline form there are still a small percentage of non-organized cellulose chains that are amorphous in nature.

Hemicellulose contains lower molecular weight polysaccharides than cellulose, including pentoses (D-xylose, L- arabinose), hexoses (D-mannose, D-glucose, D-galactose) and uronic acids (4-O-methyl-glucuronic, D-galacturonic and D-glucuronic acids). The predominant linkage in hemicellulose is a β-1-4-linkage, but sometimes β-1-3- glycosidic linkages are observed. The type of hemicellulose in hardwood is glucuronoxylan whereas glucomannan is present in softwoods (Girio et al., 2010; Pérez et al., 2002).

Lignin is the most complex, most abundant and least characterized polymer in nature. Structurally, lignin possesses high molecular weight non-water soluble heteropolymers and is amorphous in nature. The function of lignin in plants is to provide structural strength, impermeability, oxidative stress and resistance against microbial attacks. Lignin is composed of phenylpropanoic acid units joined together in three dimensional structures with different types of linkages. Lignin polymerization is synthesized through a free radical mechanism which is reason for random and highly non-homogenous nature of lignin composition and structure. Coniferyl alcohol is the major component in softwood lignins, whereas in hardwood the principal components are guaiacyl and syringyl alcohols (Pérez et al., 2002; Srinivasan, 2010). Because of the 31

complexity in the structural make-up, lignin is highly resistant to chemical and microbial degradation. Except few fungi, no other microorganisms have the required enzymes to break lignin apart. Moreover, it was found that lignin degrades only in aerobic conditions while in anaerobic environment it can persist for longer periods (Y. C. Sun et al., 2012;

Van Soest, 1994).

Cellulose can be consumed by few eubacteria and fungi, however, most of the organisms do not have cellulolytic activity to degrade cellulose into hexoses and pentoses that can be utilized by the cells. To overcome this problem, pretreatment of lignocellulosic biomass is commonly employed to increase the surface availability of cellulose for enzymes and reduce its crystallinity, thereby increasing the formation of fermentable sugars through enzyme hydrolysis. The three common modes of pretreatments available are physical (grinding/milling), chemical (application of solvents/acid) and biological (cellulolytic microorganisms). Among them, the most prevalent methods are alkali treatment, concentrated acid hydrolysis, dilute acid hydrolysis, steam explosion, ammonia explosion and supercritical fluid treatments. The characteristics of an ideal and effective pretreatment are: maximum sugar recovery, minimum formation of toxic substance during degradation, low capital cost and low energy demand (Alvira et al., 2010; Nigam & Singh, 2011; Prasad et al., 2007;

Srinivasan, 2010; von Sivers & Zacchi, 1995).

Srinivasan & Ju (2010) reported that supercritical CO2 pretreatment followed by enzyme hydrolysis gave a 77 % conversion of glucose and 86% of total reducing sugars from guayule bagasse. These results surpassed dilute-acid pretreatment and delignification pretreatment previously employed to saccharify guayule bagasse. The 32

term bagasse here refers to the remaining biomass after the latex extraction through wet milling (Srinivasan & Ju, 2010). Leaf stream biomass has higher cellulose and hemicellulose contents (and less acid insoluble materials) than bagasse (Table 2.4). The higher acid insoluble contents in bagasse might be due to the remaining rubber and resins not completely removed by the wet milling and extractions, but has not yet been verified

(Srinivasan & Ju, 2010). Boateng et al (2009) estimated the elemental composition of guayule biomass which is presented in Table 2.5. Inedible, high energy, biomass from industrial crops like guayule, offers great potential for production of biofuels like butanol.

Research on direct fermentation of cellulose-containing biomass with cellulolytic solventogenic Clostridia to butanol in a single step has gained prominence over the last few years. Clostridium thermocellum and C. cellulolyticum are well-known cellulose degrading thermophilic solventogenic Clostridia, and they majorly produce ethanol

(Demain, Newcomb, & Wu, 2005). Another thermophilic species C. thermosaccharolyticum can produce ethanol and butanol almost in equal amounts

(Patakova et al., 2012).

It was recently found that several Clostridia species such as C. acetobutylicum,

C.thermupapyrulyticum and some Clostridium sp. naturally have either the cellulose degrading extracellular enzyme complex, called cellulosome, or cellulolytic activity.

This feature enables the organisms to directly degrade cellulose for solvent production.

Among them, C. thermupapyrulyticum and some Clostridium sp., reportedly utilize cellulose directly to butanol but still these organisms need to be studied in more detail

(Mendez et al., 1991; Virunanon et al., 2008). However, C. acetobutylicum is ineffective

33

in degrading cellulose, apparently because of little active mini-cellulosomes which led to lower expression levels of cellulosome enzymes.

Attempts to develop an active mini-cellulosome in C. acetobutylicum was made by inserting a gene man5k (encoding the mannanase Man5K) from the thermophilic bacterium Clostridium cellulolyticum and it was found that the engineered strain secreted functional mini-cellulosome. These results suggest that cellulosomal genes can be transformed into C.acetobutylicum and other strains in an effort to develop cellulose utilizing microorganisms. However, optimal expression and secretion of the heterologous mini-cellulosome in the engineered strains remain to be the bottle-neck.

Overcoming this obstacle will allow researchers to radically improve the butanol production from abundantly and inexpensively available lignocellulosic biomass (Bayer et al., 2004; Y. S. Jang et al., 2011; López-Contreras et al., 2004; Mingardon et al., 2005;

Perret et al., 2004; Sabathé & Soucaille, 2003; Virunanon et al., 2008; Zverlov et al.,

2006).

Though starch and sugar based feedstocks are used predominantly as common feedstocks for butanol production, the ability of Clostridia to consume hexose and pentose sugars make them well suited for fermenting sugars derived from pretreated lignocellulosic biomass. Using cheaper agricultural residues, or wastes such as corn cobs, corn stover, packing peanuts, orchard waste, dried distillers grains and soluble

(DDGS), corn fiber, sugar cane bagasse, wheat straw, barley straw, grass and municipal solid waste (MSW), confer a more sustainable option to starch and oil, offering a lower carbon footprint, reduced greenhouse emissions, in addition to the significantly reduced cost of the feedstocks (Green, 2011; Kumar & Gayen, 2011; Nigam & Singh, 2011; 34

Qureshi, 2011). Recent investigations of Clostridia revealed that these bacteria can secrete carbohydrate degrading enzymes such as amylase, xylanse, β-glucanase, β- glucosidase, invertase, glucosidase, glucoamylase, pullulanase, amylopullulanase and both extracellular and intracellular β-fructofuranosidase (inulinase) to digest complex polysaccharides into simple monosaccharides that can be transported into the cell for subsequent metobolization (Ezeji et al., 2007; Kumar & Gayen, 2011; Patakova et al.,

2012).

2.3.2 Glycerol utilization

The asporogenous mutant C. pasteurianum DSM 525 produced 1-butanol

(0.44g/g of substrate) from ethanol stillage that contained glucose, glycerol and lactic acid as carbon source (Ahn, Sang, & Um, 2011). Another strain of this same species C. pasteurianum ATCC 6013TM produced 1-butanol, 1,3-propanediol and ethanol when pure or crude glycerol was used from biodiesel production (Taconi et al., 2009). Strains of C.acetobutylicum can utilize glycerol only if it is mixed with glucose in the medium.

In these strains, glycerol inhibited hydrogen formation and eliminates acetone synthesis while increasing 1-butanol and ethanol production. The rationale behind this phenomenon is the necessity to regenerate double the amount of NAD+ than the amount generated using glucose alone (Andrade & Vasconcelos, 2003; Patakova et al., 2012;

Vasconcelos et al., 1994). Zverlov et al. reported that C. ljungdahlii can utilize carbon monoxide and hydrogen from synthesis gas (reformer gas) as a carbon source for butanol production (Zverlov et al., 2006).

35

2.3.3 Microalgae

Another promising feedstock for biofuels production is microalgae. The definition of microalgae is unicellular and simple multicellular structure from both prokaryotic and eukaryotic microorganisms. Microalgae can be autotrophic or heterotrophic. Autotrophic organisms absorb sunlight, nutrients from the aquatic habitats, and assimilate carbon dioxide and store carbohydrates in them that can be used as biomass. Microalgae reproduce themselves; their entire life-cycle completes in a few days, requires less attention, have year round biomass production and can grow in different environmental conditions in both aquatic and terrestrial environments (Brennan

& Owende, 2010; Demirbas, 2010; Frac et al., 2010; Mata et al., 2010). Also, they can serve to bioremediate polluted environments and biofertilizers through binding of atmospheric nitrogen. Microalgae, depending on species, produce lipids, proteins, hydrocarbons, and some complex oils that can be possibly used to generate biofuels.

Currently, biodiesel production from algal oils has been the major interest because of the abundant availability of renewable microalgal biomass and rapid growth (Chisti, 2007;

Frac et al., 2010; Williams & Laurens, 2010).

Many algae contain more than 50% (dry weight) of starch, cellulose and glycogen in its composition, especially in the cell walls. The harvested algae can be a good substrate for several saccharolytic Clostridium species. Ellis et al. demonstrated ABE production from wastewater algae biomass using C. saccharoperbutylacetonicum N1-4.

From10% pretreated algae, this strain produced 2.74 g/L of total ABE whereas 7.27 g/L and 9.74 g/L of total ABE was produced when the pretreated algae is supplemented with

1% glucose and enzymes (xylanase and cellulase), respectively. Attempt to use non- 36

pretreated algae produced only meager 0.73 g/L of Total ABE (Ellis, Hengge, Sims, &

Miller, 2012). Selection of the appropriate microalgae species, engineered novel microorganisms, development in cultivation techniques, better understanding of algal biomass, economic feasibility and viability of continuous production of biofuels at low cost, are the important challenges that impedes development of algal biofuel technology

(Brennan & Owende, 2010; Pienkos & Darzins, 2009; Scott et al., 2010).

2.3.4 Food processing wastes

The food processing industry sector has attained greater significance over the years and has become pivotal for the U.S. economy. In 2007, US food processing industries exported products worth $38.7 billion and imported $34.7 billion (Source: U.S

Department of Commerce)28. The food processing industry primarily uses biological materials (agricultural, animal or aquatic) as its raw materials and consumes significant amounts of energy and water. Processing of raw materials into products, however, generates large quantities of organic wastes that are mostly biodegradable. Food processing wastes may generally contain large amounts of sucrose, starch and cellulose that could supply fermentable sugars for production of biofuels while improving sanitation, and reducing environmental impacts caused by waste disposal while minimizing energy demands.

Food industry wastes can be categorized into three groups:

(1) Losses during manufacturing/processing food

(2) Food products thrown away as municipal solid waste (MSW)

28 U.S. Department of Commerce Industry Report, 2008 http://trade.gov/td/ocg/report08_processedfoods.pdf

37

(3) Discarded cardboard, wrappers, containers, and tins

These three categories can be further divided into liquid and solid wastes (Ukita et al., 2005). Except for metal containers, tins, vessels, and wrappers, almost every other food wastes, with or without hydrolysis, may be effectively used as a substrate for anaerobic fermentation. The major classifications of food processing wastes based on type of industry and their possible ways of utilization are presented in Table 2.6.

Voget et al. tested C. acetobutylicum NRRL B596 and strains of C. beijerinckii

NRRL B592, B593 for butanol production from apple pomace. He found that the apple pomace contains 10.8% of total carbohydrates (fructose: 67%, glucose: 23% and sucrose:

10%) and the strains yielded between 1.9 and 2.2 % of butanol when the sugar content from the apple pomace was adjusted to 40 g/L (Voget et al., 1985).

The solvent-producing ability of several Clostridial strains from starch medium

(sugar content: 45-48 g/L) made from unhydrolyzed and hydrolyzed (using amylolytic enzymes) potatoes was tested. It was found that there was no significant difference in solvent production for most of the strains between unhydrolyzed and hydrolyzed starch medium. Strains of C. acetobutylicum DSM 1731, C. beijerinckii NCIMB 8052 and

NRRL 592 were found to have good solvent yield. Strains of C. acetobutylicum ATCC

824, NCP 260, and NCP 262 produced little solvent and were considered to be deficient in amylolytic activity. Tests using external enzymes to hydrolyze starch into free glucose also did not improve solvent production of these strains. Gutierrez et al. concluded C. acetobutylicum DSM 1731 was the most productive strain with highest productivity of

0.24 g/L/h on unhydrolysed potato (Gutierrez et al., 1998). Grobben et al (1993) observed that C. acetobutylicum DSM 1731 produced maximum total ABE of 19 g/L 38

after 30h when 14% (w/v) potato powder was used as a medium. Direct perstraction

(perstraction uses permeable membrane to separate desired products from the fermentation broth using extraction solvents) and microfiltration perstraction methods were evaluated to find a suitable integrated solvent recovery method. Direct perstraction using a polypropylene and /decane mixture as the extractant gave higher product yield (total ABE/ potato dry weight). The product yield increased to 77% (0.13 g/g to 0.23 g/g) and 46% (0.13 g/g to 0.19 g/g) using direct perstraction and microfiltration perstraction, respectively (Grobben et al., 1993).

Direct fermentation of sago starch by C. acetobutylicum P262 in batch scale was investigated by Madihah et al. (2001). They found that solvent production from 30 g/L sago starch was comparable to fermentation with corn starch whereas it was twice the solvent production on potato and tapioca starch. The highest total ABE production from sago starch was 18.82 g/L obtained at the medium concentration of 50g/L starch. The study revealed that individual concentrations of nitrogen and carbon in the medium influenced the solvent production to a greater extent than the carbon to nitrogen (C/N) ratio. Addition of inorganic nitrogen source (yeast extract + NH4NO3) improved starch consumption by C. acetobutylicum P262 and increased total ABE production to 24.47 g/L

(i.e. total solvent yield of 0.45 g/g of substrate consumed). Increased starch concentration in the medium above 70g/L was found to decrease solvent production significantly. The possible reasons for this phenomenon are the unavailability of a sufficient amount of enzymes to hydrolyze high starch concentrations to fermentable sugars and increase in apparent viscosity of the medium with an increase in starch concentration. Gelatinized starch, in the bottom of the bottle after autoclaving high 39

starch concentrations, is known to resist enzymatic and biological reactions.

Continuously stirred tank reactors might help complete mixing of the medium for better performance (Madihah et al., 2001).

Fermentation of 60g/L whey-permeated medium (lactose concentration of 48.4 g/L) using C. acetobutylicum P262 in a batch reactor resulted in total ABE production of

9.34 g/L after 120 h. Increasing the lactose concentration up to 225 g/L did not inhibit the solvent production. However, further increases in lactose concentration negatively affected the fermentation and drastically reduced solvent production. Continuous solvent recovery by perstraction using oleyl alcohol produced 98.97 g/L of total ABE while the lactose concentration was 227 g/L, giving a yield of 0.44g/g and a productivity of 0.21 g/L/h (Qureshi & Maddox, 2005). For continuous steady-state solvent production, it is imperative to maintain a balance between the acid-producing vegetative cells and the solvent-producing cells (Ennis & Maddox, 1989).

Foda et al. compared the butanol-producing ability of C. acetobutylicum DSM

792 and C. acetobutylicum AS 1.224 in a batch reactor using lactose and cheese whey medium. C. acetobutylicum DSM 792 was identified as the best strain and it performed better in cheese whey medium than pure lactose medium (Foda et al., 2010).

Immobilized C. beijerinckii LMD 27.6 cells in whey permeate medium at fermentation temperature of 30°C and a dilution rate of 0.1 h-1 was found to be ideal for continuous production of butanol with higher yield (Schouten et al., 1985).

C. beijerinckii BA101 produced total ABE of 24.7 g/L, 21.7 g/L and 20.2 g/L from different media using pure starch, waste packing peanuts and model agricultural waste, respectively (Jesse et al., 2002). Soy molasses, a by-product of the soy processing 40

industry, was tested for butanol production because of its relatively high carbohydrate content. Clostridium beijerinckii BA101 produced 10.7 g/L total ABE when 80 g/L spray dried soy molasses was used in the medium. Total ABE production increased to

23.8 g/L when the medium was supplemented with 25.3 g/L glucose. Qureshi et al. found that C. beijerinckii BA 101 could utilize glucose, sucrose, fructose, galactose, maltodextrin but was unable to ferment raffinose, pinitol, verbascose and stachyose because this strain lacks α-1-6 glycosidase enzymatic activity. Addition of tri-calcium phosphate (28.8 g/L) improved total ABE production from 23.8 to 30.1 g/L but addition of sodium chloride above 10 g/L was found to inhibit solvent production (Qureshi et al.,

2001).

Bread waste and brewer’s spent grain (BSG) was tested for bioethanol production using Saccharomyces cerevisiae. The bread waste was used in the medium without any pretreatment (because of lesser lignocellulose content) while the milled brewer’s waste was pretreated with dilute acid followed by delignification with NaOH, used to help improve the sugar availability to the microorganisms (Olugbenga et al., 2011). Co- fermentation of different bakery products like, potato chips, wheat flour and cheese whey, hydrolyzed using enzymes, were found to increase ethanol production and decrease processing time from 60 h to 12 h (J. V. Kumar et al., 1998). Production of ethanol from wheat flour and wheat bran after hydrolyzing them using mixtures of amylolytic enzymes was also demonstrated (Neves et al., 2006; Pejin et al., 2006).

Waste products from beverage industry, such as Buckwheat tea waste (BWTW) and barley tea waste (BTW), were tested for simultaneous saccharification and fermentation by S. cerevisiae and Mucor indicus to produce ethanol (Sasaki et al., 2012). 41

Li et al. attempted simultaneous saccharification and fermentation of easily available food wastes in Korea using S. italicus KJ to produce ethanol. At a temperature of 35°C, both enzymatic hydrolysis and fermentation were carried on simultaneously with a reducing sugar consumption rate of 3.61 g/L.hr (Li et al., 2011).

Banana fruit stalks contain approximately 57% total sugar of which 27% is starch and approximately 24% is cellulose. The orange peel contains fibers (11.8% of DM), reducing sugars (9% of DM) and protein (6.4%). Effluents generated in the potato processing industry during production of potato chips, slices, and shred potatoes have high starch content (19.5 g/l) and also have little protein (2.9 g/l) (Thomsen, 2005). All these wastes, including the wastes that have been tested for ethanol fermentation can be effectively used for ABE fermentation without the need for hydrolysis.

2.3.5 Inulin as substrate

Butanol production from inulin polymer (a polyfructose) using strains of

Clostridia was investigated in the early 1900s. The inulinic structure has to be broken down by acid hydrolysis or by application of inulinase prior to fermentation to facilitate availability of fermentable sugars. Marchal et al. investigated C. acetobutylicum ATCC

824 and two selected strains C. acetobutylicum IFP 902 and IFP 904 on enzymatically hydrolyzed inulin extract from Jerusalem artichoke (Helianthus tuberosus). The strain C. acetobutylicum IFP 904 was isolated from Jerusalem artichoke tubers. It was found that this strain naturally possesses little inulolytic activity and has butanol-resistant, high solvent-producing capability. Addition of ammonia to spike the pH of Jerusalem artichoke juice between 6.5-6.7 before the solventogenic phase gave maximum ABE

42

production of 23-24 g/L after 36h for the strain IFP 904. The same effect was observed for strain IFP 902 (Jones & Woods, 1986; Marchal et al., 1985).

Although inulin has been known for more than a century, studies using it as a feedstock for butanol production have not been looked at in detail. Details about substrate preference of Clostridia, testing of various species and strains, and optimum parameters of the fermentation process have not been explored. However, there has been considerable research on the production of ethanol from hydrolyzed inulin (Allias et al.,

1987; Bonciu et al., 2010; Duvnjak et al., 1981; Ge & Zhang, 2005; Negro et al., 2006;

Ohta et al., 1993; Onsoy et al., 2007; Razmovski et al., 2011; Thanonkeo et al., 2011;

Toran-Diaz et al., 1985; Yuan et al., 2008; Yuan et al., 2012; T. Zhang et al., 2010). As stated earlier, substrates which have been tested for ethanol production can always be effectively used for butanol production as well.

2.4 Classification of fructans

Fructans are acid-labile, water-soluble, primary reserved carbohydrates synthesized in the plant vacuole in 15% of higher plants and also present in wide range of bacteria and fungi (Valluru & Van den Ende, 2008). The term fructan refers to linear or branched fructose polymers with β 2→1 and/or β 2→6 fructosyl-fructose linkages i.e. fructans include inulin, levan and graminan. Fructans can have at least one glucose unit with α (1-2) linkage, which is typically the starting link in the polymer chain, connected to long chain of fructose polymer. An individual fructan having a glucose molecule preceding fructose is designated as GFn, where G refers to the terminal glucose unit, F refers to the fructose units and n depicts the number of fructose units found in the fructan

43

chain. For example, GF2 is a fructan oligomer with a terminal glucose unit followed by two fructose units (Figure 2.8). A fructan with no glucose unit is designated as Fn

(literature also uses Fm), where n represents the number of fructose units recur in the fructan. For example, F3 represents an oligomer of three fructose units with two fructosyl-fructose linkages (Figure 2.9).

2.4.1 Fructan Biosynthesis and its chemical structure

Fructans are formed from a starting molecule of sucrose which explains the presence of single glucose unit in the polymer (Wack & Blaschek, 2006). Depending on the linkage type between the fructosyl residues and the position of the glucose residue several fructan types can be distinguished. Inulin has β 2→1 glycosidic linkage, levan is linked by β 2→6 linkage and both are linear fructans whereas graminan is a branched type fructans which has both β 2→1 and β 2→6 linkages (Schroeven et al., 2009; Valluru

& Van den Ende, 2008; Van den Ende et al., 2003; Van den Ende et al., 2004).

Inulin type fructan accumulates as long term reserve carbohydrates in underground storage organs such as roots and tubers. Graminan, levan and neokestose- derived fructans mainly act as a short-term energy storage compounds in stems, tiller bases, leaf sheaths, elongating leaf bases and to a small extent in leaf blades and roots

(Valluru & Van den Ende, 2008). Inulin is predominant in dicotyledonous species

(especially compositae); levan type fructan is mostly found in bacteria and monocotyledon plants; graminan type fructan is widely found in gramineae family

(wheat, barley, cereals and temperate fodder grasses); Neo-inulin and neo-levan types are found in monocotyledons such as Lolium, Apasaragus and Allium (Lasseur et al., 2006;

44

Lasseur et al., 2009; Van den Ende et al., 2003; Van den Ende et al., 2004; Van den Ende

& Valluru, 2009).

The diversity of fructans in plants can readily be explained by the different types of fructan biosynthetic enzymes (also termed fructosyltrasnferases [FTs]) involved in their synthesis. Two distinct categories of enzymes have been characterized that use different donor substrates to build the fructan. They can be classified as S-type FTs

(using sucrose as a donor substrate) and F-type FTs (using fructans as donor substrate)

(Lasseur et al., 2006; Schroeven et al., 2009; Tamura et al., 2009; Van den Ende et al.,

2011). Inulin-type fructan is the most common and well-studied fructan till date.

The enzymes involved in the synthesis of inulin are 1-SST and 1-FFT. At first, the enzyme sucrose-sucrose 1-fructosyltransferase (1-SST; EC 2.4.1.99) catalyzes the production of the trisaccharide 1-kestose (1K, 1-Kestotriose) by transferring a fructose unit via β 2→1 linkage to the sucrose molecule from another sucrose molecule.

Thereafter, the enzyme Fructan:fructan-1-fructosyltransferases (1-FFT; EC 2.4.1.100) catalyzes the elongation of fructose units on 1-kestose via a β 2→1 linkage, which generally results in beginning of inulin polymer. It was found that in dicots, 1-kestose acts as a preferential donor substrate for the elongation enzyme 1-FFT, producing inulin- type fructans (Edelman and Jefford, 1968; Van den Ende and Van Laere, 2007) with higher degrees of polymerization. Therefore, 1-FFT is able to synthesize fructan molecules with a DP of above three. From a structural/polymeric viewpoint, linear inulin can be considered as a polyoxyethylene backbone to which fructose moieties are attached, as are the steps of a spiral staircase.

45

1-FFTs are considered as important enzymes in dicots and the differences in inulin pattern between different species of Compositae is due to the differences in the affinity of 1-FFT for their acceptor substrates. In high DP inulin, the 1-FFTs favor longer inulin chain as the acceptor whereas 1-FFTs in low DP inulin prefer shorter chain. Globe thistle (Echinops ritro), Viguiera discolor Baker and artichoke (Cynara scolymus) have high DP-inulin; Chicory (Chicorum intybus) and Jerusalem artichoke (Helianthus tuberosus) have low DP inulin (Schroeven et al., 2009).

A more complex cocktail of FTs can be involved in the monocots (Ende et al.,

2011) depending upon the species. In cereals, the key enzyme Sucrose:fructan 6- fructosyltransferases (6-SFT) uses sucrose as a donor substrate and primarily transfers a fructose unit from sucrose to 1-kestose (acceptor) to synthesize the tetrasaccharide bifurcose (1&6-kestotetraose). The structure of 1, 6-kestotetraose has both 1-kestose and

6-kestose in its arrangement (Figure 2.10). 6-kestose can be used as an acceptor and it can be further elongated by adding fructose units via β 2→1 linkage into linear levan by enzyme 6-SFT. Branched fructans (graminans) are usually formed by the combined action of enzyme 6-SFT and 1-FFT. Another trisaccharide neokestose is biosynthesized by fructan; fructan 6G-fructosyltransferase (6G-FFT) from 1-kestose which is used as a donor substrate. 6G-FFT catalyzes the transfer of a fructose unit from 1-kestose to the C6 carbon of the glucose unit of sucrose, which is an acceptor, neokestose is produced.

Further elongation occurs by the action of a putative 6-SFT (neokestose levan series) or1-

FFT (neokestose inulin series). The enzymes involved and the respective fructans synthesized are schematically represented in Figure 2.11. (Edelman & Jefford, 1968;

Jerry Chatterton & Harrison, 2003; Joudi et al., 2012; Lasseur et al., 2009; Portes et al., 46

2008; Ritsema & Smeekens, 2003; Ritsema et al., 2005; Sprenger et al., 1995; Tamura et al., 2009; Valluru & Van den Ende, 2008; Van den Ende et al., 2000; Van den Ende et al., 2003; Van den Ende et al., 2004). All fructosyltransferase enzymes are believed to have evolved from vacuolar-type invertases and this process occurred independently in monocots and dicots (Lasseur et al., 2009; Schroeven et al., 2009; Van den Ende et al.,

2011).

Fructans are synthesized as extracellular polysaccharides with large degree of polymerization in bacteria and fungi. Most of the bacterial fructans are synthesized by levansucrases (EC 2.4.1.10) that synthesize levan-type fructan polymers from sucrose predominantly with β (2→6) linkages (Banguela & Hernández, 2006; Vijn & Smeekens,

1999). Bacteria with β (2→1) linked inulin-type fructan occurrence are very rare and reported only in Streptococcus mutants (Ebisu, Kotani, & Misak, 1975; Wolff et al.,

2000), Lactobacillus reuteri 121 (Olivares-Illana et al., 2002) and Leuconostoc citreum

(Van Hijum et al., 2002).

The enzymes involved in synthesize of bacterial fructan are inulosucrases (EC

2.4.1.9) and levansucrases (EC 2.4.1.10), which help synthesizing high DP fructans of inulin and levan type from sucrose, respectively. Levan produced from bacteria can have much large degree of polymerization ranging from 10,000- 100,000 and highly branched

(Wack & Blaschek, 2006). On the other hand, mostly fungi synthesize fructooligosaccharides (FOS) of DP 3-10 with linear β (2→1) linked fructans (Banguela

& Hernández, 2006). Both inulosucrases and levansucrases can transfer fructosyl units from the donor substrate sucrose to a variety of acceptor substrates such as water (sucrose

47

hydrolysis), sucrose (kestose synthesis), fructan (fructan polymerization), glucose

(sucrose synthesis) and fructose (bifructose synthesis).

The generation of high DP fructan or FOS between different bacterial species is highly dependent on specific enzyme activity, affinity for sucrose or other fructosyl acceptors, and the ratio of hydrolysis versus transfructosylation activities of both levansucrases and inulosucrases (Banguela & Hernández, 2006). The reason for longer chains of inulin/levan production by bacteria is that the longest chains are the most preferable acceptors of fructose units, along with the starting sucrose, which helps building longer chains. But in the case of plant inulin, 1-kestose and other short oligomers are also used as a substrate for chain elongation which causes the shorter inulin chain length from plants (Barclay et al., 2010).

Fructan depolymerization in plants is naturally accomplished by the concerted action of 1-fructan exohydrolase (1-FEH) or inulinase which cleaves the terminal fructose residue via β 2→1 linkages and by fructan 6-exohydrolase or levanase that hydrolyze the β2→6 linkages and releases terminal fructose monomers. In graminans, activities from both specific enzymes are needed for complete hydrolysis of fructans.

Many plants lack the invertase activity necessary to breakdown the sucrose molecule and they generate one sucrose molecule as an end product (Joudi et al., 2012; Lasseur et al.,

2009; Livingston et al., 2009; Portes et al., 2008; Ritsema & Smeekens, 2003; Van den

Ende et al., 2003; Van den Ende et al., 2004). Fructan exohydrolase enzymes are believed to have evolved from cell wall-invertases or from an ancestral β-fructosidase type of enzyme, capable of degrading sucrose and fructans (Schroeven et al., 2009; Van den Ende et al., 2004; Van den Ende et al., 2011). 48

2.4.2 Role of fructans in plants

Fructans breakdown to supply energy when plants need them e.g. to support leaf growth after defoliation or for early spring growth. Fructan accumulation has several advantages over starch as a protectant in abiotic stress. Unlike starch, fructan has high water solubility, resistance to membrane-damaging crystallization at subzero temperatures and reportedly fructan synthesis functions even at low temperatures.

Fructans are found to play physiological roles other than carbohydrate storage, lowering sucrose concentration in the cell, thus preventing sugar-induced feedback inhibition of photosynthesis. Another function of fructans is to protect plants against water deficit caused by drought or extreme low temperatures, possibly by stabilization of membranes under such stressful conditions (Cairns, 2003; Jerry Chatterton & Harrison, 2003; Lasseur et al., 2009; Livingston et al., 2009; Portes et al., 2008; Van den Ende et al., 2011). The more detailed functions of fructans are reported by Livingston et al (2009).

2.4.3 Normal occurrence of fructan

Fructans, after starch, are the most abundant non-structural polysaccharides found naturally in a vast variety of plants and in some bacteria. Fructans are commonly present among 1,200 species of grasses, whereas 15 % of flowering plants, especially dicots, produce fructans in significant amounts. Fructan is present in several monocotyledons and dicotyledonous families including Liliaceae, Amaryllidaceae, Gramineae and

Compositae. Among them, they are largely spread within the Liliaceae (3500 species), and most frequently among the Compositae (also known as Asteraceae) (25000 species) eg: Chicory dahlia, Jerusalem artichoke, yacon, and Kazak dandelion (Franck & De

Leenheer, 2005). Bacterial fructans are mostly levan type and its primary functions in 49

bacteria are to provide protection and temporary energy. Pseudomonaceae,

Enterobacteriaceae, Streptococcaceae, Actinomycetes, Bacillaceae and Lactobacillaceae are known to produce levan (Banguela & Hernández, 2006; Kaur & Gupta, 2002; Lasseur et al., 2006; Vijn & Smeekens, 1999).

Inulin is mostly stored in bulbs, tubers and tuberous roots in the Liliaceae,

Amaryllidaceae and Compositae plant families. Extraction of inulin from these plants is favored for industrial applications because of the absence of interfering compounds and easy extraction processes (Franck, 2002; Kaur & Gupta, 2002). Inulin present in foods that are consumed by humans mostly belong to the Liliaceae (leek, garlic, onion and asparagus) and Compositae (Kaur & Gupta, 2002).

2.5 Inulin

Inulin is a polysaccharide made up of majorly linear chains (little percent of branched chains exists in some species) of fructose units mostly with, but not exclusively,

β 2→1 fructosyl-fructose linkages produced in plants. Both levan and inulin can be with or without terminal glucose moiety. Those fructans without a terminal glucose can be the result of internal rearrangements or depolymerization reactions in fructan metabolism

(Vijn & Smeekens, 1999). The average degree of polymerization (DP) of inulin varies from 2-100 or more fructose units depending on chain length and polydispersity, plant species and the stage of life cycle of the plant. Isolation of inulin was first reported by a

German scientist named Rose, when he extract a peculiar substance from the plant Inula helenium in 1804 and this substance later named as inulin by Thomson in 1818.

However, the structure of inulin was first revealed by a plant physiologist Julius Sachs in

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1864, he used microscope to detect the spherocrystals of inulin in Dahlia, Helianthus tuberosus and Inula helenium after precipitating inulin with ethanol (Franck & De

Leenheer, 2005).

2.5.1 Nomenclature

The term native inulin refers to inulin with a mixture of residual sugars such as glucose, fructose, sucrose and small oligosaccharides. These mixtures of sugars are extracted from fresh roots and didn’t undergo any separation or fractionation to remove the oligomers and monomers. The reason for the presence of monomers and oligomers in the extract is because of inherent inulinase activity present in the plant as well as acid hydrolysis due to change in pH in plant physiology. This native inulin differs from commercial inulin, where they remove short oligomers and monomers that come with the plant inulin; resulting in high average DP for a particular plant inulin i.e. commercial inulin does not represent actual inulin DP range of plants (Franck & De Leenheer, 2005;

Wouters, 2009). The other three common terms which have often been used in literature are inulin, oligofructose and FOS. In broader sense these three terms represent the difference in DP of inulin compounds, however, the appropriate use of the terms appear highly inconsistent. Kelly, 2008 in his review paper has summarized the nomenclature of inulin that has widely been accepted and used in literature (Table 2.7).

The other generic terms used to represent inulin are high molecular weight inulin or long chain inulin (DP ≥10) and low molecular weight inulin or short chain inulin (DP

< 10). In some cases, DP of less than 10 can further be subdivided into short-chain (DP of 2-4) and medium chain (DP of 5-9) (Kelly, 2008).

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2.5.2 Physicochemical properties of Inulin

Commercially available inulin powder appears white in color, odorless, bland neutral taste, without any off-flavor or aftertaste. Standard chicory inulin has mild sweetness, about 10% of the sweetness of sucrose. The slight sweetness is attributed to the presence of short oligosaccharides along with the long-chain inulin, which is not sweet at all. Inulin is moderately soluble in water at room temperature (10% soluble), but it is highly soluble in water between 50°C to 100°C. When inulin is mixed thoroughly mixed with water, it forms a white creamy gel-like structure with a short spreadable texture. This unique property of inulin makes it possible to use it as bulking agent in foods to improve body and mouthfeel. Also, addition of inulin in foods exhibits humectant properties; reduced water activity ensures high microbiological stability, and affects boiling and freezing points. In general, short chain oligofructose is much more soluble in water (80% in water at room temperature) and has more sweetness (35 % in comparison with sucrose) than long chain inulin. So, oligofructose is often used in foods with other intense sweeteners for well-sustained flavor and improved stability. The β

(2→1) linkage is very labile under very acidic conditions; Inulin hydrolysis into fructose is more pronounced when inulin undergoes low pH, high temperature and low dry substance conditions. Commercially available inulin is predominantly produced from chicory. The physicochemical properties of chicory inulin are summarized in Table 2.8.

(Franck, 2002; Ranawana, 2008).

2.5.3 Applications and uses of inulin

Inulin is widely used in chemical, industrial, food and pharmaceutical applications. Inulin is chemically modified (neutral, anionic, and cationic modification as 52

well as cross-linking and slow release applications) to use as industrial reagents and biodegradable compound for many chemical and pharmaceutical applications (Stevens et al., 2001). Cross-linked inulin forms hydrogel with improved stability and this aspect is used to deliver drugs slowly targeting the colon to allow delayed absorption of drugs that have adverse effects in the stomach. Inulin is also being used a protective agent to improve stability of protein and peptide drugs in human therapeutics (Barclay et al.,

2010). Inulin, oligofructose, and FOS are classified as soluble fibers and it has been majorly used as functional food ingredients as a means of dietary fiber or to replace sugars or fats. Over the years, plenty of research investigated prebiotic effects of inulin and recent studies confirmed its significance in improving the overall gastrointestinal health.

Using inulin and oligofructose in dairy products, frozen desserts and meal replacers improves mouthfeel, taste and reduces syneresis of the product (Franck, 2002).

Average daily consumption of inulin in US and Europe is estimated to be between 1-4 g and 3-11 g respectively. Wheat, onion, banana, garlic and leek are the most common sources of inulin in human diet. Inulin and oligofructose consumption improves mineral absorption (calcium, copper and magnesium), modulate the composition of gut microflora, and reduces the risk of colon cancer.

Studies on effects of inulin supplementation to improve bone health and lipid metabolism remain inconclusive. It was found that shorter-chain inulin-type prebiotics produce adverse abdominal side effects than longer-chain inulin (Kaur & Gupta, 2002;

Kelly, 2008; Roberfroid, 2002).

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2.5.4 Inulin Hydrolysis

Hydrolysis cleaves the glycosidic bond of inulin into monosaccharides through addition of water. At room temperature and neutral pH, inulin does not undergo decomposition. However, low pH and increase in temperature boosted the rate of inulin hydrolysis (Barclay et al., 2010). The rate of hydrolysis of inulin is contingent upon the type and properties of glycosidic bond. It was found that the fructosyl-fructosyl bond is

4-5 times more susceptible to acid hydrolysis than the glucosyl-fructosyl bond.

Furthermore, internal molecules are more resistant than terminal fructose molecules to hydrolysis and this phenomenon is most likely due to the change of conformation being more easily achieved by the terminal group than the internal one. Regardless of these attributes of inulin, the rate of hydrolysis for short-chain oligomers is directly proportional to the concentration of inulin (up to 40%/w) and the hydrolysis follows first order or pseudo first order kinetics. Decomposition of hydrolyzed monosaccharides occurs at extreme temperature, pH or both. This monomeric decomposition can be avoided by using methanolysis instead of hydrolysis, since the methylated monosaccharides are more stable. Avoiding oxygen when hydrolysing with HCl or trifluroacetic acid was also found to prevent decomposition of the monosaccharides

(Barclay et al., 2010).

2.5.4.1 Thermal hydrolysis of inulin

Increase in temperature in the presence of water influenced hydrolysis of inulin.

Glibowski and Bukowska found that inulin chemical stability decreases at pH ≤4 with an increase in heating time and temperature above 60°C. In neutral and basic environments

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inulin remains chemically stable irrespective of heating time and temperature up to 100°C

(Glibowski & Bukowska, 2011; L'homme et al., 2003; Matusek et al., 2009). When inulin is thermally heated (dry heating) for 60 minutes at temperatures between 135°C to

195°C, degradation of inulin was in the range of 20 to 100%. Thermally-treated inulin produces di-D-fructose anhydrides (DFDAs) as degradation products, which is found to be inaccessible by enzymes (Böhm et al., 2005). Thermal hydrolysis of fructans from

Agave tequilana was carried out at 110-126°C and 1.2 kg/cm3 of pressure (Ávila-

Fernández et al., 2009). In another study, agave fructan gave 98% of fermentable sugars after 25.5 h of cooking, in which fructose represented more than 80% of the total carbohydrates obtained (Waleckx et al., 2008). However, thermal hydrolysis of fructan generates many volatile and undesirable Maillard compounds including furans, pyrans, aldehydes, nitrogen, sulfur, methyl-2-furate, and 5-hydroxymethyl furfural (Mancilla-

Margalli & López, 2002).

2.5.4.2 Acid hydrolysis of inulin

Inulin linkage is very labile under acidic conditions where the acids protonate the glycosidic oxygen and activate the leaving group. The hydrogen ion concentration affects the inulin hydrolysis kinetics in a first-order manner. Inulin hydrolysis follows the Arrhenius rate law for temperatures between 7-130°C and pHs from 2.0-4.2 (Barclay et al., 2010). Temperature, time and pH are the significant variables in sulphuric acid hydrolysis exhibiting maximum inulin hydrolysis at pH ≤ 2 and temperature of ~100°C after 1 hour (Eskandari Nasab et al., 2009; Szambelan & Nowak, 2006). Pekic et al

(1985) found that complete hydrolysis of Jerusalem artichoke inulin was achieved at pH

55

2 after 2.5 h at 100°C with minimum fructose dehydration. Fructose, a major hydrolysis product of inulin, is unstable under extreme acidic conditions and it can degrade further into 5-hydroxymethyl furfural. Unlike fructose, glucose is 40 times more stable in thermal acid conditions. Fructose forms an unstable ring structure in open chain form, which is largely susceptible to enolization, which is believed to be the cause for HMF formation upon degradation of fructose (Kuster, 2006). Use of milder acidic conditions minimized fructose degradation significantly (<10%) and did not affect the quality of the hydrolyzate. Various acid and temperature conditions were proposed to minimize fructose degradation within tolerable limits (E. Kim, 2007; Nguyen et al., 2009).

2.5.4.3 Base hydrolysis of inulin

Bases can hydrolyze inulin through the i.e. bases can hydrolyze only from the reducing end of an inulin chain and it cannot hydrolyze inulin which has a terminal glucose moiety (no reducing end available). This attribute generally hampers hydrolysis of commercial inulin since it has very little reducing sugars. If inulin is thermally treated with high temperature, it will cleave the non-reducing long inulin chains into smaller chains with reducing ends; thereafter application of base-induced hydrolysis is feasible (Barclay et al., 2010).

2.5.4.4 Enzymatic hydrolysis of inulin

Enzymatic hydrolysis of inulin is preferred to acid hydrolysis by industry because of the maximal sugar yield (final concentration of fructose up to 95%) and minimal degradation (Bekers et al., 2008). Furthermore, acid hydrolysis produces undesirable coloring and flavoring of inulin hydrolysis products, formation of tasteless di-fructose 56

anhydride, and increases in ash content, which is expensive to remove (Derycke &

Vandamme, 1984; Kango, 2008; Kochhar et al., 1999). One notable limiting factor of inulin is its low solubility in water. Leite et al reported that inulin is 6% soluble at 10°C and 35% at 90°C (Leite et al., 2004).

Inulinase (also known as inulase) is a hydrolytic enzyme capable of cleaving β

2→1 inulin linkages. Based on the activity, inulinase can be classified into two types namely exo-inulinases (β-D-fructan fructohydrolase, EC 3.2.1.80) and endo-inulinases

(2,1-β-D-fructan fructohydrolase, EC 3.2.1.7). Exoinulinase hydrolyzes the terminal fructose molecule from the inulin chain whereas endoinulinase hydrolyze the internal linkages of inulin into smaller oligosaccharides such as inulotriose, inulotetraose, and inulopentoses (Carniti et al., 2004; Z. Chi et al., 2009; Kango, 2008; Vijayaraghavan et al., 2009). Inulinase enzyme is naturally produced in plants such as dandelion, chicory and Jerusalem artichoke, that can be extracted and purified, however, enzymes derived from plant sources are found to be less efficient than microbial ones, which can be used for commercial applications (Kochhar et al., 1999; Ricca et al., 2007). Since inulinase can be produced from different sources, a vast array of literature is available in different areas such as saccharification of inulin by inulinase producing microbes (Workman &

Day, 1984), simultaneous saccharification and fermentation (Bekers et al., 2008; Ge &

Zhang, 2005; Ohta et al., 1993; Saha, 2006), using recombinant inulinase producing organisms (Y. H. Kim et al., 1998; Liu, et al., 2012; L. Zhang et al., 2003), and hydrolysis using commercial inulinase enzyme. These microbial inulinases from different sources distinguish themselves by enzyme activity, pH optimum and temperature optimum. 57

Substrate concentration plays a crucial role in deciding the rate of hydrolysis of inulin. The rate of hydrolysis is generally faster if the inulin is shorter chain, attributed to the nature of exo-inulinase which acts on one end of inulin molecule. Therefore, inulin with high degrees of polymerization results in less available reactive ends within the reacting mixture (Derycke & Vandamme, 1984; W. Y. Kim et al., 1982; Ricca et al.,

2007; Zittan, 1981). Iron, mercury, silver or manganese ions in the reaction mixture can inhibit inulinase activity (Ricca et al., 2007). Attempts to produce fructose from inulin on an industrial scale have brought many technological innovations and improvements in inulin hydrolysis. Studies focused on inulinase purification (Y. J. Cho & Yun, 2002;

Fawzi, 2011; Kalil, et al., 2010; Ohta et al., 2002; Pessoni et al., 2007; Skowronek &

Fiedurek, 2006), enzyme recycling, inulinase immobilization (Catana et al., 2007; B. W.

Kim et al., 1997; Nakamura et al., 1995; Fernandes et al., 2008; Ricca et al., 2010; Rocha et al., 2006; Uzunova et al., 2002); different operating procedures and bioreactor types

(Díaz et al., 2006; Hang et al., 2011; Maria., 2011; Ricca et al., 2010; Rikir et al., 1990;

Singh et al., 2008) have been attempted to increase enzyme activity, enzyme shelf-life, stability and kinetic properties. Also, factorial design and surface response methodology has been applied to determine optimal operating conditions for inulinase enzymes

(Dilipkumar et al., 2011; Kalil et al., 2006; Rocha et al., 2006; Sheng et al., 2009; Yuan et al., 2012).

The commercial enzyme, Fructozyme L (a product of Novozyme), is a mixture of

10% of endoinulinase from Aspergillus niger and 90% Exoinulinase from Bacillus stearothermophilus (Basso et al., 2009) with inulinase activity of 2000-2500 INU/g

(Michel-Cuello et al., 2012). This enzyme is the most studied and used commercial 58

enzyme for inulin hydrolysis. Waleckx et al. (2011) found that the optimum conditions for Fructozyme L to hydrolyze agave fructan were 60°C, pH of 4.0-5.0, enzyme concentration of 0.02% (V/V) to achieve more than 90% of hydrolysis after 12 h.

Fernandez et al (2009) observed significant drop in the hydrolysis rate of agave fructans after 30 min of the reaction as substrate concentration decreases. When 3.4% v/w of

Fructozyme L is used, total hydrolysis was achieved after 30 min whereas it took 3.5 h if the enzyme concentration was reduced to 1.7% v/w. Most literature cited Fructozyme L as the most thermostable commercial enzyme; it has an optimum temperature of 60°C and the enzyme is stable at room temperature after 6 h of exposure to 60°C. The optimum pH is around 4.5; however, the enzyme retains its stability and activity between pH 4.0-6.0 (Bekers et al., 2008; Michel-Cuello et al., 2012; Muñoz-Gutiérrez et al., 2009;

Rocha et al., 2006; Šimonová et al., 2010; Vendrell-Pascuas et al., 2000). Another commercial enzyme, that has recently been used to produce oligosaccharides, is the endoinulinase enzyme (Novozyme 960) isolated from Aspergillus niger. Mutanda et al. obtained a maximum oligofructose yield of 54% after 72 h of reaction with 5% inulin at

45°C with enzyme concentration of 5 U/g of substrate (Mutanda et al., 2008; Risso et al.,

2012). A few other authors have used endoinulinase from A. niger but with different substrate and enzyme concentrations (Nemukula, 2008; Nguyen et al., 2011; Ronkart et al., 2007; Szambelan & Nowak, 2006). Recently, several authors have proposed a mathematical model to predict the kinetics of process optimization, substrate degradation and fructose production. Ricca et al. developed a model by combining reaction kinetics model and a deactivation model of the reaction of inulin enzymatic hydrolysis to predict reaction performance for substrate concentrations between 10 and 40 g/L and reaction 59

temperature of 60°C. They found that this model can also be used to optimize temperature, reduce reaction times and minimize enzyme loading (Ricca et al., 2009;

Ricca et al., 2010). A mathematical model describing kinetics of substrate consumption and fructose production was put forth by Michel-Cuello et al. (2012). They noticed the reaction kinetics was significantly influenced by temperature, substrate concentration and type of substrate.

2.6 Production of exo- and endo-inulinases

Microbial inulinases are considered to be the best choice for commercial production of inulinases because of their easy cultivation and high enzyme yields (Z. Chi et al., 2009). Inulinase enzyme is produced by different microorganisms such as bacteria, fungi and yeasts with diverse enzyme activities. Inulinase producing microorganisms and their maximum enzyme yield are reported in Table 2.9. Among the several species, fungal strain belonging to Aspergillus sp. (filamentous fungi) and yeast strains belonging to Kluyveromycessp. (diploid yeast) are the most common, well-studied, versatile source of inulinase producing organisms which are preferred choice for large-scale inulinase production (Z. Chi et al., 2009; Helen Treichel et al., 2012; Neagu & Bahrim, 2011;

Pandey et al., 1999; Singh & Gill, 2006). Researchers have also found that yeast strains can produce more inulinase than fungal and bacterial strains (Z. Chi et al., 2009). High enzyme yields were found in various yeasts namely Pichia sp., Kluyveromycesmarxianus,

Kluyveromycesfragilis and Cryptococcus aureus (Z. Chi et al., 2009; Gong et al., 2007;

Sheng et al., 2008). Inulinase production from bacteria is least explored compared to fungi and yeasts, however, bacterial inulinases are known for their thermostability and

60

they secretes endoinulinases (Neagu & Bahrim, 2011; Ricca et al., 2007). Streptomyces sp. are found to be the good inulinase producing bacterial species; other species such as

Pseudomonas sp., Arthrobacter sp., and Bacillus sp., can also produce inulinase (Neagu

& Bahrim, 2011). In fungi, Aspergillus sp. are the most characterized and the best known inulinase producers. Among them, strains of A. niger have been extensively investigated and characterized over the years. Derycke & Vandamme observed both extra- and intra- cellular inulinases, which displayed maximum activity at pH 4.3-4.4 and temperature of

55-56°C, from an A. niger strain isolated from a compositae rhizosphere, (Derycke &

Vandamme, 1984). Studies carried out on A. ficuum revealed the existence of multiple isoforms of exoinulinases and endoinulinases enzymes that are glycoproteins with high sugar content (Ettalibi & Baratti, 1990; Jing et al., 2003). As Pandey et al summarized in their review paper, most of the inulinases produced from fungi have more than 50 kDa of molecular weight (Pandey et al., 1999). It is noteworthy that commercial endo-inulinase enzyme Novozym 230 (molecular weight of 55.8-64 kDa) is produced from A. ficuum and Novozym 960 (molecular weight of 55.9-64 kDa) is from A. niger29 (Singh & Gill,

2006).

As far as inulinase production from yeasts is concerned, K. fragilis and K. marxianus were looked in detail as potential inulinase producers (Singh & Gill, 2006).

Mutant yeast strains, such as of K. marxianus var. marxianus CBS 6556 and Candida pseudotropicalis IP513, exhibited 3000 U/ml and 25,000 U/g units, respectively, which is higher than the parental strains enzyme production. Endo- and exo- inulinase activities

29 www.brenda-enzymes.org

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were noticed in Kluyveromyces sp. strain Y-85 which produced intracellular enzyme at a molecular mass of 42 and 65 kDa and extracellular enzyme at 57 kDa (Ricca et al.,

2007). Several marine yeast strains were found to have the capability to produce large quantities of inulinase enzymes. Reportedly strains of Pichia guilliermondii,

Cryptococcusaureus, Yarrowia lipolitica and Debaryomyces hansenii can produce exoinulinase above 40 U/ml (Z. Chi et al., 2009; L. Gao et al., 2007; Gong et al., 2007;

Gong et al., 2008). The enzyme yield, location of enzyme and nature of enzyme activity are determined by the choice of microorganism and the substrate used during fermentation (Ricca et al., 2007).

2.6.1 Optimum pH and temperature for inulinase production

Optimum pH and temperature values for different inulinase-producing microorganisms were often reported in many literatures. Pandey et al, stated that fungal inulinases exhibited an optimum pH between 4.5 and 7.0, yeast inulinases between 4.4 and 6.5 and bacterial inulinases between 4.8 and 7.0. Optimal temperature values reported for fungi generally are less than for bacteria and yeasts (Pandey et al., 1999;

Ricca et al., 2007). Most literature quoted that the optimum pH of fungi and yeasts are in the range of 4.5-6.0 (Z. Chi et al., 2009; Gong et al., 2008; Neagu & Bahrim, 2011; Singh et al., 2007). Inulinase produced from bacteria are found to be naturally slightly alkaline than inulinase from yeasts and fungi. Some bacterial strains such as Arthrobacter sp.

(Kang et al., 1998); Bacillus polymyxa (Z. Chi et al., 2009; Zherebtsov et al., 2002) and

Bacillus smithii (W. Gao et al., 2009), show maximum inulinase activity between pH of

7-7.5. Li et al. isolated a new bacterial strain from marine sediments and named it

Marinimicrobium sp. LS-A18, which showed maximum enzyme activity at pH 9.0 and 62

temperature 55°C (Li et al., 2012). Optimum temperatures for attaining maximum inulinase activities are significantly different for different species. Reportedly, inulinase activities for fungi and bacteria were higher between temperatures of 50-55°C (Z. Chi et al., 2009; Kang et al., 1998; Nakamura et al., 1997). Industrial processes involved in commercial production of fructose or fructooligosaccharides require higher operating temperatures which might be challenging for inulinases with lower optimum temperatures since at higher temperatures the enzyme will lose its activity due to thermal deactivation. To counteract this problem, inulinases with higher optimum temperatures are preferred to ensure proper solubility of inulin and also to prevent microbial contamination. Furthermore, higher temperatures will also bring down the cost of production because a lower amount of enzyme is employed to produce the desired product (W. Gao et al., 2009; Ricca et al., 2007). Both pH and temperature are critical in maintaining enzyme activity and stability in a process.

2.6.2 Factors affecting inulinase production

Agitation and aeration are considered to be effective in enhancing inulinase production in some microorganisms. Singh & Bhermi found that the agitation mode of cultivation increase inulinase synthesis in K. marxianus YS-1 up to 37 U/ml at 150 rpm whereas in stationary mode it produced 10.6 U/ml. The increase in inulinase production is attributed to the uniform distribution of the microbial culture in the medium with a better nutrient availability and oxygen transfer. Experiments at higher agitation showed reduced enzyme activity 23.1 U/ml. Therefore, the higher agitation rate is counter- productive because of shear stress on microbial cells and on the enzyme structure as well

(R. Singh & Bhermi, 2008). Agitation rate of 150 rpm has widely been reported as the 63

optimum speed for inulinase production from K. marxianus at shake flask level

(Selvakumar & Pandey, 1999; Silva-Santisteban, 2005; Singh & Gill, 2006; Singh et al.,

2007; Singh & Bhermi, 2008; Vranešic et al., 2002). Agitation mode of fermentation at

180 rpm for K. marxianus has also been reported (Cazetta et al., 2005). Optimization of aeration rate for inulinase production have been found to influence inulinase production

(Cazetta et al., 2010; Dinarvand et al., 2012; Silva-Santisteban, 2005; Singh et al., 2007;

Singh & Bhermi, 2008; Wei et al., 1998). Addition of mineral ions to the reaction medium had mixed effects on inulinase production, with some ions boosting inulinase production while some severely inhibited enzyme production. The effects of metal ions on microorganisms are not common and it appears the requirements of metal ions are specific to the microorganism used. For the yeast, K. marxianus, Singh & Bhermi found that inulinase production is activated by CO2+, Mn2+, Mg2+ and is inactivated by Cu2+,

Fe3+, Zn2+, Ca2+, Ba2+, Zn2+ (Kushi et al., 2000; Neagu & Bahrim, 2011; Singh & Bhermi,

2008). Some protein compounds such as pepstatin, EDTA, 1,10-phenenthroline have inhibitory effects on inulinase activity indicating that the characterized enzymes were metalloenzymes (Gong et al., 2008; Kang et al., 1998; Sheng et al., 2008). Several other factors such as surfactants, age and size of inoculum, nitrogen and carbon source, and substrate concentration were also found to have effects on inulinase production

(Dinarvand et al., 2012; Singh et al., 2006; Singh et al., 2007; Singh & Bhermi, 2008).

2.6.3 Substrates for inulinase production

Inulin is the most widely used substrate in inulinase production from different microorganisms, however, a variety of substrates have been used for enzyme production over the years. They comprise pure substrates (mono-, di-, or polysaccharide sugars), 64

naturally existing inulin-rich materials and mixed substrates. The general criterion in choosing the appropriate substrate is contingent upon the nature of inulinase enzyme activities shown by the microorganisms. If the microorganism exhibited only inulinase activity, then inulin serves as the best substrate. But, if the microorganism demonstrated inulinase activity coupled with invertase activity, sucrose functions as a better source for enzyme production (Pandey et al., 1999).

Naturally-occurring inulin-rich substrates such as aqueous extracts of chicory roots (Fawzi, 2011; Gupta et al., 1988; Park & Yun, 2001), dahlia tubers (Cruz et al.,

1998; Jain et al., 2012; Singh et al., 2007), dandelion roots (Kango, 2008), Jerusalem artichoke (Erdal at al., 2011; Ertan et al., 2003; Mohammed S. El-Hersh et al., 2011;

Saber & El-Naggar, 2009; Sirisansaneeyakul et al., 2007), yacon (Cazetta et al., 2005);

Asparagus officinalis tubers (Singh & Bhermi, 2008) and onion and garlic peels

(Ayyachamy et al., 2007; Dilipkumar et al., 2011; Mahmoud et al., 2011) were also used us substrate for inulinase production. More extensive reviews of inulinase production are reported elsewhere (Z. Chi et al., 2009; Helen Treichel et al., 2012; Neagu & Bahrim,

2011; Pandey et al., 1999; Ricca et al., 2007; Singh & Gill, 2006; Vijayaraghavan et al.,

2009).

2.7 Inulin characterization and estimation

Carbohydrates can adopt complex branched structures with individual monomeric units linked at one of several sites, which contribute to the chemical heterogeneity and structural complexities of carbohydrates (Harvey, 2003; Z. Zhang & Linhardt, 2009).

Because of the diversity of fructans, characterization and estimation of them was highly

65

challenging by using conventional chromatographic techniques such as such as paper or thin-layer chromatography or gel permeation or adsorption chromatography. The major drawbacks in employing these techniques are: 1) resolution tends to decrease with increasing analyte size and these fructans having DP more than 10 are often poorly resolved, 2) accurate mass determinations require standard compounds closer to the molecules analyzed, 3) require extensive additional methodology includes acid and various enzymatic and colorimetric procedures to identify the compounds (Stahl et al.,

1997).

High performance liquid chromatography (HPLC) offers high resolution, fast analysis, direct injection of the sample without or with little pretreatment, and easy of automation. Standards HPLC techniques using columns specifically designed for carbohydrate analysis and separating low molecular weight oligomers of inulin up to a maximum of about DP 16 are reported in literatures. Columns with different chemical functionalities separation mechanisms such as ion exclusion, ion and ligand exchange, reversed phase, normal phase and size exclusion have been employed (Barclay et al.,

2010). Since inulin has, conformationally, a flexible backbone, it causes problems in columns meant for normal carbohydrates. Almost all forms of HPLC can separate only oligomeric forms of inulin up to certain DP and longer polymer chains above that range elute as a single peak. Therefore, standard forms of HPLC are inadequate for complete inulin separation. Moreover, sensitive detection of carbohydrates after HPLC separation represents additional challenges because the carbohydrates lack chromophores and flurophores. This disadvantage may be offset by employing low wavelength UV or refractive index detection. UV detection is not suitable for underivatized carbohydrates 66

that uses organic mobile phase, since they also absorb UV lights. Though refractive index detection is widely known, its ability is severely limited where gradient elution is required. Evaporative light-scattering detection (ELSD) is insensitive to temperature fluctuations and its estimation depends on mass of the vaporized analyte (Corradini et al.,

2012).

A major breakthrough in the HPLC-analysis of oligosaccharides was the introduction of high performance anion exchange chromatography (HPEAC) using high pH eluents in combination with pulsed amperometric detection (PAD) (Schols et al.,

2000). HPEAC-PAD has the ability to separate all classes of alditols, aminosugars, mono-, oligo- and polysaccharides according to their structural features such as size, composition, anomericity, and linkage isomerism (Corradini et al., 2012). However, major issues of HPAEC PAD are lack of sensitivity for long chain polymers and inability to quantify each compound due to lack of commercial standards. Therefore, it can only provide qualitative results and not quantitative ones. It requires laborious isolations and characterization techniques to identify the unknown compounds (Barclay et al., 2010;

Borromei et al., 2009; Harrison et al., 2012).

HPAEC with pulsed electrochemical detection (PED) with gradient elution has also been looked as a viable instrument that allows mixtures of simple sugars, oligo and polysaccharides to be separated in a single run (Corradini et al., 2012). However, the major disadvantage in this approach is that peak identity is solely based on retention time and it is highly prone to misidentification due to retention time shifts (Harrison et al.,

2011; Harrison et al., 2012).

67

In the last two decades two soft ionization methods such as matrix assisted laser desorption/ionization (MALDI) and electrospray ionization (ESI) has gained much interest in molecular sizing and characterization of carbohydrates. Among them, MALDI coupled to time-of-flight (TOF) has extensively been used and demonstrated as a powerful tool for carbohydrates (Mischnick, 2012; Štikarovská & Chmelı́k, 2004).

MALDI is a method that uses high energy laser to vaporize and ionize non-volatile biological samples from a solid state phase directly into the gas phase. The sample is dissolved in a matrix which helps in absorbing and dissipating the energy provided by the laser and causing the substrate to vaporize. The resulting ions are passed on to a mass spectrometer and separated based on their mass and charge (m/z ratio). MALDI-TOF comes with two basic modes based on the path taken by the ions to the detector, i.e. linear

(LIN) and reflectron (REF) mode. Laser power influences the degree of desorption and ionization in MALDI-TOF analysis. Usually with increase in laser strength, more ions are generated, but higher laser power can also lead to fragmentation (Štikarovská &

Chmelı́k, 2004). It was reported that MALDI-MS can detect up to DP70 (Molecular mass of 11,358) (Mischnick, 2012). Electrospray ionization (ESI-MS) technique is another well-known method in which ions carry multiple charges and reduces their mass- to-charge ratio. This helps in getting mass spectra for large molecules. Mischnik reported that ESI-MS is superior to MALDI-MS in offering sequence analysis and structure information of a compound (Mischnik, 2012).

In MALDI-MS analysis, neutral carbohydrates mainly form sodium [M+Na]+ (in mass spectrometry M+ represents the molecular ion) and [M+K]+ adducts mainly with protonated molecules as minor products (Fabrik et al., 2012). These adducts can interfere 68

with peaks of low molecular weight mass analytes and thus make multiple artificial peaks. These cluster of speaks originate by combinations of molecules of matrix and/or matrix without water and cations of potassium and/or sodium. In another sense, these clusters can be considered as positive contribution for MALDI-TOF analysis, because their peaks can be used as internal calibrants (Štikarovská & Chmelı́k, 2004).

Tandem mass spectrometry technique (MALDI-TOF or ESI coupling with HPLC) can provide molecular mass profiling, sugar constituents, sequence and interresidue linkage positions and more information on stereochemistry of compounds. Coupling

HPAEC-PAD with mass spectrometry for online detection has limited application because of long run times, problems in quantifying individual compounds due to different response factors of the PAD detector, additional purification by precolumn chromatography, and lack of definitive peak identification due to difficulties in coupling to mass spectrometry.

Advancement in analyzing underivatized oligosaccharides using porous graphitized carbon high pressure liquid chromatography columns (PGC- HPLC) that couple to MS analysis was reported to be highly sensible, fast, reproducible and able to measure low DP molecules. PGC-HPLC coupled to negative electrospray ionization mass spectrometry with multiple charge state ions has been used to separate and quantify fructan isomers between DP3 to DP49 (Harrison et al., 2009). Improvements in fructan detection (DP up to 100) was noticed when PGC-HPLC was used with Exactive orbitrap

MS (Harrison et al., 2011).

Gas chromatography (GC) has also been attempted to estimate the molecular weight of inulin and its application was limited (can measure DP of only up to 9-10) due 69

to the volatilization of extracted sugars by silylation. Apolar columns capable of being heated to 440°C are required to analyze short chain oligomers. Longer polysaccharides require hydrolysis followed by end group analysis, while linkage and branching can be determined by permethylation, followed by reductive cleavage of the polymer and acetylation. Because of the complexity involved and relatively poor ability to measure longer chain molecules, this technique is least preferred (Barclay et al., 2010).

Estimation of chemical composition, choice of suitable substrate and strain, optimum fermentation conditions and results of food processing wastes are described in

Chapter 3.

70

Properties n-Butanol Gasoline Ethanol Diesel Methanol Biodiesel*

Molecular formula C4H9OH C4 – C12 C2H5OH C12 – C25 CH3OH C12-C22 Boiling point (°C) 117.7 25-225 78.4 180-370 64.5 182-338 Density at 20°C (g/ml) 0.8098 0.7-0.8 0.7851 0.82-0.86 0.7966 0.86-0.89 Solubility in 100 g of water immiscible immiscible miscible immiscible miscible immiscible Energy density (MJ.l-1) 27-29.2 32 19.6 35.86 16 32.6 Auto-ignition temperature (°C) 385 ~300 434 ~210 470 177 Heat of vaporization (MJ/kg) 0.43 0.36 0.92 0.27 1.2 -

Specific heat capacity Cp at 20°C (kJ/kg.K) 2.40 2.22 2.47 1.75 2.54 -

71 Flash point (°C) at closed cup 35 -45 to -38 8 65-88 12 100-170 Cetane number 25 0-10 8 40-55 3 48-65 Research octane number 96 91-99 129 - 136 - Motor octane number 78 81-89 102 - 104 -

Octanol/Water Partition Coefficient (as logPo/w) 0.88 3.52±0.62 -0.31 ~3.3 -0.77 - Stoichiometric Air/Fuel ratio (wt./wt.) 11.21 14.7 9.02 14.3 6.49 13.8 Latent heating (kJ/kg) at 25°C 582 380-500 904 270 1109 - Flammability limits (%vol.) 1.4-11.2 0.6-8 4.3-19 1.5-7.6 6.0-36.5 - Saturation pressure (kPa) at 38°C 2.27 31.01 13.8 1.86 31.69 - Viscosity (mm2/s) at 40° C 2.63 0.4-0.8 (20°C) 1.08 1.9-4.1 0.59 1.9-6.0 *ASTM D6751

Table 2.1 Physical and chemical properties of n-butanol compared with other fuels (References: Gholizadeh, 2010; C.Jin et al., 2011; Rakopoulos et al., 2010;Joshi & Pegg, 2007, NREL 2009 report)

Properties n-Butanol Chemical Structure Melting point (°C) -89.3 Ignition temperature (°C) 35-37 Auto-ignition temperature (°C) 343-345 Flash point (°C) 25-29 Specific gravity 0.810-0.812 Critical pressure (hPa) 48.4 Critical temperature (°C) 287 Oxygen, %wt 21.6 Water solubility 9.0 ml/100ml (7.7g/100ml at 20°C) Relative vapor density (air:1.0) 2.6 Vapor pressure (kPa at 20°C) 0.58 72 Properties 1-butanol 2-butanol Tert-butanol Iso-butanol Density (kg/m3) 809.8 806.3 788.7 801.8 Research octane number 96 101 105 113 Motor octane number 78 32 89 94 Boiling temperature (◦C) 117.7 99.5 82.4 108

Enthalpy of vaporization (kJ/kg) at Tboil 582 551 527 566 Self-ignition temperature (◦C) 343 406.1 477.8 415.6 Flammability limits vol.% 1.4–11.2 1.7–9.8 2.4–8 1.2–10.9 Viscosity (mPa s) at 25 ◦C 2.544 3.096 – 4.312

Table 2.2 Comparison of properties of n-butanol with its isomers (References: Gholizadeh, 2010; C. Jin et al., 2011; Wallner, Miers, & McConnell, 2009)

Butanol isomers Molecular structure Major applications

1 - Butanol v Solvents for paints, resins, dyes, etc. Plasticizers, chemical intermediate – for butyl esters or butyl ethers, etc. Cosmetics including eye makeup, lipsticks, etc. Gasoline additive

Solvent 2 - Butanol Chemical intermediate – for butanone, etc. Industrial cleaners – paint removers Perfumes or in artificial flavors

Solvent and additive for paint iso-Butanol Gasoline additive

73 Industrial cleaners – paint removers Ink ingredient

Solvent tert-Butanol Denaturant for ethanol Industrial cleaners – paint removers Gasoline additive for octane booster and oxygenate Intermediate for MTBE, ETBE, TBHP, etc.

Table 2.3 Major applications of butanol isomers (Reference : C. Jin et al., 2011)

Biomass composition (%wt) Guayule bagasse Guayule leaf stream biomass* Cellulose 21 ± 3 30.9 ± 0.9 Hemicellulose 13 ± 1 29 ± 2 Acid insolubles 54 ± 2 37.3 ± 0.2 Acid soluble lignin 0.7 ± 0.1 0.52 ± 0.02 Others 11 ± 3 3 ± 3

Table 2.4 Composition of guayule biomass (Reference: Srinivasan & Ju, 2010).*Leaf stream biomass analyzed is without any prior extraction

Components (% dw) Whole shrub Guayule bagasse C 47.09 55.19 H 5.23 6.35 N 1.38 0.65 S 0.62 0.20 O 32.88 34.36 HHV (kJ/kg) 18,329 22,385

Table 2.5 Elemental composition of guayule biomass (Reference: Boateng et al., 2009)

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Type of Waste description Type of processing End product food industry Stems, stalks, leaves, rotten  Composting (Anaerobic treatment.)  Bio-fertilizer. Bio-gas & Fertilizer fruit and vegetables  Landfill.  Fertilizer  Fermentation  Biobutanol; Bioethanol, Biogas Fruit Seeds, pulp, peel,  Extraction of pectin.  Pectin for jams, etc. & contaminated or unfinished  Extraction of oleo-resins  Various food ingredients vegetable products, etc. antioxidants,colors, essential oils,  Biobutanol, Bioethanol, Biogas, processing (post processing) Enzymes vinegar, residual dietary fibers, etc.  Fermentation of residual sugars & 75 carbohydrates.  Skins & hides, blood  Salting, tanning, valorization  Leather, dresses, food ingredients, Meat  Offals  Cleaning & processing animal feeds, Industrial & pharma &  Bones  Crushing & processing compounds, fertilizers poultry  Fat  Fat extraction; Fermentation;  Sausage casings, animal feeds, fertilizers. processing  Leftover meats after Transesterification  Bone meal, mineral supplements, gelatin. prime  Cleaning, mincing; Fermentation  Cooking fats, Soaps, Biodiesel, biogas cuts  Minced Meats, sausages, pet foods biogas etc.

Table 2.6 Types of food processing wastes and its modes of utilization

Table 2.6 continued

Type of Waste description Type of processing End product food industry  Skins, bones, heads, leftover  Size reduction & steam cooking; Oil  Fish meal & fish protein meat after filleting extraction & refining; Hydrolyzing; concentrate; Fish Oil, Fish Protein Sea food  Skins & bones Composting; Transesterification hydrolyzate; Biofertilizer; Biodiesel processing  Shells, legs, etc. of  Cold water extraction, Hot water  Collagen; Gelatin crustaceans extraction  Chitins & chitosans  Pulverizing & Processing

76  Whey from cheese making;  Precipitation, concentration; Cleaning  Whey powder, whey proteins, Cheese granules & processing; Fermentation lactose Cheese preparations, Dairy  Fat  Centrifugation, dehydration animal feeds; Butter oil (ghee) Industry  Waste fat; Buttermilk  Tray drying; Transesterification  Soap  Buttermilk powder; Biodiesel

(Adapted from: Unit 17. Waste Management in Food Processing Industry) http://vedyadhara.ignou.ac.in/wiki/images/a/ad/MVP-001-Block_4-Unit-17.pdf

FOS Short-chain, inulin type fructan mixes synthesized from sucrose Oligofructose Inulin-type fructan mixes with DPmax <10 that is produced by partial hydrolysis of inulin and then undergone physical separation to remove all long chain (DP ≥10) inulin. Inulin Hot water extracts that has inulin fructans and not undergone further processing Inulin HP Exclusively long-chain high-molecular weight inulin-type fructans (inulin with DP < 10 physically removed). FOS-enriched inulin Proprietary mixes that enrich inulin with FOS FOS-enriched inulin HP Proprietary mixes that enrich inulin HP with FOS Oligofructose-enriched inulin Proprietary mixes that enrich inulin with oligofructose Oligofructose-enriched inulin HP Proprietary mixes that enrich inulin HP with oligofructose

Table 2.7 General nomenclature used in Inulin studies

77

Physicochemical Properties Standard inulin High performance inulin Oligofructose powder Chemical structure GFn (2 ≤ n ≤ 60) GFn (10 ≤n ≤ 60) GFn + Fn (2 ≤ n ≤ 7) Average degree of polymerisation 12 25 4 Dry matter (%) 95 95 95 Inulin/oligofructose content (% on d.m.) 92 99·5 95 Sugars content (% on d.m.) 8 S 0·5 5 PH (10% w/w) 5–7 5–7 5–7 Sulphated ash (% on d.m.) < 0·2 < 0·2 < 0·2 Heavy metals (ppm on d.m.) < 0·2 , <0·2 <0·2 Appearance White powder White powder White powder Taste Neutral Neutral Moderately sweet 78 Sweetness (v. sucrose = 100 %) 10% None 35% Solubility in water at 258C (g/l) 120 25 >750 Viscosity in water (5 %) at 108C (mPa.s) 1·6 2·4 <1·0 Functionality in foods Fat replacer Fat replacer Sugar replacer Synergism Synergy with gelling Synergy with gelling agents Synergy with intense agents sweeteners

Table 2.8 Physicochemical properties of inulin Adapted from Franck (2002).

Micro-organism Maximum Enzyme Reference yield [U/ml] FUNGI Aspergillus sp. 75 Pandey et al., 1999 A. aureus MTCC 151 160 Pandey et al., 1999 A. ficuum 3000∗ Pandey et al., 1999 A. fischeri MTCC 150 1–1.2 Pandey et al., 1999 A. flavus MTCC 277 1–1.2 Pandey et al., 1999 A. nidulans MTCC 344 1–1.2 Pandey et al., 1999 A.niger 100 Ge and Zhang, 2005 52.5 Kango, 2008 176 Kumar et al., 2005 60 Poorna and Kulkarni, 1995 A. niger 817 0.0685 Pandey et al., 1999 A. niger A42 4600∗ Pandey et al., 1999 A. niger MTCC 281 1–1.2 Pandey et al., 1999 A. niger mutant 817 160 Pandey et al., 1999 A. niger mutant selection 35.18 Skowronek and Fiedurek, 2003 A. niger mutant UV1 120 Pandey et al., 1999 A.parasiticus 2.9 Neagu & Bahrim 2011 Cladosporium sp. 10.9 Pandey et al., 1999 Fusarium sp. 0.080 Pandey et al., 1999 Penicillium sp. 50 Pandey et al., 1999 Penicillium sp. 91–4 3.74 Pandey et al., 1999 P. rugolosum 54 Pandey et al., 1999 P. spinulosum 1.67 Ertan et al., 1999 P. trzebinskii 11 Pandey et al., 1999 Streptomyces sp. 32 Pandey et al., 1999

Table 2.9 Inulinase producing microorganisms and their maximum yield

79

Table 2.9 continued

Micro-organism Maximum Reference Enzyme yield [U/ml] FUNGI

S. rochei E87 Pandey et al., 1999 1 Trichoderma viride Ertan et al., 1999 1.18 Rhizoctonia solani Ertan et al., 2003 1.792 Chrysosporium pannorum Xiao et al., 1988 115 Geotrichum candidum Mughal et al., 2009 45.65 Aspergillus ficuum Chen et al., 2011 193.6a YEASTS C. pseudotropicalis IP513 25000* Pandey et al., 1999 Kluyveromyces fragilis 7 Pandey et al., 1999 K. fragilis ATCC 12424 355 Pandey et al., 1999 K. lactis 43.7 Pandey et al., 1999 K. marxianus 56000* Pandey et al., 1999 K. marxianus 176 Silva-Santisteban and Filho, 2005 K. marxianus 127 Kalil et al., 2001 K. marxianus ATCC 36907 260 Pandey et al., 1999 K. marxianus ATCC 52466 0.418 Pandey et al., 1999 K. marxianus CDBB-L-278 82 Pandey et al., 1999 K. marxianus var. bulgaricus 107

80

Table 2.9 continued

Micro-organism Maximum Enzyme Reference yield [U/ml] YEASTS K. marxianus ATCC 16045 176 Santisteban et al., 2005 208 Santisteban et al., 2009 194.1 Kalil et al., 2010 127 Kalil et al., 2001 18743 Kushi et al., 2000 K. marxianus YS-1 50.2 Singh and Bhermi, 2008 47.1 Singh et al., 2006 K. marxianus var. marxianus CBS 3000 Kushi et al., 2000 6556 58000* Pandey et al., 1999 K. marxianus CBS 6556 212 Pandey et al., 1999 K. marxianus UCD (FST) 55–82 60.1 Pandey et al., 1999 Pichia guilliermondii 47.2 Gong et al., 1999 K. marxianus NRRL Y-7571 250 Mazutti et al., 2010 1139 Mazutti et al., 2007 1294 Sguarezi et al., 2009 1317 Treichel et al., 2009a 2620.9* Treichel et al., 2009b Golunski et al., 2011

81

Table 2.9 continued

Micro-organism Maximum Enzyme yield Reference

[U/ml]

BACTERIA Bacillus sp. 5.14 Pandey et al., 1999 B. subtilis 430 A 50–70 Pandey et al., 1999 C. acetobutylicum IFP 912 43.7 Pandey et al., 1999 C. acetobutylicum ABKn8 6.06 Pandey et al., 1999 C. thermosuccinogenes 0.011 Pandey et al., 1999 Flavobacterium mulivorum 0.456 Pandey et al., 1999 Pseudomonas sp. 65 15* Pandey et al., 1999 Staphylococcus sp. 0.634 Pandey et al., 1999 Streptomyces sp. GNDU 1 0.552 Gill et al., 2003 Streptomyces sp. ALKC4 0.524 Gill et al., 2006 9400* Sharma et al., 2007; Yields are expressed in U/ml if not otherwise specified. Data marked with “*” are U/g

a - Units/gram of dry substrate (U/gds)

82

Others* Biofuels 3.3% Biofuels and and waste Nuclear 12.9% 5.8% waste Oil Others* 10.2% 41.3% Natural gas 0.8% 15.2%

Natural gas 20.9%

Oil Hydro 32.8% 2.3% Coal/peat Electricity 10.0% 17.3% Coal/pea t 27.2% World's total energy World's total primary energy consumption in 2009 supply in 2009

12,150 Mtoe 8,353 Mtoe *Mtoe – Million tonnes of oil equivalent

Figure 2.1 World’s total energy supply and consumption in 2009 (IEA, Key World Energy Statistics, 2011). *Others include geothermal, solar, wind, heat, etc.

83

Biofuels

Primary Secondary

3rd generation Firewood, wood 1st generation 2nd generation Substrate: Algae, seaweeds chips, pellets, Substrate: Edible crop Substrate: lignocellulosic biomass

animal waste, products (seeds/grains) Biodiesel from microalgae forest and crop Bioethanol and biodiesel

residues, landfill Bioethanol or biobutanol by produced from novel starch, oil and Bioethanol from 84 gas. fermentation of starch sugar crops such as Jatropha, (from wheat, barley, corn, cassava or Miscanthus; microalgae and seaweeds potato) or sugars (from Hydrogen from green sugar cane, sugar beet, etc.) Bioethanol, biobutanol, syndiesel produced from lignocellulosic microalgae and microbes Biodiesel by materials(e.g. straw, wood, and transesterification of oil grass) crops (rapeseed, soybeans, sunflower, palm, coconut, Methanol, Fischer-Tropsch used cooking oil, animal gasoline and disesel, mixed fats, etc.) alcohol, dimethyl ether and green diesel by thermochemical processes.

(Dragone et al., 2010; Nigam & Singh, 2011) Biomethane by anaerobic digestion

Figure 2.2 Major classifications of biofuels

Solar 1% Geothermal 2% Coal Nuclear Wind 13% 20.1% 8.5% Biomass waste 5% Renewable Biofuels 21% Natural Energy Biomass 48% gas 9.5% 25.5% Wood 22% Petroleum 36.3% Hydropower 36%

Total: 97.18 quadrillion BTU Total: 9.13 quadrillion BTU

Figure 2.3 U.S. Primary energy consumption by energy sources, 2011 85

NGPL Hydropower 3.7% 34% Crude oil Nuclear Geothermal 15.4 10.6% 2% 15.4% Solar/PV 2% Renewable Wind 13% Natrual gas Energy 30.1% 11.8% Biofuels 22% Biomass 49 % Coal *Other 28.4% Biomass 27%

Total: 78.16 quadrillion BTU Total: 9.24 quadrillion BTU

Figure 2.4 U.S. Primary energy consumption by energy sources, 2011 (*Source: U.S. Energy Information Administration, Monthly Energy Review, Table 1.2, 1.3 & 10.1 (August 2012)

Acidogenic phase- Growth phase Solventogenic phase-sporulation

Figure 2.5 Acidogenic and solventogenic phase of Clostridia

86

87

Figure 2.6 Metabolic pathway of Clostridium acetobutylicum (Thick arrows indicate reactions which activate the whole fermentative metabolism. Gray letters indicate genes and enzymes responsible for the reactions. CAC and CAP numbers are the ORF numbers in genome and megaplasmid, respectively). (References: Lee et al., 2008; C. Jin et al., 2011).

Figure 2.7 Pretreatment of lignocellulosic structure (Reference: Mosier et al., 2005)

88

Figure 2.8 GF2 Fructan

Figure 2.9 F3 Fructan

(Reference: Kelly, 2008)

89

a)

b)

c)

d) 1&6-kestotetraose (bifurcose)

Figure 2.10 Structures of fructo-oligosaccharides *The starting molecule sucrose is shown in boxes. (References: Chatterton et al., 1993; Livingston et al., 2009)

90

91

Figure 2.11 Enzymes involved in fructan biosynthesis (Reference: Ende et al., 2004) 1-FEH, fructan 1-exohydrolase; 1-FFT, fructan: fructan 1-fructosyltransferase; 1-SST, sucrose: sucrose 1- fructosyltransferase; 6-FEH, fructan 6-exohydrolase; 6-SFT, sucrose: fructan 6-fructosyltransferase; 6 SST, sucrose:sucrose 6-fructosyl transferase; 6G-FFT, fructan:fructan 6G-fructosyl transferase;

CHAPTER 3: BIOBUTANOL PRODUCTION FROM INDUSTRIAL

FOOD PROCESSING WASTES

3.1 Introduction

Food waste has been a global problem for decades and has grown to enormous proportions in most countries (provide reference). Sources of food wastes differ significantly among countries. While food wastes are mostly generated by retail and consumer shops in developed countries, inadequate post-harvest technology and infrastructure, resulting in food spoilage and loss, are the major cause of food wastes generation in the developing countries (Venkat, 2012). The Environmental Protection

Agency (EPA) defines food wastes as the organic residues generated at the pre-consumer level (wastes during manufacturing, processing and handling) and at the post-consumer level (left-over foods)30. Recent reports revealed that about 40% of food produced in the

U.S. goes to garbage every year, equivalent to a loss of $165 billion31,32,33. Food waste contributes to almost 14% of total Municipal solid wastes generated in the U.S. and more than 55 million metric tonnes of avoidable food wastes (edible food wasted by

30 http://www.epa.gov/osw/conserve/materials/organics/food/fd-gener.htmL 31http://www.washingtonpost.com/business/economy/in-us-food-is-wasted-from-farm-to- fork/2012/08/21/2d5fed94-ebdb-11e1-9ddc-340d5efb1e9c_story.htmL. 32http://news.blogs.cnn.com/2012/08/22/40-of-u-s-food-wasted-report-says/. 33http://www.nrdc.org/food/wasted-food.asp.

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consumers) are generated annually (Venkat, 2012). In addition to monetary losses, redundant use of land, water, and chemicals for food production in tandem with the generation of greenhouse gases are major consequences of inefficient food use. Notably, avoidable food wastes in the U.S., which has a retail value of $198 billion, generate greenhouse emissions of at least 113 million metric tons of CO2 per year (2% of total national emissions). It is estimated that food waste alone accounts for more than one fourth of the global total freshwater consumption and nearly 300 million barrels of oil per year (Hall et al., 2009). The energy entrained in food wastes is much more than that produced by the well-established energy-producing strategies such as total annual ethanol production of the U.S. and closer to annual crude oil production from the U.S. outer continental shelf. Webber & Cuellar concluded that the amount of energy present in the food wastes generated in U.S amounts to 2030 ± 160 trillion BTU (British thermal units) or 2.21×1018 Joules (Cuéllar & Webber, 2010). While this amount is markedly greater than 1158 trillion BTUs from total ethanol produced (13.60 billion gallons in 2011)34, it is lower than 3480 trillion BTUs from total crude oil production (25.20 billion gallons in

2010)35..

The food industry is an energy-intensive sector which produces enormous quantities of wastes in both solid and liquid forms. Industrial food processing wastes contribute a substantial proportion of the total food waste especially in developed countries like U.S where the food industries are an integral part of the country’s

34 www.ethanolrfa.org/pages/statistics#A 35 www.whitehouse.gov/sites/default/files/fact_sheet_expanding_oil_production.pdf

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economy. Food processing wastes are termed residual materials produced during the conversion of agricultural commodities into marketable food items, and include wastes from raw materials, pre- and post-processing wastes, industrial effluents and sludge36.

Liquid wastes from food industries are usually disposed of in sewers (public waste water treatment systems) and in water bodies like lakes and streams. On the other hand, the normal disposal modes of solid wastes are composting and landfill applications (Hang,

2006; Levis et al., 2010).

Only about 3% of the food wastes are recycled in the U.S. for beneficial uses such as animal feed, composting or anaerobic digestion (provide reference). Several reasons have been given for such low reprocessing of food wastes which include inadequate infrastructure to process the enormous quantity of food wastes, monetary restrictions of recycling facilities, and the presence of potential contaminants in some food wastes

(Goldstein & Emmaus, 2012). Generally, disposal of wastes from food processing industries are often challenged by several factors such as limitations on direct landfill availability for solid wastes or the cost of transporting them to farm land. Direct discharge of liquid wastes to the wastewater treatment plant is mostly restricted to certain amounts or may entail higher sewer surcharges. All these factors encouraged food industries to consider anaerobic digestion as a viable option to mitigate waste disposal problems (Goldstein & Emmaus, 2012).

Notably, food processing wastes are largely biodegradable organic matter which may consist of different components such as fats, oils, protein, carbohydrates (mono-,

36 http://www.pacode.com/secure/data/025/chapter287/chap287toc.htmL

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oligo- and polysaccharides), pigments and antioxidants. These wastes have variable amounts of suspended solids, high chemical oxygen demand (COD) or biological oxygen demand (BOD) (Hang, 2006; Litchfield, 1987). These characteristics make many organic food wastes suitable for anaerobic digestion, and some are suitable for fermentation.

Production of energy-rich biofuels, like butanol, from food processing wastes could be a cost- effective way to utilize these wastes. Apart from being a viable, alternate transportation fuel, butanol can help meet industrial energy needs, offer economic benefits by reducing the amount of energy purchased from overseas, and minimize waste disposal costs and environmental pollution (Wang, 2008).

One of the objectives of this study was to obtain food processing waste samples from major food processing companies in Ohio. To accomplish this goal, a list of food processing companies was obtained from the Ohio BioProducts Innovation Center,

Columbus. We sorted this list based on annual revenue. Food processing facility with annual revenue of more than $1 million is regarded as a major food manufacturer for the study. A total of 256 companies matched the revenue criterion and we requested waste samples from all of them. Ten of the companies responded positively and provided a total of 34 different samples ranging from vegetables wastes to digester sludge wastes. In addition to these, we also obtained 14 different waste samples from the Parker Food

Science and Technology Pilot plant, Columbus, The Ohio State University. The different types and description of the 48 collected wastes are presented in Appendix B.

Eliciting information about food waste from any kind of food industry is a key challenge. Also, reporting the amount of food wastes generated by the food processing

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facilities is not mandatory by law and providing the information to EPA is purely voluntary. Earlier attempts by several researchers to gather data about total food processing waste biomass availability in Ohio were not fruitful. Companies were unwilling to disclose the amount of wastes they generate and a very poor response rate was obtained by a food waste biomass survey (Jeanty et al., 2004). Although, some food processing facilities might keep track of waste for budgeting purposes or for monitoring production efficiency, the scale they use to measure food wastes is neither uniform nor follows any specific standards. Furthermore, most facilities depend on service contracts to haul their food wastes. The service contractors also may keep track of waste based on hauling costs or the number of compactor bins filled over time. These wastes might not be just food scraps, unless they purposely separate the food wastes or have steps in the process where it can easily be tracked. All these complications make it hard for food processing facilities to have exact amount of annual food wastes generated in their facility and thus, may be a contributing factor to the poor response rate (Personal communication from EPA’s Environmental planner).

Ohio food processing firms are concentrated mostly in Hamilton, Franklin,

Cuyahoga and Stark counties and these counties probably produce large amounts of food processing wastes (Jeanty et al. 2004). In the quest to create a food biomass inventory in

Ohio, Jeanty and coworkers (Jeanty et al. 2004) drafted food biomass maps for all the counties in Ohio using GIS (Geographical Information System) techniques. Their study was based on the premise that the volume of the waste generated is proportional to the food production. Company size, the type of products produced, and annual sales were

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considered prior to generating the inventory (Jeanty et al., 2004). If actual amount of food processing biomass waste generated in Ohio is known, it would help streamline the use of waste streams for Ohio’s product and energy needs.

Our aim for obtaining different industrial food processing wastes is to characterize the wastes and evaluate their suitability for cost-effective conversion to value-added products in lieu of expensive disposal. Our preferred end use is butanol production. If a waste substrate proves unfit for butanol production, then other possible applications of the substrate will be suggested. The probable modes of applications of all the wastes are reported in the Table 3.6. The type of wastes we acquired and how industries treat them at present are described in Appendix A.

3.2 Materials and Methods

In order to classify food wastes based on type, energy potential and mineral composition, we analyzed percent total solids, ash, pH, energy content, elemental composition, total carbon and nitrogen content of each sample as follows.

3.2.1. Determination of total solids and moisture

Total solids and moisture content of samples were estimated according to

TMECC’s (Test Methods for the Examination of Composting and Compost) method

03.09-A. Instead of the recommended drying temperature (70±5°C), we opted to dry food waste samples at 50±5°C in a hot air oven to prevent caramelization of sugars present in the food waste samples which might affect the energy content estimation.

Also, at higher drying temperatures, loss of volatile compounds and changes in concentrations of nutrients are possible (Bhuiyan et al., 2010, Cornish et al., 2013). 97

Since conventional oven drying of liquid and fat-rich samples posed some challenges such as low-melting temperature of saturated fats, we dried them using freeze drier (Model 259044, VirTis Freezemobile).

All the samples were dried until further weight change was undetectable. Total solids were reported as percentage of dry solids contained in the fresh samples.

3.2.2. Determination of ash content

TMECC’s method 03.02-A (Un-milled material ignited at 550°C without inert removal) was followed to measure ash and volatile content of food waste samples. Pre- weighed samples were ignited at 550°C in the presence of excess air for 2-3 h in a forced air muffle furnace (Barnstead Thermolyne 30400-Series Furnace; Model: F30428C-80).

The remaining ash from each sample was cooled in a desiccator at room temperature and its final weight noted. The percentage of ash content (on dry matter basis) of the samples was estimated.

We also measured the ash content of waste samples using bomb calorimeter

(Model: C 2000 Basic, IKA) i.e. after complete combustion of samples, the residual ash remaining in the combustion vessel was measured. This ash measure was less and did not correlate with ash measured through the standard method. The reason for this disparity could be more complete combustion of samples in the bomb calorimeter.

However, it is more likely that the cause is spurting of samples out of the combustion vessel during combustion we observed (results in loss of ash) in the bomb calorimeter.

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3.2.3. Estimation of Total Organic Carbon (TOC)

The total organic carbon (TOC) represents sugars, starches, proteins, fats, hemicelluloses, cellulose and lignocellulose contents present in food processing waste samples that are biodegradable. TOC of samples was measured in accordance with

TMECC’s standard method 04.01-A (Combustion with CO2 Detection). According to

TMECC’s definition, total organic carbon does not include inorganic carbonate fractions such as calcium and magnesium carbonates.

A carbon analyzer (Model: vario MAX CN- Elementar Americas) was used to determine total organic carbon in the waste samples. The analyzer works on the principle of total combustion of a sample in an oxygen-rich atmosphere of a 1370°C (2500°F) resistance furnace. The CO2 produced by the combustion is channeled into an oxygen stream through anhydrone tubes to scrub water vapor from the stream. Then, the dehydrated CO2 stream is fed into the infrared detector and the amount of CO2 produced is measured. All the carbon in the sample emits as CO2 and all the hydrogen in the sample wind-up as H2O.

3.2.4. Estimation of Total Nitrogen (TN)

Total nitrogen represents the sum of Kjeldahl nitrogen (sum of organic nitrogen and ammonia nitrogen), nitrate nitrogen and nitrite nitrogen. Total nitrogen is usually used to denote the carbon to nitrogen ratio (C:N) of any samples.

The total nitrogen content was measured using TMECC’s 04.02-D oxidation by dry combustion method. Quantitative determination of nitrogen was carried out using the principle of Dumas in an automated analyzer (Model: vario MAX CN- Elementar

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Americas) according to the manufactures’ instructions. In the Dumas method, a known mass of sample is combusted at a temperature of about 900°C in a chamber in the presence of oxygen. During combustion, the sample release carbon dioxide, water and nitrogen. The gas stream is then passed over columns to remove carbon dioxide and water leaving out nitrogen. A column containing a thermal conductivity detector at the end is then used to separate the nitrogen from any residual carbon dioxide and water and the remaining nitrogen content is measured.

Air oven dried and milled samples with no inert materials were used for both

TOC and total nitrogen determination. All the samples were stored separately in sealed coin envelopes in a hot air oven at 60°C before using them for the estimation.

Both total organic carbon and nitrogen content values are reported as percentage of the dry weight of samples.

3.2.5. Determination of major, minor and trace elements

The total elemental composition of samples was measured by inductively coupled plasma optical emission spectroscopy (ICP-OES) (Teledyne Leeman Labs Prodigy Dual view ICP). The completely dried samples were ground and weighed before transferring them to polytetrafluroethylene (PTFE), also known as Teflon, vessels and subjected to microwave digestion with 7 mL of concentrated HNO3 (TMECC 04.12-A). Microwave digestion is considered to have significant advantages over other digestion procedures because of shorter digestion time, due to high pressure and temperature achieved within the vessels, and the closed vessel system prevents cross-contamination and loss of volatile elements (D. H. Sun et al., 2000). The digested samples were then cooled at

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room temperature and transferred to an ICP-OES autosampler for analysis. The ICP ionizes the sample digest with the help of argon gas (~10-15 L/min) in an applied radio frequency field. Each element after ionization, exhibits a characteristic emission spectrum and the intensity of each spectrum reflects the concentration of the element in the sample. The detectors identify elements and measure their spectral intensity to derive qualitative and quantitative analysis of the elements (Plank, 1992)37. TMECC’s standard

04.05, 04.06 and 04.07 were followed for estimating the major, mineral and trace elements. The estimated major elements and minor elements are reported in mg and µg, respectively, per g of sample (on dry matter basis).

All the above analyses were done in STAR lab located on the OARDC campus,

Wooster, Ohio.

3.2.5. Measurement of pH

The pH of the food waste samples was measured using TMECC’s standard method of 04.11-A (1:5 Slurry Method). Fresh waste samples were blended with deionized (DI) water at a ratio of 1:5, w/w basis. Samples were shaken at 180 rpm for 20 to 30 minutes at room temperature to allow salts solubilize in the DI water. The pH was measured with an electrometric pH meter probe directly in the resultant slurry while swirling. If there was a change in pH reading, swirling of the sample flask was continued until a stable pH reading was attained. Often pH change of the waste samples is used as an indicator of stability, mobility and availability of metals and nutrients.

37 http://www.shimadzu.com/an/elemental/oes/oes.htmL

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3.2.6. Determination of energy content

A food’s calorific value is determined by its content of fat, carbohydrate and protein. To determine the gross calorific value of solid and liquid waste samples, we used a combustion calorimeter (Model: C 2000 Basic version 1, IKA). In this, the decomposition vessel (Model: C 5010) is filled with a weighed quantity of sample which is incinerated in the presence of pure oxygen. The instrument estimates the gross calorific value as the quotient of the amount of heat liberated upon total combustion and the weight of the original sample. The working principle of the calorimeter is to capture the released heat energy with a reservoir of water, which has a high capacity for absorbing heat. The temperature of the water reservoir is measured at the beginning and at the end of the experiment. The increase in temperature (in °C) times the mass of the water (in g) gives the amount of energy captured by the calorimeter, in kilocalories.

Energy content of food processing waste samples was estimated with a minimum of 3 repetitions until consistent values were obtained for a sample. Liquid samples can be fed to the calorimeter directly, but we opted for freeze drying of the liquid samples before energy estimation due to ease of handling and making direct estimations based on dry matter possible.

Results of food processing wastes analyses are grouped together based on predetermined criteria and are presented in Figs 3.1-3.4.

3.2.7 Selection of an ideal substrate for butanol production

Butanol can be produced from cellulosic, hemicellulosic, and starchy substrates, albeit lignin and fat-rich substrates are not suitable due to their low carbohydrate content.

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While cellulose and hemicellulose must be hydrolyzed to simple sugars prior to fermentation by solventogenic Clostridia, direct utilization of starch by this group of microorganisms has been demonstrated by several investigators (Gutierrez et al., 1998;

Grobben et al., 1993; Madihah et al., 2001; Jesse et al., 2002). The goal of this study is to evaluate locally-available food processing wastes for low-cost butanol fermentation.

Direct use of food wastes for fuel production may help reduce industrial waste processing and transit costs, and could encourage industries to consider this option for the disposal of their wastes. Given that starch is rapidly fermented into butanol by solventogenic

Clostridia, we shortlisted food wastes that whose major carbohydrate component is starch. From this list of starchy wastes, after taking into consideration energy content, percent carbon and nitrogen, C/N ratio and amount of elements present in the samples

(Figure 3.2d, 3.3d, 3.4d), we chose eight different food wastes and quantified their starch content.

Generally, anaerobic microorganisms can consume carbon 25-30 times faster than nitrogen and a C/N ratio of 20-30 is regarded as ideal (Alshiyab et al., 2008). However, the ideal C/N ratio for butanol production is unclear and has scarcely been reported in literature. Madihah et al. (2001) concluded in their studies that there is no clear relationship between C/N ratio and butanol production. Also, they showed that individual concentrations of carbon and nitrogen are more significant factors in maximizing the butanol production than the ratio. Since no clear demonstration of effect of C/N on butanol production is available, we choose the substrates with C/N ratio (range

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of 15-30) that is ideal for maximum decomposition rate of organic matter. An ideal substrate also should have low total salts, and high total energy content.

The two acquired dairy wastes are milk dust powder and whey liquid waste. The milk dust powder was recovered from the dust collector during spray-drying of milk.

According to the manufacturer (International dairy ingredients, Inc, Wapakoneta, OH), it is basically a blend of different milk powders. Milk dust powder is identified as one of the hazardous materials in that it could cause explosion in the food industry and severe damage to people and properties. Suspension of milk dust in the air (between concentration 75-1000 g/m3 of air)38 can explode or undergo self-ignition if it comes in contact with hot surfaces (Davis et al., 2011; Ministry of Labour, New Zealand, 1993). A drier with 10 metric tonnes/h capacity could generate 80.30 kg of airborne milk dust powder39. The projected milk powder production (includes skimmed milk and whole milk powder) in the USA in 2012 is estimated to be 1.02 million metric tons, which translates into 116.44 metric tons/h (USDA, 2012) and thus, the approximate amount of milk dust powder generation in the U.S. is estimated to be around 900 kg/h. Other than being used as animal and cattle feed, this enormous amount of dairy waste (1.15×1014

Joules/year) upon careful handling has many potential applications in biofuel production40. The milk dust powder constitutes mainly of lactose, protein, fat and some minerals. Further, it retains all the natural properties of milk such as color, flavor and solubility. On mixing with water, it resembles typical raw milk in appearance.

38 www.hse.gov.uk/food/dustexplosion.htm 39 http://dustexplosions.blogspot.com/2008/12/milk-powder-combustible-dust-hazards.html 40 http://www.usdec.org/files/PDFs/US08_G.pdf

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The leftover substance after precipitation of casein during cheese making is known as whey liquid waste. Whey liquid has 55% of the total milk nutrients, specifically 45-50 g/L of lactose, 6-8 g/L soluble proteins, 4-5 g/L lipids, 0.5% minerals and 93.50% of water. The major mineral salts are KCl (more than 50%), NaCl and calcium salts (mostly phosphate). Whey liquid waste also contains low amounts of lactic acid, citric acid, urea, uric acid and B group vitamins (Athanasiadis et al., 2002;

Chatzipaschali & Stamatis, 2012; Foda et al., 2010; Napoli, 2009). Global annual production of whey is estimated to be approximately 200 million metric tonnes with an increment rate of 2% every year (Illanes, 2011). In 2006, U.S produced a total of 41.05 million metric tonnes of whey liquid (Ling, 2008).

Lactose is a disaccharide molecule made up of glucose and galactose, and

Clostridia have the ability to transport lactose into their cells (Servinsky et al., 2010; Yu et al., 2007). Butanol production from whey liquid (an industrial dairy by-product) has been well-researched and documented in the last two decades (Ennis & Maddox, 1989;

Foda et al., 2010; Qureshi & Maddox, 2005). Although using an industrial dairy waste directly without any kind of processing (protein removal, concentration or purification of substrate) has not been reported. Hence, the suitability of direct use of either whey liquid waste or milk dust powder was examined after estimating the free lactose availability in the substrates.

The milk dust powder has higher lactose content (~425 g/L) than whey liquid

(~27 g/L). The whey liquid would need to be concentrated to increase the sugar

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concentration before use as a fermentation substrate which is an additional process and cost.

In conclusion, after estimating starch and lactose contents of obtained food wastes, we selected the best three starchy (highest starch content) and milk dust powder wastes for batch fermentation trials. The selected food wastes were inedible dough

(supplied by ConAgra Foods Inc, Troy, Ohio), breading and batter liquid (provided by

Lake Erie Frozen foods Co Inc, Ashland, Ohio) and milk dust powder (International

Dairy Ingredients Inc, Wapakoneta, Ohio). The starch and lactose contents of these food wastes are presented in Table 3.1 and 3.2, respectively.

3.2.8 Selection of microorganism, culture maintenance and inoculum preparation

The best strain reported till date for starch utilization is C. beijerinckii BA 101, a mutant strain of C. beijerinckii NCIMB 8052 (Qureshi & Blaschek, 2000). Use of C. acetobutylicum ATCC 824 for ABE fermentation using starch as substrate has also been reported (Gutierrez et al., 1998). We have stocks of C. acetobutylicum ATCC 824 and C. beijerinckii NCIMB 8052 in our culture collection, and these two microorganisms were tested for ABE production using selected sample wastes as substrates.

C. beijerinckii NCIMB 8052 (ATCC 51743) and C. acetobutylicum ATCC 824 were procured from American Type Culture Collections, Manassas, VA, USA. These microorganisms were maintained as stock spore suspensions in sterile double-distilled water at 4°C. For culturing, 200µL of spores from each microorganism stock were heat- shocked at 75°C for 10 min, cooled on ice for 3 min, and then inoculated into 10mL of tryptone-glucose-yeast extract (TGY) medium and incubated anaerobically at 35±1°C for

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12-14 h (Please provide reference). When the optical density (OD600 nm) of the culture reached between 0.9-1.1, 8 mL of the actively growing culture was transferred into 92 mL anoxic presterilized TGY medium and incubated for 3-4 h (OD of 0.9-1.1 at 600 nm).

To bring about anaerobiosis, loosely capped 150 mL Pyrex screw bottles containing sterilized 100 mL TGY medium was kept overnight (14-16 h) in an anaerobic chamber

(Coy Laboratory Products Inc., Ann Arbor, MI) with an atmosphere of 82% N2, 15%

CO2, and 3% H2 prior to inoculation with C. beijerinckii NCIMB 8052 or C. acetobutylicum ATCC 824. Keeping the growth medium inside the anaerobic chamber for 14-16 h period facilitates exchange of gases between the medium and the gases present in the chamber and thus, removes residual oxygen from the TGY medium (Ezeji

& Blaschek, 2008; Richmond et al., 2012; Zhang et al., 2011).

3.2.9 Medium preparation and ABE fermentation

Batch fermentation studies of food processing wastes were conducted in 150 mL

Pyrex screw capped media bottles. Both C. beijerinckii NCIMB 8052 and C. acetobutylicum ATCC 824 were tested separately to determine their butanol producing ability from selected substrates. The starchy sample wastes were reconstituted in Pyrex bottles with water (with 1 g/L yeast extract) to make medium equivalent to ~50 g/L starch, and autoclaved at 121°C for 15 min. The amount of substrate required to make 50 g/L starch medium is shown in Appendix D. The autoclaved media were cooled to 40°C before transferring them into the anaerobic chamber (Coy, Ann Arbor, MI) and kept at

35±1°C for 14-16 h. P2 medium (60 g/L glucose and 1 g/L yeast extract) was used as a control for both microorganisms. Prior to the inoculation with 6 mL culture of C.

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beijerinckii NCIMB 8052 or C. acetobutylicum ATCC 824 cells to the 91 mL medium, filter-sterilized P2 stock vitamin (0.1 g/L para-amino-benzoic acid; 0.1 g/L thiamine;

0.001 g/L biotin), buffer (50 g/L KH2PO4; 50 g/L K2HPO4; 220 g/L ammonium acetate) and mineral (20 g/L MgSO4.7H2O; 1 g/L MnSO4.H2O; 1 g/L FeSO4.7H2O; 1 g/L NaCl) solutions of 1 mL each were added (Richmond et al., 2012; Zhang et al., 2011).

Additionally, the milk dust powder medium was prepared by autoclaving a mixture of ~12 g milk dust powder and 1g yeast extract at 121°C for 15 min. Following cooling to 40 C, the mixture was transferred into the anaerobic chamber and ~80 mL anoxic sterilized water (reconstituted to 11-12% total solids in the medium) was added to bring the concentration of lactose in the medium to 50 g/L. Prior to inoculation with either C. beijerinckii NCIMB 8052 or C. acetobutylicum ATCC 824, the medium was supplemented with P2 stock solutions as described above.

The pH of breading and milk dust powder media was pH 6.5 after addition of P2 stock solutions, whereas the pH of inedible dough and batter liquid media was 5.4 and

5.6, respectively. Therefore, the pH of the inedible dough and batter liquid media was adjusted to 6.5 using 3M KOH prior to inoculation with either C. beijerinckii NCIMB

8052 or C. acetobutylicum ATCC 824. Three milliliters of samples were taken every 12 h to measure pH, residual sugars, ABE and acid production. Unless otherwise stated, the start pH of all the fermentation media was 6.5, temperature was maintained at 35 ±1°C, and no agitation or pH control was employed. All fermentations were done in triplicate.

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3.2.10 Analytical methods

Growth of C. beijerinckii NCIMB 8052 or C. acetobutylicum ATCC 824 was determined by plate counts (viable cell counts). Each counted C. beijerinckii NCIMB

8052 or C. acetobutylicum ATCC 824 colony is regarded as a colony forming unit

(CFU). Culture samples were serially diluted in 10 mL of TGY medium and plated in 10 mL of semi-solid TGY agar (0.45% agar in TGY medium). The plates were incubated anaerobically for 24-48 h at 35±1°C and the number of CFUs was counted and expressed as CFU/ml of original culture (Jesse et al., 2002; Nielsen et al., 2009). The residual concentrations of glucose, maltose, maltotriose and lactose were measured by high performance liquid chromatography (HPLC) with a refractive index (RI) detector

(Agilent Technologies 1200 Series) and an organic acid column (Rezex ROA-Organic

+ Acid H column, 300 mm X 7.8 mm). The mobile phase was 0.0025M H2SO4 (Fluka,

50% sulfuric acid, 0.35 ml diluted with millipore water to 1 L) operated at a flow rate of

0.6 mL/min. All samples were injected by automatic sampler and the injection volume is

10µL. The column and detector temperature were 80°C and 55°C, respectively.

The concentration of fermentation products, such as acetone, butanol, ethanol, acetic acid and butyric acid, was measured using a gas chromatography (GC) system

(7890A Agilent Technologies Inc., Santa Clara, CA, USA), equipped with a flame ionization detector (FID) and 30m X 320 µm (length x internal diameter) with a 0.5µm

(HP-Innowax film) J x W19091N-213 capillary column (Zhang et al., 2011). Yield was calculated as the maximum amount of ABE produced per gram of substrate utilized and is expressed in g/g. ABE productivity was estimated as maximum ABE produced (g/L)

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divided by the corresponding fermentation time (h) or when the fermentation ceased divided by the total fermentation time (h) (Ezeji & Blaschek, 2008; Richmond et al.,

2012). For both HPLC and GC analyses, the samples were diluted 5 times with deionized water and centrifuged at 10,000 x g for 8 min to extract clear supernatant prior to analysis.

The starch content of the food waste samples was measured using the slightly modified megazyme enzymatic starch assay as described by Galicia et al. (2008). The sample (20 mg) was wetted with 40 µL of 80% aqueous ethanol and stirred for 5 min.

Then, 600 µL of α-amylase in MOPS (3-(N-morpholino)propanesulfonic acid) buffer at pH 7 was added and incubated for 6 minutes in a boiling water bath. On cooling the samples to 50°C, 800 µL of sodium acetate buffer and 20 µL of amyloglucosidase were added and incubated at 50°C for 30 minutes. The entire content was transferred to a 50 mL plastic corning tube and18.54 mL of distilled water was added and centrifuged at

3000 rpm for 10-20 minutes until a clear solution was obtained (Galicia et al., 2008).

From the supernatant, one mL of the content was further diluted with 9 mL of distilled water and mixed thoroughly. From this diluted mixture, 500 µL of the mixture was pipetted into a glass tube and 1000 µL of anthrone reagent (Acros Organics) was added.

The contents of the glass tube were incubated for 10 minutes (95-100°C) and cooled for

10 minutes on ice. The cooled samples were vortexed and analyzed in a DU800 spectrophotometer (Beckman Coulter Inc., Brea, CA) at 630 nm . The OD obtained was correlated to a standard calibration curve from pure glucose and thus, the starch content was estimated using the formula:

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∗∗∗∗ Starch (mg/100 mg) of samples =

Where, x = glucose concentrations (mg/ml), df is the dilution factor, V is the original volume of starch extract (20 ml), dw = original weight of samples (20 mg) and hf is the starch hydrolysis factor 0.9. All starch assays were done in triplicate with a duplicate of standard starch.

3.3 Carbohydrate metabolism

It is known that Clostridium sp. can utilize a wide variety of carbohydrate substrates as energy source. The metabolic pathways involved in lactose and starch utilization by C. acetobutylicum ATCC 824 and C. beijerinckii NCIMB 8052 are briefly described below:

3.3.1 Lactose metabolism in solventogenic Clostridium species

The two different enzymes involved in lactose metabolism are β-galactosidase, which hydrolyzes lactose into glucose and galactose, and phospho-β-galactosidase which hydrolyzes lactose 6-phosphate into glucose and galactose 6-phosphate. Yu et al (2007) found that C. acetobutylicum ATCC 824 grown in lactose medium exhibited increased phospho-β-galactosidase activity than in glucose medium. Also, a basal level of β- galactosidase activity was observed when C. acetobutylicum ATCC 824 was grown in glucose and lactose media. Further, it was revealed that lactose is transported by this microorganism through a phosphoenol-pyruvate (PEP) dependent phosphotransferase system (PTS) and the phosphorylated derivative (lactose 6-phosphate) is further metabolized by the enzyme phospho-β-galactosidase into glucose and galactose 6-

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phosphate. It is plausible that glucose is phosphorylated further and incorporated into the glycolytic pathway whereas galactose 6-phosphate is metabolized by the tagatose 6-P pathway (Yu et al., 2007). The genes needed for lactose and galactose metabolism are found in the genome of C. acetobutylicum ATCC 824 around CAC2950 and CAC2966 region. Higher expression of these genes is noticed only in media with lactose or galactose (Servinsky et al., 2010). Detailed reports on C. acetobutylicum ATCC 824 genome and lactose metabolism are available elsewhere (Servinsky et al., 2011; Yu et al.,

2007).

It also was demonstrated that C. beijerinckii NCIMB 8052 has been shown to have 47 sets of PTS II genes it uses to metabolize complex carbohydrates (Shi et al.,

2010). Detailed information on lactose metabolism in C. beijerinckii NCIMB 8052 has not been reported. However, it is plausible that lactose transport and metabolism in C. beijerinckii NCIMB 8052 could be similar to C. acetobutylicum ATCC 824 (Servinsky et al., 2011; Yu et al., 2007) or sucrose, another disaccharide molecule (Mitchell et al.,

1995; Tangney et al., 1998).

3.3.2 Starch metabolism in solventogenic Clostridium species

Butanol production from starch by solventogenic Clostridium species is an age- old fermentation technique. Starch has to be cleaved into glucose before it is transported into the cells of Clostridium species. Concerted actions of two enzymes, namely α- amylase and glucoamylase, are necessary to breakdown the starch. The α-amylase action is endo-acting and non-specific in that it hydrolyzes starch into oligo-, di-, and monosaccharides, whereas glucoamylase is exo-acting, and it breaks the terminal glucose

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units from the non-reducing end of the starch molecule, yielding only glucose as its hydrolysis product (Paquet et al., 1991). Most solventogenic Clostridium species produce both these enzymes and consume the resultant glucose from the starch hydrolysis

(Madihah et al., 2001). Starch is metabolized in the same pathway as glucose and maltose. The only difference in the metabolism is the transportation of starch. Starch is hydrolyzed extracellularly and the glucose is transported into the cells. A detailed account of starch metabolism in C. acetobutylicum ATCC 824 is reported by Servinsky et al. (2010). C. beijerinckii NCIMB 8052 also utilizes starch in the same glucose metabolic pathway (Lee & Blaschek, 2001; Shi et al., 2010).

3.4 Results and discussion

3.4.1 ABE production from milk dust powder

When milk dust powder was mixed with water and autoclaved at 121°C (for 15 minutes), the medium coagulated and had a curd-like appearance. Fermentation of this curd was poor and the maximum butanol produced was 1.2 g/L and 0.8 g/L, from C. acetobutylicum ATCC 824 and C. beijerinckii NCIMB 8052, respectively (data not shown). Though butanol production showed an increasing trend over fermentation time for both microorganisms, the concentration of butanol in the fermentation broth never exceeded 1.2 g/L in 72 h fermentation. The cell growth was poor which signaled poor lactose utilization by both the microorganisms (data not shown). However, it was clear from these experiments if C. acetobutylicum ATCC 824 used the milk dust powder medium better than C. beijerinckii NCIMB 8052.

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Milk powder contains two major proteins namely casein (~80%) and whey protein (~20%) (Farrell Jr et al., 2004). The coagulation of milk dust medium upon autoclaving is attributed to casein coagulation and gel formation due to increased amounts of denatured whey protein and K-casein interactions. The stability of casein in milk powder during heat treatment is contingent upon availability of optimal amounts of calcium, phosphate and citrate content. Application of severe heat (over 90°C-120°C for

10 or more minutes) affect the salt equilibrium and precipitates calcium and magnesium as phosphates or citrates, which results in casein coagulation41. Notably, raw milk dust powder has 7.08 mg of calcium, 13.05 mg of potassium and 6.38 mg of phosphorus per gram of dry matter (Figure 3.3d). Coagulated casein arranges themselves into a three- dimensional lattice and hold sugar, water, fat and minerals in position42. This phenomenon might have affected the availability of free sugars in the milk dust medium.

Furthermore, 11-12 % total solid concentration of milk dust medium might have increased the gel formation and strength. High milk concentration in the medium increases its protein concentration which in turn, increases the number of protein interactions and formation of firm gels (Anema, 2008; Feary, 2010). All these physico- chemical changes of milk dust medium during heating might have severely inhibited nutrient utilization and consequently, resulted in low acetone butanol ethanol (ABE) production.

41 chestofbooks.com/food/science/Experimental-Cookery/Coagulation-Of-Milk-Part-3.html 42 Dairy proteins prepared by the Wisconsin center for dairy research and the Wisconsin milk marketing board. www.cdr.wisc.edu/programs/dairyingredients/pdf/dairy_proteins.pdf

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To overcome this issue of curd formation, autoclaving the milk dust powder

(121°C for 15 minutes) without reconstitution was explored. Upon heating, the color of milk dust changed from light yellow to dark brown. After cooling, anoxic sterile water

(80 ml) was added to the milk dust inside the anaerobic chamber to make 91 ml fermentation medium (~50 g/L lactose). This new medium resulted in maximum butanol production of 3.12 g/L and 5.43 g/L after 72 h by C. beijerinckii NCIMB 8052 and C. acetobutylicum ATCC 824, respectively (data not shown). But, both microorganisms showed a long lag phase and produced merely 0.50-1.00 g/L butanol after 36 h fermentation. Furthermore, the autoclaved milk dust formed a stone-like structure at the bottom of the fermentation bottle which left the liquid medium clear (i.e. milk dust did not dissolve in the liquid medium). The hardened stone-like formation was because of the conglomeration of milk solids, denatured proteins, and calcium and magnesium phosphates when the milk dust is heated above 60°C43,44. The structural changes of milk dust medium are illustrated in Figure 3.14. Manually breaking this hardened surface before inoculation was found to reduce the lag phase by 12 h and increased the butanol production to 5.80 g/L and 7.24 g/L by C. beijerinckii NCIMB 8052 and C. acetobutylicum ATCC 824, respectively (Figure 3.5a & 3.8a). Increasing surface area upon breaking the milk dust increased the substrate availability to the microorganism which, in turn, was reflected in butanol production.

43 http://drinc.ucdavis.edu/dairyp/dairyp5.htm 44 http://www.encyclopedia.com/doc/1O39-milkstone.html

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Maximum total ABE produced by C. beijerinckii NCIMB 8052 was 8.15 g/L

(Figure 3.5b) (ABE yield of 0.29 g/g of substrate; ABE productivity of 0.13 g/L/h) whereas C. acetobutylicum ATCC 824 produced total ABE of 10.25 g/L (Figure 3.8b)

(ABE yield was 0.30 g/g of substrate; ABE productivity of 0.17 g/L/h). The fermentation ceased after 60 h and there was no further increase in ABE production thereafter. C. acetobutylicum ATCC 824 consumed a total of 34.72 g of lactose whereas C. beijerinckii

NCIMB 8052 consumed 27.67 g. The substrate consumption pattern of the two species is illustrated in Figure 3.11. Glucose P2 medium was used as a control in this study.

Although fermentation of this waste without any pretreatment by both microorganisms was successful, the fermentations were slower than the control. The control showed ABE yield of 0.44 g/g for C. acetobutylicum ATCC 824 and 0.43 g/g for C. beijerinckii

NCIMB 8052. The kinetic parameters of the fermentation for the two species were compared and are presented in Table 3.5. The initial medium pH of 6.5 was reduced to

5.5 after 12 h for both Clostridium species, and thereafter continued to drop till 48 h.

After 48 h, the pH stabilized and stayed between 5.2 and 5.3. At the end of the fermentation, the concentration of acetic acid and butyric acid for C. beijerinckii NCIMB

8052 was 1.38 g/L and 2.17 g/L (Figure 3.7), respectively, whereas the fermentation medium with C. acetobutylicum ATCC 824 had 1.18 g/L of acetic acid and 1.69 g/L of butyric acid (Figure 3.10, Table 3.5). Therefore, C. acetobutylicum ATCC 824 produced and re-assimilated acids (acetic acid and butyric acid) better than C. beijerinckii NCIMB

8052 which substantiates the maximum ABE production from the organism. It should be

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noted that in ABE fermentations, acids are intermediate metabolites produced by the microorganism before they are reassimilated for acetone and butanol production.

C. acetobutylicum ATCC 824 had 1.80×107 cells per ml at zero hour whereas

1.17×107 cells per ml of C. beijerinckii NCIMB 8052 was determined. Both the species displayed the same pattern of cell growth i.e. steady increase in cell growth up to 36 h and then a decline in growth (Figure 3.12 & 3.13). C. acetobutylicum ATCC 824 and C. beijerinckii NCIMB 8052 grew to a peak cell density of 7.67×109 and 3.67×109 cells per ml at 36 h of fermentation, respectively (Appendix C). The difference in growth and

ABE production between the two organisms could be due to the difference in availability of relevant genes and enzymes for lactose transport and metabolism. Phospho-β- galactosidase and β-galactosidase are the two necessary enzymes for lactose metabolism and levels of activity of these enzymes determine the efficacy of lactose utilization. Also, it is likely that presence of excess minerals and protein content in the milk dust medium might have played a role in inhibition for C. beijerinckii NCIMB 8052 more than C. acetobutylicum ATCC 824, since the later consumed more lactose and produced more

ABE than the former.

Ennis and Maddox (1986) conducted whey permeate fermentation using C. acetobutylicum P262 and found that ABE production is highly influenced by medium pH.

They observed that the highest butanol production rates occurred in the pH range of 5.1-

5.5. The pH of the milk dust fermentation medium was in accordance with the pH range for both microorganisms. The lowest pH measured for C. beijerinckii NCIMB 8052 was

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5.16 (Figure 3.6b) whereas pH 5.21 was measured for C. acetobutylicum ATCC 824 fermentation (Figure 3.9b) after 36 h.

It should be noted that continuous and fed-batch fermentation of lactose medium with different solvent recovery systems was attempted earlier by several researchers

(Qureshi & Maddox, 1990; Napoli et al., 2010 & 2011; Ennis et al., 1986; Friedl et al.,

1991). It was found that integrated fermentation and solvent recovery technique improved solvent yield, productivity, substrate utilization and concentration of solvent stream (Friedl et al., 1991; Qureshi & Maddox, 1991; Napoli et al., 2010). Since high lactose concentration can be used in a continuous fermentation mode, milk dust powder, which is rich in lactose content, could be a good substrate for large-scale continuous fermentation coupled with solvent removal systems.

3.4.2 ABE production from starchy food processing wastes

Breading

The total energy content of breading was 17.66 MJ/kg and the estimated starch content of the dry weight was 61.0% (Table 3.1). The total carbon content was 42.2% and the nitrogen content was 1.87% (C/N ratio of 22.61) (Figure 3.2d). Direct fermentation of breading with C. beijerinckii NCIMB 8052 produced total ABE concentration of 14.8 g/L at 72 h, of which butanol and acetone amount to 10.47 g/L and

2.77 g/L, respectively. The ABE yield was 0.36 g/g and productivity was 0.20 g/L/h. On the other hand, fermentation using C. acetobutylicum ATCC 824 produced total ABE concentration of 13.98 g/L after 72 h (9.74 g/L butanol and 3.32 g/L acetone) with ABE yield of 0.33 g/g and productivity of 0.19 g/L/h. It is obvious that C. beijerinckii NCIMB 118

8052 produced more solvents from the breading medium than C. acetobutylicum ATCC

824 did.

Although both the organisms showed a similar pattern of cell growth, C. acetobutylicum ATCC 824 adapted to the breading medium sooner and grew better than

C. beijerinckii NCIMB 8052. Both microorganisms showed maximum cell growth at 36 h; C. acetobutylicum ATCC 824 had 6.50×1010 and C. beijerinckii NCIMB 8052 had

3.90×1010 cells per ml (Figure 3.12 & 3.13). The initial medium pH of 6.50 dropped to around 5.1 during the acidogenic phase and increased later on to 5.50. However, the increase in medium pH was observed after 36 h for C. acetobutylicum ATCC 824 whereas it was observed after 60 h for C. beijerinckii NCIMB 8052. This result corroborates our observation that C. acetobutylicum ATCC 824 adapted to breading medium faster than C. beijerinckii NCIMB 8052. The total amount of starch utilized after 72 h of fermentation by C. acetobutylicum ATCC 824 was 42.0 g/L whereas C. beijerinckii NCIMB 8052 consumed 40.4 g/L (Table 3.3 & 3.4).

Inedible dough

The inedible dough had 63.1 % total solids (Figure 3.1d), 43.8% carbon, 2.6 % nitrogen (C/N ratio of 17.0), an energy content of 17.7 MJ/kg (Figure 3.2d), and a total starch content of 60.38 % on a dry weight basis (Table 3.1). It can be seen that the energy content, carbon and starch content of the inedible dough is comparable to that of breading. However, the ABE yield and productivity from inedible dough was better than that of breading. Inedible dough gave ABE yield of 0.37 g/g (14.44 g/L total ABE of which 9.26 g was butanol; productivity was 0.24 g/L/h) for C. beijerinckii NCIMB 8052 119

while C. acetobutylicum ATCC 824 had the yield of 0.38 g/g (16.30 g/L total ABE; 11.20 g/L butanol; 0.23 g/L/h productivity). The total amount of starch utilized by C. acetobutylicum ATCC 824 was greater (43.26 g) than C. beijerinckii NCIMB 8052

(39.45 g) (Table 3.3 & 3.5). In general, the fermentation was faster and the total ABE production at the end of 36 h was 16.13 g/L and 13.54 g/L for C. acetobutylicum ATCC

824 and C. beijerinckii NCIMB 8052, respectively. After 36 h, the increase in ABE production for both the microorganisms was very minimal (Figure 3.5 & 3.8). The cell growth of both microorganisms followed a similar pattern by showing peak cell density at 36 h followed by decline in growth (Figure 3.12 & Figure 3.13). At zero hour, C. acetobutylicum ATCC 824 had 1.92 g/L of acetic acid and 0.67 g/L of butyric acid whereas C. beijerinckii NCIMB 8052 had 1.64 g/L of acetic acid and 0.81 g/L butyric acid. Acetic acid and butyric acid production showed the typical increase during the acidogenic growth phase and then decrease during the solventogenic growth phase

(Richmond et al., 2011). (Figure 3.7 & 3.10)

Batter liquid

The liquid batter waste is composed of 62.1% starch on a dry weight basis (Table

3.1), 33.0 % total solids (Figure 3.1d), C/N ratio of 28.8 (42.6% carbon and 1.48% nitrogen), and an energy content of 17.3 MJ/kg (Figure 3.2d). Among all the starchy wastes fermented, batter liquid emerged as the best substrate for ABE production by both microorganisms. While C. acetobutylicum ATCC 824 produced 17.4 g/L of ABE (11.76 g/L butanol, ABE yield of 0.43 g/g, and productivity of 0.24 g/L /h), C. beijerinckii

NCIMB 8052 produced 15.1 g/L ABE (10.00 g/L butanol, ABE yield of 0.37 g/g, 120

productivity of 0.31 g/L/h) using batter liquid as substrate. The reason for low ABE productivity by C. acetobutylicum ATCC 824 is because the maximum ABE was produced at 72 h whereas in C. beijerinckii NCIMB 8052 the maximum ABE production was observed at 48 h (Figure 3.5 & 3.8). The peak cell density for C. acetobutylicum

ATCC 824 was observed at 36 h (8.86×1010 cells per ml) whereas it was observed at 60 h

(2.80×1010 cells per ml) for C. beijerinckii NCIMB 8052 (Figure 3.11& 3.12; Appendix

C). Unlike the breading and inedible dough medium, the accumulation and reassimilation of acids for ABE production was much more clearly demonstrated by both microorganisms, especially C. beijerinckii NCIMB 8052. Higher amount of ABE from this waste by the two species could be attributed to the better reassimilation of acids by the organisms.

The maximum ABE production, productivity, and cell growth from the starchy waste fermentation by C. acetobutylicum ATCC 824 follows this pattern: Batter liquid >

Inedible dough > Breading (Figure 3.8, Table 3.4). This same pattern was observed from

C. beijerinckii NCIMB 8052 with regard to ABE productivity and onset of solventogenesis. However, this microorganism displayed varied pattern with regard to maximum ABE production (Figures 3.5, Table 3.3). The maximum ABE production by

C. beijerinckii NCIMB 8052 from starchy wastes is as follows: Batter liquid (15.11 g/L)

> Breading (14.80 g/L) > Inedible dough (14.44 g/L).

While maximum growth of C. beijerinckii NCIMB 8052 obtained when breading and inedible dough were used as substrates were 3.90×1010 and 3.50×1010 cells per mL, approximately 2.80×1010 cells per ml was obtained when batter was used as substrate.

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Therefore, there was no clear relationship between cell growth of C. beijerinckii NCIMB

8052 and ABE production. It should be noted that the difference in ABE production and cell growth of C. beijerinckii NCIMB 8052 from the starchy media were not as prominent as it was with C. acetobutylicum ATCC 824. Nonetheless, batter liquid gave the maximum ABE production for both the microorganisms.

Though all the three substrates were similar in starch, energy, carbon and nitrogen, and mineral contents, the appearance of the media differed slightly among them. The batter liquid medium was colloidal; the inedible dough medium was semi- solid (more of a jelly) while the breading formed a firm gel (i.e. did not shake). To put it simply, the order of least to most gelled starchy medium follows this pattern: Batter liquid > Inedible dough > Breading. Notably, ABE production by C. acetobutylicum

ATCC 824 from these starchy substrates followed the same pattern. This observation led to the hypothesis that retrogradation of starch did slow down the fermentation or reduce

ABE production. When starch-water mixture is heated, the starch granules absorb water molecules and irreversible loss of crystallinity structure and solubilization of amylose occur (which is known as gelatinization) (Morris, 1990; Zeng et al., 1997). In simple terms, gelatinized starch is a hot amylose solution with swollen granules as filler in an amylopectin skeleton (Srichuwong et al., 2005; Morris et al., 1990). On cooling to room temperature, amylose molecules rearrange and form double-helical associations (Tamaki et al., 2011) whereas amylopectin forms crystalline structure by reassociation of external short branches (Singh & Sandhu, 2007). The phenomenon of starch molecules to re- arrange into a more rigid crystalline state and forms firm gel upon cooling is known as

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retrogradation. Though both amylose and amylopectin are responsible for retrogradation, amylose content of starch plays a major role in retrogradation of starch which readily takes place when cooled to room temperature. On the other hand, the rate of retrogradation of amylopectin is much slower in general (i.e. long-term storage induces gel formation) (Srichuwong et al., 2005; Lauro et al., 1997; Keetels et al., 1996). At higher concentrations of amylose (<1%), the crystallized amylose form a three- dimensional network which traps water to create a gel which increases the viscosity of the medium (Case et al., 1996; Keetels et al., 1996). It should be remembered that availability of amylose varies between 20-30% depends on plant source (Tamaki et al.,

2011; Case et al., 1996). Apart from the ratio of amylose and amylopectin content in starch, other factors such as degree of polymerization, temperature, concentration of starch in the medium, botanical source of starch are also known to influence rate of retrogradation (Jacobson et al., 1997; Ezeji et al., 2005). Srichuwong et al (2005) summarized that presence of minor ingredients such as phospholipids and phosphate monoesters in starch are found to impact the retrogradation properties. Therefore, these factors can be attributed for the difference in gelling and pasting characteristics of the starchy media tested.

Retrogradation of starch (upon cooling of autoclaved media) increased the apparent viscosity of the media. Retrogradation was known to significantly lower the availability of starch for enzymatic action (Ezeji et al., 2005; Fredriksson et al., 2000).

Therefore, it resists enzymatic activity and slows down the rate of hydrolysis. This phenomenon could have affected the amount of fermentable sugars available in the

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swelled-starchy media. In general, reduction in medium viscosity was observed as the fermentation progressed in all three starchy wastes. Madihah et al (2001) found, at high starch concentrations at batch level, that deposition of starch in the bottom of the bottle upon gel formation limits solvent production. They also concluded that the effect of gel formation and viscosity of the medium on solvent production was more conspicuous when there was no agitation due to decreased mass transfer in the medium. Mass transfer rate is contingent upon medium viscosity i.e. higher viscosities decrease diffusion rate and thus limit the activity of α-amylase and glucoamylase enzymes on starch molecules

(Madihah et al., 2001). It should also be noted that increased medium viscosity reduces cell motility which might have also contributed to the slowness of ABE fermentation in retrograded starchy media.

The control experiment with C. beijerinckii NCIMB 8052 in which glucose was used as substrate produced total ABE of 14.97 g/L which is comparable to inedible dough

(14.44 g/L), breading (14.80 g/L), and batter liquid (15.11 g/L) fermentations.

Interestingly, maximum cell counts of C. beijerinckii NCIMB 8052 grown in all the three starchy media was greater than that of the glucose control. It should be noted that the control had ABE yield of 0.43 g/g which is higher than all the starchy fermentation media tested.

C. acetobutylicum ATCC 824 produced total ABE of 14.62 g/L from glucose control which is higher than the breading medium (13.98 g/L) but lesser than the inedible dough medium (16.30 g/L) and the batter liquid medium (17.41 g/L). Only the batter

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liquid medium had the ABE yield (0.43 g/g) closer to the ABE yield of glucose control

(0.44 g/g).

The fermentation of starchy wastes by both microorganisms was faster than that of the glucose control. The presence of some mineral salts and differences in carbon and nitrogen contents of the starchy wastes media may have influenced the fermentation and

ABE production. However, C. acetobutylicum ATCC 824 adapted to the starchy media faster and better than C. beijerinckii NCIMB 8052.

3.4.3 Feasibility of using other starchy wastes

Other industrial starchy food processing wastes obtained in this study such as popcorn waste, donut dry mix, donut wet mix, potato waste products, chipped raw potato, landfill food wastes, and inedible pizza also can be directly fermented into butanol. All these wastes have significant amounts of starch contents and least amounts of mineral salts; therefore it is conceivable that they can easily be fermented to ABE.

3.5 Conclusions

3.5.1 Milk dust powder

Batch experiments have shown that from milk dust powder medium C. acetobutylicum ATCC 824 produced more solvents (10.25 g/L ABE) than C. beijerinckii

NCIMB 8052 (8.15 g/L ABE). Increased ABE concentration, yield, productivity and maximum lactose utilization were widely demonstrated previously by employing integrated ABE recovery system in a continuous fermentation mode. Hence, continuous or fed-batch fermentors integrated with product recovery could be an option for milk dust fermentation for large-scale industrial or commercial applications. The most noticeable 125

feature of this fermentation is the feasibility of direct use of this industrial dairy waste for butanol production.

3.5.2 Starchy food processing wastes

Among the three starchy wastes tested, batter liquid medium yielded maximum

ABE production by both microorganisms. Overall observation of the fermentation results

(such as maximum cell growth, total ABE produced, ABE yield and amount of substrate consumed) showed that C. acetobutylicum ATCC 824 was the best organism for the starchy wastes. Butanol production using these starchy wastes might have been limited due to butanol toxicity to the microbial cells given that butanol concentration in the medium reached 10-12 g/L which was comparable to the control experiment.

Retrogradation of the starchy media appeared to have delayed and restricted the ABE production by both microorganisms.

Direct utilization of some of the food processing wastes for butanol production was successfully demonstrated at batch level. Adopting these food processing wastes for butanol production by food industries would help meeting local energy demands and provide easy disposal of these wastes. Directing these wastes from the food processing facility to an on-site fermenter will cut down hauling and transportation costs involved in handling and disposal of these wastes. Also, this waste use will help mitigate environmental problems associated with landfill/compost of these wastes.

Although this study was done at a batch level, these results could encourage food processing industries to consider waste processing as a viable option for disposal and generation of additional revenue. The fermentation can be scaled-up to match the waste

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availability in the industries and would invariably employ continuous or fed-batch fermentation. Continuous or fed-batch fermentation with vigorous agitation and integrated butanol recovery techniques would help prevent starch gelatinization, enhance starch utilization, could handle higher concentrations of starch, and improve total ABE production from these wastes.

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Food wastes Starch content (% of dry weight)

Popcorn waste 52.14 ± 2.418 Inedible dough 60.38 ± 2.16 Donut dry mix 54.35 ± 2.85 Breadings 60.97 ± 3.42 Batter liquid 62.08 ± 4.68

Potato waste products 57.53 ± 2.69

Floating sludge 13.76 ± 3.87 Landfill food wastes 32.33 ± 4.74

Table 3.1 Amount of starch present in selected food wastes

Dairy wastes Lactose Glucose Galactose

Milk dust powder 424.16 ± 2.47 (g/kg) 0 0

Whey liquid waste 26.97 ± .21 (g/L) 0 0.43 ± 0.10

Table 3.2.Amount of sugars present in dairy wastes

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Parameters Control Inedible dough Breading Batter liquid Acetone (g/L) 3.12 ± 0.12 2.78 ± 0.34 2.78 ± 0.17 2.95 ± 0.52 Ethanol (g/L) 1.33 ± 0.48 2.44 ± 0.75 1.56 ± 0.18 2.46 ± 0.15 Butanol (g/L) 10.51 ± 0.65 9.26 ± 0.75 10.47 ± 0.59 10.00 ± 0.34 Total ABE (g/L) 14.97 ± 1.25 14.44 ± 1.37 14.80 ± 1.03 15.11 ± 1.30 Initial glucose (g/L) 60.86 ± 0.37 0.89 ± 0.13 0.96 ± 0.02 0.83 ± 0.11 Initial maltose (g/L) - 2.33 ± 0.04 2.46 ± 0.39 - Initial maltotriose (g/L) - 1.20 ± 0.02 0.30 ± 0.03 - Initial starch (g/L) - 49.91 ± 1.31 49.32 ± 0.39 50.55 ± 0.71 Final glucose (g/L) 26.24 ± 2.44 0.61 ± 0.64 4.45 ± 0.44 1.00 ± 0.05

129 Final maltose (g/L) - 4.05 ± 2.37 6.51 ± 0.61 1.93 ± 0.11 Final maltotriose (g/L) - 3.42 ± 2.15 8.31 ± 0.19 2.09 ± 0.28 Final Starch (g/L) - 10.46 ± 3.92 8.96 ± 0.42 9.91 ± 0.61 Total starch utilized (g) 34.62 ± 2.08a 39.45 ± 2.61 40.37 ± 0.58 40.64 ± 1.32 ABE yield (g/g of substrate) 0.43 ± 0.02 0.37 ± 0.03 0.36 ± 0.02 0.37 ± 0.03 ABE productivity (g/L/hour) 0.31 ± 0.02 0.24 ± 0.02 0.20 ± 0.01 0.31 ± 0.02 aTotal glucose utilized

Table 3.3 Performance and kinetic parameters of ABE production from starchy-food processing wastes using C. beijerinckii NCIMB 8052 after 72 h fermentation

Parameters Control Inedible dough Breading Batter liquid Acetone (g/L) 2.84 ± 0.82 3.18 ± 0.13 3.32 ± 0.61 4.02 ± 0.05 Ethanol (g/L) 1.39 ± 0.33 2.12 ± 0.10 0.92 ± 0.01 1.91 ± 0.05 Butanol (g/L) 10.65 ± 0.69 11.20 ± 0.31 9.74 ± 0.53 11.76 ± 0.28 Total ABE (g/L) 14.62 ± 1.74 16.30 ± 0.32 13.98 ± 1.14 17.41 ± 0.41 Initial glucose (g/L) 60.89 ± 0.16 0.81 ± 0.93 0.94 ± 0.11 0.64 ± 0.47 Initial maltose (g/L) - 2.74 ± 0.34 2.09 ± 0.32 0.04 ± 0.02 Initial maltotriose (g/L) - 0.92 ± 0.32 0.57 ± 0.09 - Initial starch (g/L) - 49.71 ± 0.91 49.92 ± 1.02 49.01 ± 0.17 Final glucose (g/L) 27.44 ± 1.92 1.19 ± 0.33 1.33 ± 0.40 0.99 ± 0.38

130 Final maltose (g/L) - 5.67 ± 0.51 4.81 ± 0.03 2.46 ± 0.59 Final maltotriose (g/L) - 4.23 ± 0.03 6.62 ± 0.34 2.74 ± 0.03 Final Starch (g/L) - 6.45 ± 0.07 7.96 ± 0.07 9.70 ± 0.02 Total starch utilized (g) 33.45 ± 1.77a 43.26 ± 0.68 42.00 ± 0.56 39.40 ± 0.14 ABE yield (g/g of substrate) 0.44 ± 0.01 0.38 ± 0.01 0.33 ± 0.03 0.43 ± 0.01 ABE productivity (g/L/hour) 0.25 ± 0.01 0.23 ± 0.01 0.19 ± 0.02 0.24 ± 0.01 aTotal glucose utilized

Table 3.4 Performance and kinetic parameters of ABE production from starchy-food processing wastes using C. acetobutylicum ATCC 824 after 72 h fermentation

Parameters Control Clostridium Control (Glucose) Clostridium (Glucose) acetobutylicum beijerinckii Clostridium ATCC 824 NCIMB 8052 Clostridium beijerinckii acetobutylicum NCIMB 8052 ATCC 824 Acetone (g/L) 2.84 ± 0.82 1.70 ±0.07 3.12 ± 0.12 1.21±0.11 Ethanol (g/L) 1.39 ± 0.33 1.45±0.21 1.33 ± 0.48 1.16±0.21 Butanol (g/L) 10.65 ± 0.69 7.25±0.32 10.51 ± 0.65 5.80±0.12 Total ABE (g/L) 14.62 ± 1.74 10.25±0.62 14.97 ± 1.25 8.15±0.79 Initial lactose (g/L) 60.89 ± 0.16a 49.94±1.61 60.86 ± 0.37a 49.08±1.26 Final lactose (g/L) 27.44 ± 1.92a 14.36±1.66 26.24 ± 2.44a 22.26±1.43 a a

131 Total lactose utilized (g) 33.45 ± 1.77 34.72±1.64 34.62 ± 2.08 27.67±0.70 ABE yield (g/g of substrate) 0.44 ± 0.01 0.30±0.03 0.43 ± 0.02 0.29±0.02 ABE productivity (g/L/hour) 0.25 ± 0.01 0.17±0.01 0.31 ± 0.02 0.13±0.01 aTotal glucose utilized

Table 3.5 Performance and kinetic parameters of ABE production from milk dust powder after 72 h fermentation

Types of waste Possible applications Almost 40% of cabbages are discarded as wastes and have largely been used for composting or animal feed applications (Nilnakara et al., 2009). Approximately 90 % of the dry weight of cabbage consists of one-third of fiber and two-third of low molecular weight carbohydrates mostly glucose, fructose, galactose, sucrose, raffinose, sedoheptulose and (Nilsson et al., 2006; Martinez-Castro et al., 2011). Also, cabbage has significant amounts of anticarcinogenic compounds, Cabbage trimmings antioxidants, phenolic compounds and vitamin C.

Production of dietary fiber (DF) powder rich in anti-carcinogenic compounds and anti- oxidants for food applications has been demonstrated earlier by several authors (Tanongkankit et al., 2011; Jongaroontaprangsee et al., 2007; Nilnakara et al., 2009; Nilsson et al., 2006; Tanongkankit et 132 al., 2010; Podsedek 2007).

Cabbage leaves and trimmings have high volatile solids which is the biodegradable part of the organic matter. Thus fermentation of these wastes is also an attractive option. Anaerobic fermentation for biogas production (RongHou et al., 2008; Velmurugan & Ramanujam, 2011 ), aerobic/anaerobic fermentation for industrial enzyme production (Chandrasekaran & Krishna, 1995; Das & Ghosh, 2009; Choi et al., 2002; Choi & Park, 2003), fermentation by lactic acid bacteria (Yoon et al., 2006) are the other applications. Effluents and digestate from the lactic acid fermentation using cabbage wastes could be used for animal feed or human food applications, since the lactobacillus is a well-known probiotic microorganism.

Table 3.6 Different types of wastes and their possible applications

Table 3.6 continued

Types of waste Possible applications Sauerkraut juice or brine is the waste liquid generated during the fermentation process of the shredded and salted cabbages. It contains organic compounds such as lactic acid, sugars and proteins (Fuchs et al., 2003). The low pH (3.76) of the sauerkraut juice explains the acidic nature of the juice. It also has high ash content (46.00 %) (Figure 3.1a) and potassium content (34.21 mg/g of dry matter) (Figure 3.3a). Biogas production through anaerobic fermentation has been demonstrated (Fuchs et al., 2003) and it can be used to utilize this waste stream. Sauerkraut juice contains high COD and BOD content (Fuchs et al., Sauerkraut Juice 2003; Shih & Hang, 1996), which is a suitable medium for various yeasts. Enzyme production (Sim & Hang, 1996; Hang & Woodams, 1990) and carotenoid production (Shih & Hang, 1996) from yeasts has also

133 been reported. Waste tea leaves have high energy content (20.35MJ/kg) and have 55% carbon (Figure 3.2d), which implies the presence of high carbohydrate content. Tea leaves mostly contain starch, cellulose, lignin and a small amount of protein (Sasaki et al., 2012). Anaerobic fermentation of leachate from tea wastes for biogas Waste tea leaves production has been reported (Goel et al., 2001). Production of ethanol from tea wastes is also an option. However, it requires enzymatic hydrolysis to break down the complex polymers into monomers to be consumed by the yeasts (Sasaki et al., 2012). Gasification, biodiesel production and production of other hydrocarbon fuel gases from tea wastes are stated in literature (Sasaki, Hashimoto, Asada, & Nakamura, 2012)

Table 3.6 continued

Types of waste Possible applications Except oregano spent (14.42 MJ/kg), all the spent wastes have energy content between 17.00 – 20.00 MJ/kg, while the organic content of all these spent wastes is around 40-45% (Figure 3.2d). Though Wet celery spent these substrates appear good for fermentation, they contain excess minerals such as phosphorus, potassium, Dry celery spent calcium, magnesium and sulphur (Figure 3.3d) which might severely inhibit fermentative microorganisms. Parsley spent Densification of these wastes into briquettes and pellets will further increase the energy content, and then it Oregano spent can be used to produce syngas through gasification. Syngas can be used in combustion engines to generate All spice spent electricity (Boerrigter & Rauch, 2006). The other option is pyrolysis (thermochemical decomposition) Fennel spent which can produce high value bio-oil, char and non-condensable fuel gases. The bio-oil can be used in

134 heating or electricity generation applications while the char and fuel gases can be used as low-grade fuels (Aho et al., 2008; Bridgewater et al., 1999). Biogas production through anaerobic fermentation might also work.

Table 3.6 continued Types of wastes Possible applications Though all these wastes came from different sources, they all have high amounts of fat and high energy content. It should be noted that these wastes were unable to be oven-dried since the fat melted when heated above 50-60°C; hence these wastes were freeze dried. The wastewater and wastewater solids contain excessive grease Uncooked meat and it could be considered alongside other fatty wastes. Cooked meat Meat and poultry industry occupies a significant portion in U.S agriculture sector. There were 6728 meat Inedible cheese and poultry slaughtering and processing plants in U.S and total production from these plants in 2011 was around Wastewater 41.87 million metric tonnes. This huge sector would produce thousands of tons of meat wastes per year45. Wastewater solids These wastes, rich in oils and fats can be used for biodiesel production in a small-scale industrial setup. Soybean-oil- Crude fats and oils from these wastes have to be extracted and filtered before being tranesterified into biodiesel. 135 shortening waste Crude glycerol is the byproduct of biodiesel production (Popescu & Ionel, 2011). Biodiesel blended with fossil Fryer siftings fuels can be used in engines (Phan & Phan, 2008) and whereas the crude glycerol can be used as an additive in Fried donut waste animal feed and as a feedstock for various chemicals (Yang et al., 2012). Biodiesel production from wastewater and wastewater solids is also feasible (Kargbo, 2010; Chakrabarti et al., 2008). Interestingly, the residual matter (after fat and oil extraction) can be composted/landfilled as a fertilizer to farmlands or it can be anaerobically fermented to produce biogas46 or can be used as animal feed or animal feed additive (Prior et al., 1988). These multi-utilization ways could be a viable option to reduce production costs in industries. Biogas production from animal fats and oils have also been demonstrated by several researchers (Wu et al., 2009; Hejnfelt & Angelidaki, 2009; Woon & Othman, 2011).

45 www.meatami.com/ht/d/sp/i/47465/pid/47465 46 http://www.thebioenergysite.com/news/8677/producing-biofuel-from-waste-crisps-and-pies

Table 3.6 continued

Types of waste Possible applications These wastes, which are co-products of anaerobic digestion, are widely been used for land application as fertilizer. The bottom sludge has 20.93 MJ/kg and 46.71 % carbon Digester bottom whereas the floating sludge has an energy value of 28.84 MJ/kg and 60.06% carbon (Figure sludge 3.2c). Thermal processes such as gasification or pyrolysis could help improve utilization of Digester floating these wastes. In particular, pyrolysis is an attractive option since the energy content of the sludge wastes are recovered as bio-oil and the sludge nutrients are recovered as char (Bridle & Pritchard, 2004; Hospido et al., 2005; Stammbach et al., 1989; Miikki et al., 1999). Wastewater treatment plant (WWTP) sludge is a combination of microbes and

136 biomass that accrues during wastewater treatment. The sludge has 3.77% total solids (Figure 3.1c), energy content of 21.32 MJ/kg and 46.92% carbon. Bellisio liquid wastes contain total solids of 7.94 %, energy value of 23.80 MJ/kg and 43.53% carbon (Figure WWTP sludge 3.2c). Bellisio liquid wastes Anaerobic digestion of these wastes for biogas production followed by pyrolysis of the digested/composted sludge can increase the energy recovery from these wastes (Cao & Pawlowski, 2012; Cao & Shan, 2011). Land application of the digested sludge could pollute soil if the sludge contains heavy metals, and possibly some pathogenic microorganisms and toxic compounds (Rulkens, 2007). Particularly, WWTP sludge has 18.06 mg of phosphorus and 0.42 mg of strontium per g of dry matter (Figure 3.4c)

136

Table 3.6 continued

Types of waste Possible applications These wastes are rich in starch and have only small amounts of minerals; hence these wastes are ideal for Landfill food butanol production (Table 3.1 & Figure 3.3d, 3.4d). Direct utilization of starchy food processing wastes for butanol wastes production has already been demonstrated in this thesis. Fermentation of these wastes by yeasts for ethanol Donut dry mix production could be an alternative option; however, enzymatic hydrolysis of starch is required prior to fermentation. Donut wet mix Another interesting application could be as a substrate for biodegradable starch-based plastic wares and Inedible pizza dinner wares. Currently most of the starch-based plastics are made from corn starch or cereal starch, which will likely Popcorn waste increase the food-fuel conflicts. Therefore, diverting starchy food processing wastes for bio-plastic manufacturing would reduce the dependence on corn-starch as well as production cost.

137 Global potato production in 2010 was 324.18 million metric tonnes of which 18.34 million metric tonnes were produced in USA47 (FAO, 2011). Potato wastes have high water (80-85%) and carbohydrates (12-13%) Potato waste content. Potato pulp has starch as the major carbohydrate that can be fermented to produce butanol as stated earlier. products The one common application of potato waste is as animal feed (Tawila et al., 2008; Radunz et al., 2003). A new Chipped raw application of potato pulp is extraction of rhamnogalacturonan I.(a pectic polysaccharide) to use as a functional food potato ingredients (Byg et al., 2012). Raw potato Potato peel is an important waste produced in huge amounts in potato processing industries. It has 52.14% starch (on dry weight basis), and have considerable amounts of cellulose, hemicellulose and lignin (Arapoglou et al., 2010). Direct fermentation of potato peel for butanol production is feasible. Other possible application of potato peel is extraction of antioxidants (Schieber & Saldana, 2009; Chang, 2011).

47 faostat.fao.org/site/339/default.aspx

Table 3.6 continued

Types of waste Possible applications Anaerobic fermentation of vegetable processing wastes for biogas or hydrogen production has Coriander widely been in practice worldwide for the past few decades (Sagagi et al., 2009; Velmurugan & Minced pepper Ramanujam, 2011; Bouallagui et al.,2003; Thompson 2008). Raw pepper Upon hydrolysis these wastes can be used for ethanol (Patle & Lal, 2007; Qazi, 2005) and Onion peel butanol production. However, the vegetable waste hydrolyzate has to be concentrated to have the amount Beans of sugars required for fermentation (Claassen et al., 2000). Canned broccoli Another applications are extraction of antioxidants from coriander (Guerra et al., 2005; waste Wangensteen et al., 2004 ; Rajeshwari et al., 2012) and broccoli (kaur et al., 2007), flavonoids from onion 138 peel (Martino & Guyer, 2007; Kefalas & Makris, 2008), pigments from pepper (Richins et al., 2010; Valle et al., 2003). The cost involved in these extraction processes have to be taken into consideration to decide the efficacy of this process. Biological or thermochemical conversion of the residual matters into biofuels could improve the economics of the extraction processes.

Table 3.6 continued

Type of waste Possible applications

World tomato production in 2010 was 151.70 million metric tonnes in which USA Tomato pulp produced12.85 million metric tonnes (FAO, 2011)48. Globally more than 35 million metric tonnes Canned tomato juice waste of tomatoes are processed in a year49. Estimated 11 million metric tonnes of tomato wastes were produced in 2007 of which tomato pulp alone amounted to 4 million metric tonnes50. Both tomato pulp and peel have high moisture content (80-98%) and thus necessitates drying to prevent fouling and to preserve these wastes for alternate applications. The composition of tomato pulp (in dry weight basis) is: protein (17-22%), fat (10-15%), crude fiber (33-57%), 25.73% total sugars, 7.55% pectins18 (Mirzaei-Aghsaghali et al., 2011; Del 139 valle et al., 2006). The most well-known-applications of tomato pulp are its use as animal feed ingredient (Weiss et al., 1997; Caluya et al., 2000; Yuangklang et al., 2010, Mirzaei-Aghsaghali et al., 2011) and a substrate for biogas production (Ulusoy et al., 2009). The other applications of tomato pulp are: tomato pulp powder as thickening agent in tomato ketchup (Farahnaky et al., 2008), extraction of phytochemicals (polyphenols, carotenoids, antioxidants) for food applications (Savatovic et al., 2010; Kalogeropoulos et al., 2012).

48 faostat.fao.org/site/339/default.aspx 49 www.tomatonews.com/resources.html 50 www.feedipedia.org/node/689

Table 3.6 continued

Types of waste Possible applications Tomato peel and seed comprises 1-4% (w/w) of total tomato processed for products (Kalogeropoulos et al., 2012). Fiber (46.10%) (Lazos & Kalathenos, 1988), protein (10-20%) Tomato peel (Knoblich et al., 2005) on dry matter basis, and pectin are the major compounds of tomato peel which has reflected in the energy content (22.95 MJ/kg) (Figure 3.2a). Owing to the high fiber content, production of micro and nano fibers for food applications (Kocak, 2010) and fillers for rubber manufacturing processes are new applications of tomato peel. Apart from fiber, tomato peel also contains considerable amount of lycopene (an antioxidant) 800-1200 µg/ g of dry matter (Knoblich et al., 2005; Topal et al., 2006) and cutin (a wax-like polymer) (Osman et al., 1999). Extraction of lycopene (Lavecchia & Zuorro, 2008; Knoblich et al., 140 2005; Naviglio et al., 2008; Nobre et al., 2012), biodegradable plastics and biofilm production51,52 and biogas production are some other applications of tomato peels (Atem et al., 2010; Dinuccio et al., 2010).

51 http://blog.productosecologicossinintermediarios.es/2012/05/a-biodegradable-plastic-obtained-from-tomato-skin/ 52 http://www.bvsde.paho.org/bvsacd/iswa2005/food.pdf

Table 3.6 continued

Type of waste Possible applications

Calcium sulfate dihydrate (CaSO4 .2H2O) (also known as gypsum) which appears as a white solid Calcium sulfate when dried in an oven can be used as a fertilizer in agricultural lands. Heating removes the water molecules

stream and then it can be turned into anhydrous calcium sulfate (CaSO4), which has enormous applications in various industries. Some noticeable applications include filler material in paper and rubber polymer, and cement manufacturing. EPA defines coal ash as remnants gathered during burning of coal in boilers. This ash is non- Coal boiler ash combustible (might contain little carbon from incomplete combustion of coal), and contains 80-90% mineral matter (accrued from coal) mostly silicon, aluminum, iron and calcium. Combustion or fermentation of this

141 waste is not possible. Few well-known applications of coal ash are as a building material for concrete/cement blocks and filler material for paints, plastics and rubbers53. Whey liquid has lactose as the primary sugar component. Production of butanol from whey liquid Whey liquid waste is a well-established research area, but the whey liquid we obtained has only 26.97 g/L lactose (Table 3.2) which only makes half the amount of sugar required for butanol production. Concentrating the whey liquid would be an option to make it suitable for butanol production. The animal feed waste is the left-over or wasted feed materials gathered in the facility. Application Animal feed waste of whey liquid and animal feed wastes as cattle feed is a good option (which the company currently adopted to deal with these wastes). Biological conversion of these wastes for biogas production is another choice to consider.

53 http://www.epa.gov/radiation/tenorm/coalandcoalash.html

(a) Percent Total solids, Ash, and pH of different vegetable wastes

60

40

20 142

0

Vegetable wastes %TS %Ash pH

Figure 3.1 Percent total solids, ash and pH (a) vegetable wastes, (b) fat-rich industrial wastes, (c) industrial sludge wastes, (d) starchy and other industrial wastes

Figure 3.1 continued

(b)

Percent Total solids, Ash and pH of fat-rich industrial wastes

100

75 143 50

25

0 Uncooked inedible Cooked inedible Cheese inedible Fryer siftings Fried donut waste Soybean oil meat meat shortening waste

Fat-rich industrial wastes % TS % Ash pH

Figure 3.1 continued

Percent Total solids, Ash and pH of different industrial sludge wastes (c) 36

24 144

12

0 Con Agra Con Agra Dannon Dannon WWTP Bellisio liquid Bellisio Bellisio Bellisio wastewater wastewater whey liquid animal feed sludge waste bottom floating landfill food solids sludge sludge waste Industrial sludge wastes

% TS % Ash pH

Figure 3.1 continued

(d) Percent Total solids, Ash, and pH of starchy and other industrial wastes

100

75

50 145

25

0

Starchy and other industrial wastes % TS % Ash pH

Mean calorific value, percent carbon, nitrogen and C/N ratio of different (a) vegetable wastes

60

50

40

30

20 146 10

0

Vegetable wastes

Calorific value (MJ/kg) % Carbon % Nitrogen C/N ratio *Mean ± 1.96 MSE

Figure 3.2 Mean calorific value, percent carbon, nitrogen and C/N ratio (a) vegetable wastes, (b) fat-rich industrial wastes, (c) industrial sludge wastes, (d) starchy and other industrial wastes

Figure 3.2 continued

(b) Mean calorific value, percent carbon, nitrogen and C/N ratio of fat-rich industrial wastes

80

60

147 40

20

0 Cheese inedible Uncooked inedible Cooked inedible Fried donut waste Fryer siftings Soybean wastes meat meat Fat-rich industrial wastes

* Soybean oil Shortening wastes : Mean % Nitrogen: 0.15 ± 0.07; Mean C/N: 521.61 ± 0.61 Calorific value (MJ/kg) % Carbon % Nitrogen C/N ratio

Figure 3.2 continued

(c) Mean calorific value, percent carbon, nitrogen and C/N ratio of different industrial sludge wastes

75

60

45 148 30

15

0 Con Agra Con Agra Dannon whey Dannon Dannon Bellisio liquid Bellisio Bellisio Bellisio waste water waste water liquid animal feed WWTP waste bottom floating landfill food Solids sludge sludge sludge waste

Industrial sludge wastes *Mean ± 1.96 MSE Calorific value (MJ/kg) % Carbon % Nitrogen C/N ratio

Figure 3.2 continued

(d) Mean calorific value, percent carbon, nitrogen and C/N ratio of starchy and other industrial wastes

60

40 149

20

0

Starchy and other industrial wastes *Cardboard waste: Mean C/N: 97.39 ± 2.40 Calorific value (MJ/kg) % Carbon % Nitrogen C/N ratio

(a) Concentration of major elements present in different vegetable wastes

100

75

50

25 150 Amount matter) dry of (mg/g Amount

0

Vegetable wastes

P K Ca Mg S Al B Cu Fe Mn Mo Na Zn

Figure 3.3 Concentration of major elements present (a) vegetable wastes, (b) fat-rich industrial wastes, (c) industrial sludge wastes, (d) starchy and other industrial wastes

Figure 3.3 continued

(b) Concentration of major elements present in fat-rich industrial wastes

45

30 151

15 Amount matter) dry of (mg/g Amount

0 Cheese inedible Uncooked inedible Cooked inedible Fried donut waste Fryer Siftings Soybean oil meat meat shortening wastes

Fat-rich industrial wastes

P K Ca Mg S Al B Cu Fe Mn Mo Na Zn

Figure 3.3 continued

(c) Concentration of major elements present in different industrial sludge wastes

80

60

40 152

20 Amount matter) dry of (mg/g Amount

0 Con Agra Dannon Bellisio liquid Bellisio Bellisio Bellisio Whey liquid Dannon Waste water Waste water WWTP waste bottom Digester Landfill food waste Animal feed Solids sludge sludge floating waste sludge Industrial sludge wastes

P K Ca Mg S Al B Cu Fe Mn Mo Na Zn

Figure 3.3 continued

(d) Concentration of major elements present in starchy and other industrial wastes

80

60

40 153 20 Amount matter) dry of (mg/g Amount

0

Starchy and other industrial wastes

P K Ca Mg S Al B Cu Fe Mn Mo Na Zn

(a) Concentration of minor elements present in different vegetable wastes

800

600

400

200 Amount matter) dry of (µg/g Amount

154 0

Vegetable wastes

As Ba Be Cd Co Cr Li Ni Pb Sb Se Si Sr V Tl

Figure 3.4 Concentration of minor elements present (a) vegetable wastes, (b) fat-rich industrial wastes, (c) industrial sludge wastes, (d) starchy and other industrial wastes

Figure 3.4 continued

(b) Concentration of minor elements present in fat-rich industrial wastes

300

150 155 Amount matter) dry of (µg/g Amount

0 Cheese inedible Uncooked inedible Cooked inedible Fried donut waste Fryer siftings Soybean oil meat meat shortening waste

Fat-rich industrial wastes

As Ba Be Cd Co Cr Li Ni Pb Sb Se Si Sr V Tl

Figure 3.4 continued

(c) Concentration of minor elements present in different industrial sludge wastes

1800

1200 156

600 Amount matter) dry of (µg/g Amount

0 Con Agra Dannon Bellisio liquid Bellisio bottomBellisio floating Bellisio landfill Dannon whey Con Agra waste water WWTP sludge waste sludge sludge food waste liquid wastewater Solids

Industrial sludge wastes

As Ba Be Cd Co Cr Li Ni Pb Sb Se Si Sr V Tl

Figure 3.4 continued

(d) Concentration of minor elements present in starchy and other industrial wastes

1800

1200

157 600 Amount matter) dry of (µg/g Amount 0

Starchy and other industrial wastes

As Ba Be Cd Co Cr Li Ni Pb Sb Se Si Sr V Tl

(a) 12

8 Butanol(g/L)

4

0 0 24 48 72 Fermentation time (h)

20 Glucose control (b) Inedible dough Breading 16 Batter liquid Milk dust powder

12

8 Total (g/L) ABE

4

0 0 24 48 72 Fermentation time (h)

Figure 3.5 Fermentation of food processing wastes using C. beijerinckii NCIMB 8052 (a) butanol production, (b) total ABE production

158

5 (a)

4

3

2 Acetone(g/L)

1

0 0 24 48 72 Fermentation time (h)

7.0 (b) Glucose control Inedible dough 6.5 Breading Batter liquid Milk dust powder 6.0 pH

5.5

5.0

4.5 0 24 48 72 Fermentation time (h)

Figure 3.6 Fermentation of food processing wastes using C. beijerinckii NCIMB 8052 (a) acetone production, (b) change in pH

159

6

(a)

4

Acetic(g/L) acid 2

0 0 24 48 72 Fermentation time (h)

4 Glucose control Inedible dough (b) Breading 3 Batter liquid Milk dust powder

2 Butyric acid (g/L) 1

0 0 24 48 72 Fermentation time (h)

Figure 3.7 Fermentation of food processing wastes using C. beijerinckii NCIMB 8052 (a) acetic acid production, (b) butyric acid production

160

16

(a)

12

8 Butanol(g/L)

4

0 0 24 48 72

Fermentation time (h)

24 Glucose control Inedible dough (b) 20 Breading Batter liquid Milk dust powder 16

12

Total ABETotal (g/L) 8

4

0 0 24 48 72

Fermentation time (h)

Figure 3.8 Fermentation of food processing wastes using C. acetobutylicum ATCC 824 (a) butanol production, (b) total ABE production

161

5 (a) 4

3

Acetone(g/L) 2

1

0 0 24 48 72 Fermentation time (h)

7.0 Glucose control Inedible dough (b) 6.5 Breading Batter liquid Milk Dust Powder

6.0 pH

5.5

5.0

4.5 0 24 48 72

Fermentation time (h) Figure 3.9 Fermentation of food processing wastes using C. acetobutylicum ATCC 824 (a) acetone production, (b) change in pH 162

5

(a) 4

3

2 Acetic(g/L) acid

1

0 0 24 48 72

Fermentation time (h)

3 Glucose control Inedible dough Breading (b) Batter liquid Milk dust powder 2

1 Butyric acid (g/L) Butyric

0 0 24 48 72

Fermentation time (h)

Figure 3.10 Fermentation of food processing wastes using C. acetobutylicum ATCC 824 (a) acetic acid production, (b) butyric acid production

163

C. beijiernckii NCIMB 8052 C. acetobutylicum ATCC 824 60 50 40 30 20

Lactose (g/L) Lactose 10 0 0 24 48 60 72

Fermentation time (h)

Figure 3.11 Milk dust powder consumption by C. beijerinckii NCIMB 8052 and C. acetobutylicum ATCC 824

1e+11

1e+10

1e+9

CFU/ml 1e+8

Glucose control Inedbile dough 1e+7 Breading Batter liquid Milk dust powder 1e+6 0 24 48 72 Fermentation time (h)

Figure 3.12 Cell growth of C. beijerinckii NCIMB 8052 in food processing waste media 164

1e+12

1e+11

1e+10

CFU/ml 1e+9

Glucose control 1e+8 Inedible dough Breading Batter liquid Milk dust powder 1e+7 0 24 48 72

Fermentation time (h)

Figure 3.13 Cell growth of C. acetobutylicum ATCC 824 in food processing waste media

Coagulated milk dust Stone-like formation Agitated milk dust medium in milk dust medium medium

Figure 3.14 Structural changes of milk dust medium

165

CHAPTER 4: BIOBUTANOL PRODUCTION FROM INULIN-RICH

BIOMASS

4.1 Introduction

Biofuel production from traditional annual crops such as soybean, corn and sorghum are often blamed for increasing food prices, negative impacts on existing land and water resources, and declining biodiversity (Dale et al., 2010). The growing demand for biofuels impels the research community to look into additional crops and wastes that could fulfill multiple societal needs for food, fuel, feed and fiber. This challenge could be met by the development of new and multipurpose industrial crops that offer valuable products and energy-rich biomass, and are able to be grown on marginal lands. To ensure sufficient food and cattle feed production and to mitigate their price rise, biofuel production from non-food related industrial crops is preferred. Two non-food industrial crops, which are gaining prominence as alternate rubber producing crops in the U.S., are guayule (Parthenium argentatum Gray) and Buckeye gold, also known as Kazak dandelion (Taraxacum kok-saghyz), hereafter referred to as TKS. Efforts to find domestic sources of natural rubber in the U.S. during World War II, when natural rubber supplies (from Hevea brasiliensis) from Asian countries were cut-off, resulted in concerted research, planting and development of both of these plants. However, the

166

aftermath of the war saw normalization of rubber supplies and the resurgence of Hevea rubber thwarted all commercial-scale alternative rubber production in the U.S54.

Increasing global demand for natural rubber, and problems associated with Hevea rubber cultivation, such as prolonged and complex breeding cycles, labor-intensive rubber extraction processes and latex allergy have prompted research into guayule and

TKS (van Beilen & Poirier, 2007). Guayule is a perennial shrub, but widely seen as a biannual or multiannual rubber producing plant which stores rubber in the cells of bark and woody tissues55. Guayule produces hypoallergenic rubber with molecular weight comparable to Hevea and is suitable for medical applications (Cornish, 1996). On the other hand, TKS is an annual crop which stores rubber in its roots. Rubber extracted from TKS has excellent properties and is considered suitable for non-medical applications such as manufacturing of automobile and aircraft tires (van Beilen &

Priorier, 2007). Guayule cultivation has been commercialized whereas TKS is still pre- commercial and being evaluated for high value markets. Although current research is focused primarily on molecular and breeding techniques (Collins-Silva et al., 2012;

Schmidt et al., 2010; Kirschner et al., 2012) to improve rubber yield, the large amounts byproduct biomass produced from these plants cannot be ignored. Guayule produces non-rubber biomass of 13 tons/ha/year (van Beilen & Priorier, 2007) or more, mostly in the form of resin and residual bagasse. The resin is a complex mixture of sesquiterpene ethers, triterpenoids, and fatty acid triglycerides and has found applications as plastic

54 http://www.kultevat.com/industrial.html 55 http://www.eu-pearls.eu/UK/Background/Guayule/

167

binder, wood preservative or rubber additive (Cornish et al., 2007). The guayule bagasse primarily contains cellulose, hemicellulose, lignin, resin and some proteins and lipids

(Boateng et al., 2009). This solid biomass has been used in plastic composite materials

(Nakayama, 2004), and for soil enrichment (Nakayama, 2005). Thermochemical conversion of guayule biomass into bio-oil (~30 MJ/kg) and other useful chemical compounds (Boateng et al., 2009, 2010) and ethanol production (Srinivasan, 2010) has been reported. However, the possibility of using the residual aqueous extract (remaining liquid after rubber extraction) for biofuel production has not been previously investigated.

TKS roots contain 2-20% rubber and 25-60% inulin on a dry weight basis

(Buranov & Elmuradov, 2010; Schutz et al., 2006; van Beilen & Poirier, 2007). The residual root bagasse and leaves of TKS could also be used for cellulosic biofuel production. Currently two different rubber extraction processes are employed to extract rubber from TKS roots, namely latex extraction (Cornish, 1996) using aqueous base at room temperature and solid rubber extraction using hot water at 70-95°C (Eskew, 1946).

The overall schematic representation of TKS processing, rubber extraction and biomass production is presented in Fig. 4.1

Owing to the differences in extraction temperatures and solvents involved, the liquid extract obtained from these two methods gives different amounts of inulin with different molecular weights and degrees of polymerization (DP). It should be noted that inulin is only 10% soluble in water at room temperature but highly soluble in water between 50 to 100°C (Franck, 2002; Ranawana, 2008). Research on butanol production from hydrolyzed inulin extract was reported earlier by Marchal et al (1985) while butanol

168

from unhydrolyzed inulin was reported by Oiwa et al (1987). Though Clostridia can consume a different range of susbtrates, the capability of Clostridium sp. to hydrolyze polysaccharides differs significantly from strain to strain (Montoya et al., 2000). Hence, this study was aimed at finding a suitable butanol producing strain for the hydrolyzed and unhydrolyzed inulin medium. Chicory extract, which has similar molecular characterisitics of TKS inulin, was tested as a model substrate and its fermentation performance was compared to TKS inulin fermentation. In addition to that, guayule extract also was assessed for its suitability for butanol fermentation.

Inulin was hydrolyzed enzymatically using commerically available endo-inulinase enzyme (Novozyme 960). Production of exo-inulinase from Kluyveromyces marxianus

ATCC 52466 and endoinulinase from Kluyveromyces marxianus ATCC 16045 also was attempted.

Since TKS produces biomass in addition to rubber, commercialization of this crop is largely contingent upon finding economic uses and/or cost-effective disposal methods for this residual biomass. Production of butanol has a wide variety of potential applications as an alternate fuel, industrial solvent and chemical intermediate for the synthesis of many other products. Although this study is focused mainly on butanol production from extracted inulin, it should be noted that butanol also could potentially be produced from bagasse and leafy biomass of TKS. Scale up of the fermentation of TKS inulin could improve the economics and help commercialize TKS as a viable natural rubber producing crop in USA. Also, in-situ fermentation of inulin extract during rubber extraction could ameliorate process efficiencies by cutting down costs and time involved

169

in transporting the biomass and could also provide solvents much needed by other industries.

4.2 Materials and Methods

4.2.1 Microorganisms and culture maintenance

Stocks of C. beijerinckii NCIMB 8052, C. beijerinckii NRRL B-592, C. acetobutylicum ATCC 824 and C. saccharobutylicum P262 were routinely preserved as spore stock suspensions in sterile double distilled water at 4°C. Spores (200 µL) of all the organisms, except C. saccharobutylicum P262, were heat shocked at 75°C for 10 minutes, followed by cooling in ice for 3 minutes. C. saccharobutylicum P262 spores

(200 µL) were heat shocked for 3 minutes at 70°C and then cooled in ice for 1 minute

(Ezeji & Blascheck, 2008). The heat shocked spores were inoculated into 10 mL of TGY

(tryptone-glucose-yeast extract) medium and incubated anaerobically for 12-14 h at

35±1°C. Spores of C. saccharobutylicum P262 were incubated anaerobically for 16-18 h

(Qureshi & Maddox, 2005). Upon attaining the optimum optical density (0.9-1.1,

OD600nm), 8 mL of actively grown culture was transferred into 92 mL of anoxic presterilized TGY medium and incubated for another 3-4 h (OD of 0.9-1.1 at 600 nm) before they were transferred into solvent production medium. Loosely capped pyrex bottles containing presterilized TGY medium were placed inside the anaerobic chamber

14-16 h prior to fermentation to induce anaerobiosis. The anaerobic chamber (Coy

Laboratory Products Inc., Ann Arbor, MI) had a modified atmosphere of 82% N2, 15%

CO2, and 3% H2 (Richmond et al., 2012).

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4.2.2 Inulin extracts from different sources

Inulin extract from guayule, TKS and Chicory were analyzed for their sugar content to assess their suitability for butanol fermentation. The amount of sugars present in the raw extract is presented in Table 4.1. It is self-evident that guayule extract, which had merely 2.56 g/L of inulin effectively, cannot be used for butanol fermentation, unless it is concentrated to increase the sugar content. Similarly, TKS extracts obtained from the latex process had a limited amount of sugars (~12 g/L inulin) which is not sufficient for ABE production. Media made from extracts with low concentrations of sugars are liable to substrate exhaustion during fermentation, which limits the ability of Clostridia to produce solvents. Though both guayule and TKS extract (Latex) could be used if concentrated to bring the sugar concentration to ~50 g/L.

TKS inulin and chicory inulin extracts (syrup) obtained through the Eskew process contained enough sugars (~50 g/L) for conversion to butanol. In the Eskew process, dried and roller milled TKS roots were heated in extraction solvent at ~70°C and each extraction was carried out for 30 minutes. A total of 10 extractions were carried out, each time with fresh extraction solvent, to make sure that all the leachable carbohydrates from the roots were extracted. Between 0.40 to 0.50 g of inulin per gram of TKS roots was obtained through this process (data not shown). The first three extractions (which contained ~28-30 g/L inulin) were pooled and kept in a hot air oven (~45°C) for 24 -36 h to evaporate off excess water to a final inulin concentration of ~40 g/L, to make it suitable for ABE fermentation.

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The extraction solvent was a mixture of water and 0.1% Na2CO3. The extraction process had a solvent to root ratio of 6:1 (v/w). The TKS extract used in this study was obtained from TKS inulin-extraction studies conducted in another lab (Dr. Fred Michel’s research group, OARDC, Wooster).

Chicory inulin extract was obtained from OSU-PENRA Eskew process pilot plant located in Wooster, Ohio. The pilot plant was built to process locally cultivated TKS roots in large quantities, and chicory roots were used for a trial run. Leaching of inulin from the roots was done via an 8-stage counter-current extraction method.

Both raw TKS and Chicory extract contained appreciable amounts of short- oligomers and monomers. Although, native inulin (raw inulin) extract naturally comes with free monomers (due to inulin metabolism in plants), the increased amount of monomers presence in the inulin extracts can be attributed to inulin extraction temperature (~90°C) and additional concentration step. Temperature and pH are significant factors affecting inulin hydrolysis (Franck, 2002; Ranawana, 2008; Barclay et al., 2010). It is noteworthy to mention that the original pH of the TKS extract was 4.78 whereas it was 4.49 for chicory extract. These low pH values might be due to possible fermentation of inulin extract during storage. The fatty acid production during fermentation might have reduced the pH of the extract (It should be noted that the pH of extract solvent used was in the range of 9-10). Also, at higher temperatures inulin is more leachable from the roots than at room temperature. Multiple extractions steps and higher extraction temperatures were likely the reasons for the higher final sugar concentration in the TKS Eskew extract compared to the TKS Latex extract (Table 4.1).

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4.2.3 Media preparation

In this study, the substrate specificity of the organisms tested was identified. Both unhydrolyzed (raw) inulin and enzymatically-hydrolyzed inulin extract media were used for fermentation. The overview of inulin fermentation is presented in Figure 4.2.

Preliminary experiments were aimed at identifying suitable organisms that could consume and produce ABE from inulin and fructose medium. Since literature available on inulin and inulin-derived oligomer consumption by Clostridia is scarce, exploration of these substrates for butanol production was necessary.

4.2.3.1 Unhydrolyzed inulin media

TKS Eskew extract and chicory extract media were prepared by adding 1 g/L yeast extract. Commercially-available pure inulin (AlfaAesar) was used to prepare inulin

P2 medium (~55 g/L inulin and 1 g/L yeast extract) which was used as control for unhydrolyzed inulin fermentation.

4.2.3.2 Enzymatically-hydrolyzed inulin media

Both TKS and Chicory inulin extract were hydrolyzed using endo-inulinase

(Novozyme 960). The hydrolyzates in addition to 1g/L yeast extract were used as fermentation media. Pure fructose P2 medium (60 g/L fructose and 1 g/L yeast extract) was used as control medium for hydrolyzed inulin fermentation, since fructose is the major hydrolysis product of inulin. The appearance of TKS Eskew and Chicory inulin medium is depicted in Figure 4.3.

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Pyrex screw capped bottles (150 mL) containing 100 mL of fermentation medium were used throughout this studies. Bottles with 91 mL of fermentation media were sterilized for 15 minutes at 121°C and cooled to 40°C before placing them inside the anaerobic chamber at 35±1°C for 14-16 hours. This was followed by addition of filter- sterilized P2 stock vitamin (0.1 g/L para-amino-benzoic acid; 0.1 g/L thiamine; 0.001 g/L biotin), buffer (50 g/L KH2PO4; 50 g/L K2HPO4; 220 g/L ammonium acetate) and mineral (20 g/L MgSO4.7H2O; 1 g/L MnSO4.H2O; 1 g/L FeSO4.7H2O; 1 g/L NaCl) solutions of 1 mL each to the fermentation medium. Actively grown cells from the TGY medium were used as inoculum and the fermentation medium was inoculated with 6%

(v/v) of inoculum. In summary, 100 mL of fermentation medium consists of 91 mL of extract, 3 mL of P2 medium stock solutions and 6 mL of inoculum.

The initial pH of all fermentation media were adjusted to 6.50 after autoclaving using 3M KOH and temperature was maintained at 35±1°C, no agitation or pH control was employed and all fermentations were done in triplicate. During the course of each fermentation, 3 mL samples were taken from each flask every 12 h to determine cell growth and to analyze residual sugars, ABE and acid production.

4.2.4 Calcium carbonate as a component of the fermentation medium.

Though Clostridia can consume fructose as a sole carbon source, the fermentation rate was slower and solvent production lower, than on glucose as the sole carbon source

(Chen et al., 2010), possibly due to less efficient fructose transport into the cells.

Effects of several salts on butanol production have been reported. Addition of salts such as NH4CH3COOH (Gu et al., 2009; Kasap, 2002; Welsh et al., 1986),

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(NH4)2SO4, and carbonates (Rangaswamy & Isar, 2012) were found to influence butanol production. Han et al (2012) tested the effects of various carbonate salts, Na2CO3,

K2CO3, (NH4)2CO3, NaHCO3, NH4HCO3, and CaCO3 in P2 medium using C. beijerinckii

NCIMB 8052 and found that CaCO3 supplemented P2 medium produced more solvents than the other carbonate salts tested. Addition of certain salts could influence cell growth and solvent production, therefore, addition of calcium carbonate in the medium was assessed.

Preliminary experiments, after adding CaCO3, showed marked improvement in solvent production and fructose consumption in all of the strains. Richmond et al (2011) found that the effect of CaCO3 on Clostridium species is non-selective (influences ABE fermentation irrespective Clostridia Sp.) in action. Further they found that 4 g/L CaCO3 was an optimum concentration for ABE fermentation and any further increase in CaCO3 concentration did not show any effects on solvent production. The solubility of CaCO3 in water is very limited and contingent upon pH (CaCO3 has solubility of approximately 2 g/L at pH 6.0). A decrease in medium pH elevates dissolution of CaCO3 with subsequent proton consumption and thus increases medium alkalinity. Thus, addition of 4 g/L

CaCO3 improves buffering capacity to the decreasing pH and limits acidification during

ABE fermentation (Han et al., 2012).

Addition of CaCO3 also enhances cell growth, reduces butanol toxicity in the medium and increases substrate utilization. The dramatic improvements in overall performance of fermentation were because of elevated expressions of several proteins of the Clostridium species in the presence of CaCO3 (Richmond et al., 2011). Hence, 4 g/L

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CaCO3 was also included as one of the basal medium components and all fermentation media used in this study contained 4g/L CaCO3.

4.2.5 Enzymatic hydrolysis of Inulin

Inulin was hydrolyzed using endoinulinase (Novozyme 960; I6285 Sigma; CAS

#: 9025-67-6) from Aspergillus niger. The enzyme cleaves beta 2→1 fructosyl linkages and yields short oligomers and monomers upon hydrolysis of inulin. According to the enzyme supplier’s description, this enzyme has inulinase activity of 286 IU/g (329

INU/ml; Density 1.15 g/ml), IU refers to inulinase units. The optimum temperature of the enzyme was between 55-60°C and the optimum pH was in the range of 4.60-5.20.

Reportedly, fructose formation was greater at temperatures above 55°C. The effect of pH on endoinulinase activity is presented in Figure 4.4. Therefore, in this study, hydrolysis of TKS and chicory inulin extracts was carried out at pH 5.00 and a temperature of 60°C.

Since the hydrolyzate had to be used for butanol fermentation (ideal medium pH is 6.50), the hydrolysis was done at pH 5.00 to limit the addition of base during pH adjustment.

The enzyme comes in a mixture of aqueous glycerol solution.

The amount of enzyme used and substrate available for hydrolysis determine the rate of hydrolysis and depolymerization of inulin (Ricca et al., 2009a & 2009b; Michel-

Cuello et al 2012). Therefore, the optimum amount of endoinulinase required to obtain maximum amount of hydrolysis products, preferably monomers, was determined in this study. To evaluate the optimum amount of endoinulinase, hydrolysis with 50 IU, 100 IU, and 150 IU per gram of inulin was performed.

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The enzymatic hydrolysis experiments were carried out in 250 mL screw-capped bottles (Fisherbrand) placed in an incubator shaker (New Brunswick, Innova 4300) with agitation (250 rpm) and controlled temperature (60°C). One replicate of each experimental condition was used and commercial inulin was used as model substrate to compare its reaction rate with TKS and chicory inulin. The initial inulin concentrations in the TKS extract, chicory extract and commercial inulin, were 38.47 g/L, 54.43 g/L, and

56.20 g/L, respectively. The hydrolysis process was monitored by measuring pH change and hydrolysis products such as inulin (DP 4+), kestose, sucrose, glucose and fructose by

HPLC. Samples of the reaction media (4 mL) were taken periodically and subsequently placed in a boiling water bath at 100°C for 10 minutes to deactivate the enzymes then quenched in ice for 5 minutes. The samples were filtered using 0.45 µm (13 mm dia) non-sterile nylon syringe filters (Tisch Scientific) to eliminate the enzymes to obtain samples suitable for HPLC analysis (see method below).

The hydrolysis was continued until the fructose production reached a steady-state during hydrolysis. Endoinulinase can cleave inulin at any β (2→1) linkage, resulting in oligosaccharides that are not completely hydrolyzed, and monosaccharides as its hydrolysis products. Endoinulinase is largely used in industries for the production of fructo-oligosaccharide syrup. An ideal enzyme for inulin hydrolysis, should have both exo-inulinase and endo-inulinase activities that would predominantly produce monomers as its hydrolysis products. The commercial enzyme Fructozyme L, which is a mixture of endo-and exo-inulinase, reportedly produced more fructose than endo-inulinase enzymes

(Michel-Cuello et al 2012). However, recently this enzyme was discontinued and is not

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commercially available. So, there were not many options available for choosing commercial inulinase enzymes.

4.2.6 Analytical methods

4.2.6.1 High pressure Liquid Chromatography

Several columns were tested for their ability to separate the hydrolysates, initial attempts to use a Biorad Aminex HPX-87P column and a Waters X-bridge amide column were not successful. The Biorad column, which uses HPLC grade water as the mobile phase, had poor separation and was unable to separate inulin hydrolyzate compounds.

The Waters X-bridge amide column uses acetonitrile as the mobile phase which can precipitate enzymes and short-oligomers (Ku et al., 2002). Other polar solvents such as ethanol, acetone and propanol also were found to precipitate inulin and oligosaccharides

(Ku et al., 2002). Owing to the complexities involved, this column was decided not appropriate to analyze inulin samples.

The sugar concentrations in hydrolyzates and fermentation media were measured using a high performance liquid chromatography (HPLC) equipped with refractive index

(RI) detector (Agilent Technologies 1200 Series) with an organic acid column (Rezex

ROA-Organic Acid H+ column, 300 mm X 7.8 mm). This column is an ion-exclusion column packed with 8% cross-linked sulfonated styrene-divinylbenzene (SDVB) adducts.

This column was able to distinguish oligosaccharides up to DP 3, and any compounds greater than DP 3 co-eluted with inulin peak.

The calibration curve was prepared from nine different calibration standards using pure glucose (MP chemicals), fructose (Fisher), sucrose (Sigma), kestose (TCI, Ltd) and 178

inulin (Alfa Aesar). HPLC chromatograms of the standard samples and their calibration curves are presented in Appendix I. The mobile phase was 0.0025M H2SO4 (Fluka, 50% sulfuric acid, 0.35 ml diluted with millipore water to 1 L, pH 2) at a flow rate of 0.6 mL/min. Sample volumes of 10µL were injected automatically. The column temperature and the detector temperature were 25°C. When column operated at a temperature of

25°C, optimum selectivity, precision and selectivity of sugar compounds can be attained

(Tomlins et al., 1990).

It should be noted that column temperature played a crucial role in quantification of low DP oligosaccharides. The acidic mobile phase (pH 2.00) of the HPLC breaks down the β (2→1) linkage easily resulting in hydrolysis of oligosaccharides into monomers. This behavior was observed in our preliminary experiments. However, when pure inulin was injected into the column, no such acid-induced hydrolysis was observed.

Inulin was much more stable in acidic mobile phase even at column temperature of 80°C, whereas, at column temperature of above 30°C, kestose and sucrose hydrolyzed into glucose and fructose. Since such hydrolysis could affect accurate estimation of sugars, column temperature of 25°C was adopted for HPLC analysis.

4.2.6.2 Gas chromatography

The solvents and acids produced during fermentation were quantified using a gas chromatography (GC) system (7890A Agilent Technologies Inc., Santa Clara, CA, USA), equipped with a flame ionization detector (FID) and 30 m X 320 µm (length x internal diameter) with 0.5µm (HP-Innowax film) J x W19091N-213 capillary column (Zhang,

Han et al. 2011). Six different calibration standards containing acetone, butanol, ethanol, 179

acetic acid and butyric acid were used to prepare a calibration curve. ABE yield (g/g) was calculated as the maximum amount of ABE produced per gram of substrate utilized.

ABE productivity was estimated as maximum ABE produced (g/L) divided by the total fermentation time (h) or when the fermentation ceased (Richmond et al., 2012; Ezeji,

Blaschek 2008).

Fermentation samples were centrifuged at 10,000 x g for 8 minutes and the culture supernatant was analyzed using HPLC and GC. All HPLC samples were filtered using 0.45 µm (13 mm diameter) non-sterile nylon syringe filters (Tisch Scientific) prior to analysis. Filtering of HPLC samples is essential to remove dirt or any other small particulates associated with samples, which might interfere with the compound of interest, increase column pressure and possibly block the separation column.

4.2.6.3 Biomass concentration

Cell concentration was estimated by measuring the optical density of the culture,

(after centrifuging for 8 minutes at 10,000 × g) at 600 nm in a DU800 spectrophotometer

(Beckman Coulter Inc., Brea, CA). TKS and chicory extracts contained dirt or suspended particles (came from roots) that settle in the bottom of centrifuging tube along with cell biomass. Washing or removing the dirt from the cell pellet was challenging. To overcome this problem, both of the extracts were extensively centrifuged (14000 x g for

30 minutes) before and after hydrolysis. The supernatant hydrolyzate was collected and centrifuged again for 20 minutes (14,000 x g) to remove any remnant particulates prior to fermentation medium preparation.

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4.2.7 Production of endo-and exo-inulinase enzymes

4.2.7.1 Microorganisms

Kluyveromyces marxianus var. bulgaricus ATCC 16045 and Kluyveromyces marxianus ATCC 52466 were obtained from American Type Culture Collections,

Manassas, VA, USA. Reportedly, strain ATCC 16045 produces endo-inulinase (EC

3.2.1.7) and the strain 52466 secretes exo-inulinase (EC 3.2.1.80) (Source: www.atcc.org). Strain ATCC 16045 was maintained on 200 YM (malt-yeast) agar slants

(pH 6.20) containing yeast extract (Fluka) 3 g/L, malt extract (Difco) 3.0 g/L, glucose 10 g/L, peptone (sigma) 5g/L, and agar (Difco) 20 g/L. The other strain ATCC 52466 was maintained on YEPD (Yeast extract-peptone-dextrose) agar slants (pH 5.60) made up of

10 g of yeast extract, 20 g of peptone, 20 g of dextrose and 20 g of agar per liter

(Rouwenhorst et al., 1990). The slants were sub-cultured every 14 days to maintain strain viability and stored at 4°C for short-term perseveration. The overview of inulinase production and the process involved is illustrated in Figure 4.5.

4.2.7.2 Optimization of culture conditions

Temperature, pH, agitation

The optimum process parameters widely reported for inulinase production from

Kluyveromyces marxianus (Pandey & Selvakumar, 1999; Singh & Bhermi, 2008) were used in this study. The optimum temperature and agitation rate for both of the strains were 30°C and 150 rpm, respectively. The fermentation medium pH of ATCC 16045 was 5.00 whereas it was 6.00 for ATCC 52466.

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Inoculum preparation

Two inocula media namely 200 YM medium for strain ATCC 16045 and YEPD medium for the strain ATCC 52466 were prepared. A loopful of cells from the slant were transferred into a 100 ml conical flask containing 20 ml of culture media and incubated at 30°C at 150 rpm for 24 h in rotary shaker (Lab-line Orbital Environ-Shaker

Model 3527) (Pandey & Selvakumar, 1999). The grown inocula culture was then added to optimized fermentation medium.

Carbon sources

Inulin, sucrose, glucose, and fructose have been reported as suitable carbon sources. In particular, inulin and sucrose were extensively tested by researchers who found that both compounds induced more inulinase production than any other sugar compounds (Kalil et al., 2001; Silva-Santisteban et al.2006; Gill et al., 2003). Our preliminary experiments showed more inulinase production from inulin medium than from sucrose medium (data not shown).

Substrate concentration

The optimum substrate concentration for each strain was evaluated by varying the inulin concentration from 5 g/L-30 g/L in the fermentation medium. The two strains showed different optimal substrate concentrations: 20 g/L inulin for Kluyveromyces marxianus var. bulgaricus ATCC 16045, and 30 g/L inulin for Kluyveromyces marxianus

ATCC 52466.

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4.2.7.3 Composition of fermentation media

The fermentation medium for Kluyveromyces marxianus var. bulgaricus ATCC

16045 contained 20 g/L inulin, 5 g/L yeast extract, 10 mg/L SDS (sodium dodecyl sulphate) (Amresco) and 0.1 g/L K2HPO4 (Kalil et al., 2001). The culture medium for

Kluyveromyces marxianus ATCC 52466 had 30 g/L inulin, 5 g/L yeast extract, 10 mg/L

SDS, and 2 mM MnSO4 (Singh & Bhermi, 2008). Addition of SDS to the medium reportedly increases enzyme production because SDS increases the permeability of cell membrane and thereby influences easy release of enzyme into the medium (Singh &

Bhermi, 2008).

4.2.7.4 Shake-flask level fermentation

The fermentations were carried in 250 ml Erlenmeyer flasks with 100 ml of fermentation medium and inoculated with 5% (v/v) of cell suspension. Each flask was incubated at 30°C for 84 h in a rotary shaker at 150 rpm. The samples were withdrawn every 12 h to analyze enzymatic activity, optical density and residual sugars in the medium.

4.2.7.5 Enzyme extraction

Samples were taken every 12 h and centrifuged at 10,000×g for 15 minutes at

4°C. The extra-cellular enzyme activity in the supernatant was determined (Gupta et al.,

1994; Kushi et al., 2000).

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4.2.7.6 Inulinase enzyme assay

The enzyme assay was performed as described by Singh et al (2007). A mixture of 0.1 ml of crude enzyme (supernatant) and 0.9 ml of 0.1 M sodium acetate buffer (pH

5.00) containing 2% inulin was incubated at 50°C for 15 minutes in a water bath. After incubation, the resultant mixture was heated in a boiling water bath (100°) for 10 minutes to inactivate the enzyme and then cooled to room temperature (Singh et al., 2007; Pandey

& Selvakumar, 1999). The amount of fructose released was measured by HPLC. One unit of inulinase activity was defined as the amount of enzyme required to release one micromole of fructose per minute under standard assay conditions (Ayyachamy et al.,

2007).

4.3 Results and discussion

4.3.1 Enzymatic hydrolysis of inulin

The hydrolysis of different sources of inulin occurred progressively and displayed different patterns depending on enzyme concentration and amount of substrate(s) present in the reaction. Though fructose was found to be the major hydrolysis product of inulin, significant amounts of short oligomeric compounds such as kestose (DP 3), and sucrose

(DP 2) were also present. Since the enzyme is endo-acting in nature, the presence of short-oligomers was expected. At zero hour, the chicory extract contained 68.65% inulin,

7.04% kestose, 4.81% sucrose, 3.77% glucose and 15.72% fructose. On the other hand, the TKS extract constituted 57.53% inulin, 3.81% kestose, 2.97% glucose and 35.68%

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fructose. The commercial inulin standard had 100% inulin i.e. did not contain sugars below DP 4.

Hydrolysis conducted with an enzyme activity of 150 IU/g of inulin showed sharp degradation of inulin after 2 h compared to hydrolysis carried out with lower enzyme activities i.e. 50 IU and 100 IU per gram of inulin (Figure 4.7 & 4.8). The extent of substrate hydrolysis (%) was defined as the ratio of consumed substrate/amount of substrate provided ×100 (Michel-Cuello et al 2012). Experimental results of reactions with an enzyme activity of 150 IU/g of inulin showed that 90% of the chicory inulin and commercial inulin was degraded in two hours, while only 64% of the TKS inulin was degraded. In general, the reaction rate slowed after 2 h in all samples (Figure 4.7). After

4 h, 98% of chicory inulin had been hydrolyzed and the hydrolyzate contained 1.14 g inulin, 10.39 g kestose, 8.20 g sucrose, 12.47 g glucose and 36.16 g fructose per liter

(Figure 4.8). Only 71% and 83% of chicory inulin was hydrolyzed in reactions treated with the lower enzyme activities of 50 IU/g and 100 IU/g, respectively (Figure 4.7).

Although near complete inulin degradation (more than 95%) was achieved in the chicory inulin and commercial inulin samples (Figure 4.8), in which the amount of monomers (glucose and fructose) available was 51.34 g/L (71.50%) and 44.49 g/L

(74.31%) from chicory and commercial inulin, respectively. The highest concentration of monomers obtained from TKS inulin hydrolysis was 49.66 g/L (58.75%) (Figure 4.8). It should be noted that around 14-15 g/L inulin (~ 18% of the reaction mixture) remained in the TKS inulin samples even after 42 h of hydrolysis while this behavior was not observed in chicory and commercial inulin samples (Figure 4.8).

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As a general trend, the rate of substrate hydrolysis and fructose production was very high during the initial 2 hours. This was due to high amount of substrates present in reaction mixture. High substrate availability in the reaction renders enormous amount of points of attack for the enzyme (endo-inulinase). Higher amount of enzyme in the reaction mixture increases the rate of hydrolysis and thereby drastically reduces the reaction time (Ricca et al., 2009a & 2009b). This behavior was observed in the hydrolysis experiments for the initial 6 hours, however, after 6 h, fructose production appeared to remain constant in TKS and chicory inulin samples, while commercial inulin sample showed gradual increase in fructose production until 30 h. Although it showed that enzyme deactivation did not occur in commercial inulin samples, fructose production from commercial inulin was much slower and the reactions took a longer time (Figure 4.7

& 4.8). Since the commercial inulin employed was highly purified (~ 98% pure), it is unlikely that impurities or inhibitors slowed the hydrolysis. One plausible reason for slow reaction rate of commercial inulin hydrolysis could be due to change in pH of the hydrolyzate. The starting pH of the commercial inulin solution was 5.00 but after 6 h, pH decreased to 4.28 (Figure 4.8). It should be noted that the enzyme exhibits maximum activity between 4.60 and 5.20 (Figure 4.4), hence the slow fructose production rate from commercial inulin could be attributed to decreased enzyme activity due to pH change.

TKS and chicory inulin hydrolyzate pH did not go below 4.90 (Figure 4.9).

Novozyme 960 did not appear to have invertase activity. Presence of significant amounts of sucrose in the hydrolysis products of all the inulin samples demonstrated lack of invertase activity in the enzyme (Figure 4.7 & 4.8). The discrepancy in the rate and

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efficiency of hydrolysis between different inulin could be due to presence of impurities or inhibitory compounds, substrate inhibition, differences in the average DP of inulin molecules between samples. Michel-Cuello et al (2012) reported such disparity in hydrolysis among inulins from different sources.

Complete degradation of chicory inulin was much faster (4 h) than commercial inulin (30h) (Figure 4.7). Chicory inulin has a low DP (estimated average DP of 11-12), compared to commercial inulin (estimated average DP of 27-28) (Personal communication from Dr. Fred’s research group) and this might also have caused chicory inulin to degrade quickly in the presence of high enzyme concentrations (Michel-Cuello et al., 2012). The effect of DP on reaction kinetics is due to the number of molecules available for the reaction. The presence of lower DP inulin in the reaction medium increases molar concentration, which influences reaction rate (Ricca et al., 2009a). The reaction rate of TKS inulin (estimated average DP of 10-19) hydrolysis did not differ significantly at different enzyme concentrations (Figure 4.7). Therefore, for TKS inulin, initial reaction rate was not contingent upon enzyme activity but appear to be related to substrate concentration as observed by Michel-Cuello et al (2012). Another possibility could be substrate or product inhibition which might have hampered TKS inulin hydrolysis.

The optimum temperature (60°C), which showed maximum enzyme activity and enzyme thermo-stability as described by the enzyme manufacturer, was used as the operating temperature for the hydrolysis. It can be observed that initial reaction rate of inulin hydrolysis was very fast and subsequently it decreased quickly. This phenomenon

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was due to enzyme deactivation, which was reported earlier by Ricca et al (2009b &

2010b). According to them, optimum temperatures favor enzyme reaction kinetics but hinder activity retention and thereby increase enzyme deactivation rate. Hydrolysis conducted with lower temperatures was much more stable, and retained enzyme activity despite a slow reaction rate (Ricca et al 2009a; 2009b; 2010a & 2010b). A predictive model based on temperature optimization, reaction kinetics, deactivation rates and inulin hydrolysis for Fructozyme L (a mixture of exo-and endo-inulinases) was proposed by

Ricca et al (, 2009a; 2009b; 2010b). Such detailed studies on reaction kinetics using

Novozyme 960 would improve the potential of inulin enzymatic hydrolysis at an industrial scale, especially for fructo-oligosaccharide (FOS) production.

4.3.2 Production of exo- and endo-inulinases

4.3.2.1 Endo-inulinase production from Kluyveromyces marxianus ATCC 16045

Substrate concentration influenced endo-inulinase production from ATCC 16045.

The highest enzyme activity (5.79 IU/ml) was observed at a substrate concentration of 20 g/L (Figure 4.11a), though biomass concentration was lower than in the 30 g inulin/L culture medium (Figure 4.10a). The organism grew well in the inulin medium and the highest enzyme production achieved was similar to the 7.30 IU/ml obtained by Kalil et al

(2006a) using inulin substrate, and contrasts with the very low activity reported earlier by

Selvakumar & Pandey (1999). However, our results on inulin were much lower than the inulinase activity (> 100 IU/ml) obtained by researchers using (Kalil et al 2001; 2006a;

2006b; 2010) sucrose as a substrate. However, our preliminary experiments did not find sucrose to be an effective substrate for inulinase production from Kluyveromyces 188

marxianus ATCC 16045. Literature reported high inulinase activity from sucrose as carbon source often employed medium pH in the range of 3.50-4.50. Acid hydrolysis of sucrose due to the acidic nature of medium is one possibility which might leave an appreciable amount of monosugars in the medium. This could have resulted in false enzyme activity measurements since the inulinase assay measures the amount of monosugars released (Kalil et al 2001; 2006a; 2006b; 2010). Furthermore, inulinase assays cited in the literature do not use standard methods and there are significant differences in temperature, pH and substrate concentration as well as the amount of enzyme employed in various inulinase assays. Also, although the standard definition of one unit of inulinase activity is the amount of enzyme required to release 1 µmole of fructose per minute, some researchers define units in terms of the liberation of glucose or total reducing sugars (which includes glucose and fructose). The inulinase assay employed in this work defined units in terms of liberated fructose molecules and this could be a major contributing factor for limited inulinase activity from ATCC 16045.

Since the endo-inulinases primarily produces huge fractions of oligosaccharides with tiny portions of monosaccharides (i.e. glucose and fructose) (Sirisansaneeyakul et al., 2007), estimation of fructose to determine endo-inulinase activity is misleading and can result in reporting poor endo-inulinase activity than the actual rate of reaction. An inulinase assay capable of estimating the oligosaccharides would be more appropriate to determine the endo-inulinase activity; however, such assay was not reported in literature.

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4.3.2.2 Exo-inulinase production from Kluyveromyces marxianus ATCC 52466

Strain ATCC 52466 showed increased exo-inulinase production and cell growth with increased substrate concentration (Figure10b & 11b). The maximum enzyme activity observed was 9.87 IU/ml after 74 h of fermentation in 30 g/L inulin culture medium. The maximum biomass concentration was 2.64 (OD @ 600nm) which was much less than the maximum cell growth (13.34) observed from ATCC 16045 (Figure

10). Since 30 g/L substrate concentration in the medium did not seem to inhibit enzyme production in the batch fermentation, it is likely that higher substrate concentrations may improve enzyme production. The exo-inulinase production (9.87 IU/ml) observed in this study was 20-fold higher than the results reported by Selvakumar & Pandey (1999) and

Shin et al (2004).

Inulinase assays carried out with the mixture of crude endo- and exo-inulinase obtained in this study, did not yield increased enzyme activity. Sequential addition of crude enzymes in the inulinase assay (addition of endo-inulinase followed by exo- inulinase) also did not improve the total enzyme activity (data not shown). Although both exo- and endo-inulinases were from the same yeast species, it is clear that the assay reaction was severely inhibited (Data not shown). The presence of inhibitory metals or differences in optimal temperature and pH for maximum activity of these enzymes might have led to these results.

In summary, production of exo- and endo-inulinase was demonstrated in shake flask fermentation in this study. Though the enzyme activities reported from the organisms were encouraging, further optimization of the fermentation process and

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purification of crude enzymes would ameliorate enzyme production and enzyme activity and make it more suitable for industrial applications of inulin hydrolyzate. It is notable that both of these organisms can be grown in raw inulin extracts (Gupta et al., 1988;

Kango 2008) and still produce inulinase. Therefore, using TKS extract, in-situ production of inulinase enzymes, hydrolysis, and conversion of liberated sugar molecules into biofuels such as butanol and ethanol is feasible.

4.3.3 Butanol production from unhydrolyzed inulin media

At the start of this experiment, screening of Clostridia sp. was done to identify the best strain for use of inulin as the sole carbon source. Inulin P2 medium (~55 g/L inulin) prepared with commercial inulin and used as growth medium was used to test four different strains , namely, C. beijerinckii NCIMB 8052, C. beijerinckii NRRL B592, C. acetobutylicum ATCC 824, and C. saccharobutylicum P262. Preliminary experiments revealed that C. saccharobutylicum P262 grew well in the inulin medium and produced more solvents compared to the other strains tested (Figure 4.12). It is notable that all the strains demonstrated the ability to utilize inulin and produce solvents, though such ability was very limited. In particular, C. beijerinckii NCIMB 8052 and C. acetobutylicum

ATCC 824 strains exhibited inulolytic capability and produced maximum ABE of 4.07 g/L (3.39 g/L butanol) and 3.25 g/L (2.37 g/L butanol), respectively. This finding was in contradiction with earlier reports where Montoya et al (2000 & 2001) did not observe inulolytic activity in either C. beijerinckii NCIMB 8052 or C. acetobutylicum ATCC 824.

One possible reason for the inulolytic activity observed from these strains in our experiments could be due to the presence of calcium carbonate in the medium. Han et al

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(2012) observed increased heat shock proteins (grpE, dnaK) and activity of several important solventogenic enzymes such as coenzyme A transferase (CoAT), acetate kinase

(AK), acetoacetate decarboxylase (ACDC) and butanol dehydrogenase (BDH) in CaCO3- mediated P2 medium of C. beijerinckii NICMB 8052. Further they found that presence of Ca2+ ions in the medium alleviates stress during solventogenesis by stabilizing the membrane proteins, enhancing glucose utilization and mediating cellular signaling events

(Han et al., 2012; Kanouni et al., 1998). In the presence of Ca2+ ions in the medium, expression of relevant genes for inulolytic activity and thereafter, production of inulinase enzyme might have happened in the Clostridia species tested

In the inulin P2 medium, the maximum concentration of ABE produced by C. beijerinckii NRRL B592 was 3.82 g/L (3.28 g/L butanol) whereas C. saccharobutylicum

P262 produced 10.26 g/L ABE (7.08 g/L butanol). The average maximum cell density observed was 4.89 at 60 h for C. saccharobutylicum P262, 2-fold higher than the other strains. Accumulation of butyric acid, in the range of 3-4 g/L in C. beijerinckii NCIMB

8052 and C. acetobutylicum ATCC 824 fermented medium, showed that the switch from acidogenic phase to solventogenic phase was not complete in the inulin P2 medium. In comparison, the maximum butyric acid produced by C. beijerinckii NRRL B592 and C. saccharobutylicum P262 was 1.96 g/L and 2.31 g/L, respectively. Low butyric acid and solvent production by C. beijerinckii NRRL B592 infers that inulin utilization by this organism was relatively poorer than the other strains. This observation was further augmented by the relative poor cell growth of this organism in the inulin P2 medium

(Figure 4.12). It is notable none of the four organisms produced solvents in the first 24 h

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and this lag phase could be perceived as the time needed for the organisms to hydrolyze inulin into fermentable monosugars of sufficient concentration. Owing to the lag phase and the slow nature of fermentation, all the fermentations using inulin P2 medium were continued for 120 hours. After observing the results of inulin P2 medium with different organisms, it was clear that C. saccharobutylicum P262 was the superior strain

(maximum total ABE of 10.26 g/L). Solvent production among the other strains was not significantly different. Although C. beijerinckii NCIMB 8052, C. beijerinckii NRRL

B592 and C. acetobutylicum ATCC 824 gave low solvent yield from pure inulin medium in our experiments, the presence of relevant genes to utilize inulin in these organisms can be inferred from the results.

C. saccharobutylicum P262, which displayed higher inulolytic activity in the preliminary experiments, was chosen for ABE fermentation of unhydrolyzed raw inulin extracts. It should be noted that the raw chicory and TKS extracts contained significant amounts of glucose and fructose. Raw TKS extract medium yielded 8.48 g/L total ABE

(4.86 g/L butanol; ABE productivity was 0.12 g/L/h) after 72 h. From the chicory extract medium, C. saccharobutylicum P262 produced total ABE of 12.50 g/L (8.58 g/L butanol;

ABE yield of 0.32 g/g, 0.21 g/L/h ABE productivity) after 60 h of fermentation. Inulin

P2 medium (~55 g/L inulin) was used as control medium, which gave total ABE yield of

9.71 g/L (6.56 g/L butanol, 0.33 g/g yield, 0.10 g/L/h productivity) after 96 of fermentation (Table 4.3; Figure 4.13). The total ABE (12.50 g/L) produced from unhydrolyzed chicory extract medium was similar to the amount (9.40 g/L of total acetone and butanol) produced from unhydrolyzed Jerusalem mashed artichoke tubers

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using C. acetobutylicum ATCC 4259 (Fan et al., 1983). Also, these results were comparable to results reported by Oiwa et al., (1987) from C. pasteurianum strain I-53 using 68 g/L unhydrolyzed dahlia inulin medium (total ABE of 20. 70 g/L).

The onset of solventogenesis in the chicory extract medium reached 6.47 g/L butanol after 24 h whereas there was no butanol production at this point of time in the

TKS extract medium (Figure 4.13). An extended lag phase was observed in the TKS extract medium and butanol production was observed only after 48 h. From the pure inulin P2 medium, C. saccharobutylicum P262 produced butanol after 36 h and the reason for this lag phase, as explained in the earlier section, could be understood as the time needed for the organisms to break inulin into fermentable monosugars.

Although TKS extract medium contained 28.32 g/L of fructose at 0 h, the medium took long time to switch to solventogenesis mode. Notably, at 0 h, TKS extract medium had 6.37 g/L acetic acid which was similar to 6.39 g/L acetic acid observed in chicory extract medium (Figure 4.14). The very high acetic acid accumulation in the TKS and chicory extract medium at 0 h could be ascribed to the presence of acetic acid in the extract (probably due to fermentation during storage and chemical reactions associated with the minerals leached from roots and the extraction solvent) and addition of buffer stock (which contains ammonium acetate) could have increased the acetic acid concentration in the medium. It should be noted that KOH was added to the extract to increase the pH up to 6.50. The original pH of the TKS and chicory extracts were 4.78 and 4.49, respectively. Intriguingly, TKS extract needed much more KOH than the chicory extract for the pH adjustment. Addition of surplus KOH to the TKS extract

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might have been a contributing factor for the pH profile observed during fermentation.

The lowest pH recorded during fermentation of TKS extract was 5.98 whereas it was 5.57 and 5.17 for chicory extract and pure inulin P2 medium, respectively (Figure 4.15).

Addition of KOH during pH adjustment may have prevented drop in pH during fermentation, i.e. there was not enough acid production in the medium.

A gradual decrease in acetic acid concentration in the TKS and chicory extract media was observed and, at the end of fermentation, 4.71 g/L and 2.33 g/L acetic acid, respectively, was present. In contrast to this decreasing trend, butyric acid accumulated as fermentation progressed in both TKS and Chicory extract medium. The highest butyric acid concentration observed in the TKS and Chicory extract medium was 4.62 g/L and 4.95 g/L, respectively (Figure 4.14). This observation indicates that rapid termination of solventogenesis might have occurred after the medium switched from acidogenesis. This resulted in accumulation of butyric acid in the medium. The total concentration of undissociated acetic acid and butyric acids might have surpassed a critical threshold value, and that could have inhibited solventogenesis. Maddox et al

(2000) stated that exceeding the critical value of total undissociated acids (57-60 mmol/L) could halt further fermentation and metabolic activity, which includes substrate utilization and ABE production. This hypothesis fits well with the results obtained from chicory and TKS extract media. The overall observation of the two extract media revealed that after, Solvent production and substrate utilization both seem to have become limited when the butyric acid concentration increased to more than 4 g/L (Figure

4.13, 4.14 & 4.16), which occurred after 36h for chicory medium and 48h for TKS

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medium. This correlated to the decrease in cell density observed in both inulin extract media (Figure 4.20). Although the trend of decreasing acetic acid concentration was observed in both extract media, owing to the increase in butyric acid production, the relative concentration of butyric acid was more in the media compared to acetic acid.

The increased concentration of butyric acid in the medium might have contributed more to the cessation of ABE fermentation than acetic acid concentration. On a molar concentration, butyric acid is reportedly more toxic than acetic acid (Wang et al., 2011;

Maddox et al., 2000; Soni et al., 1987).

The fermentation results of TKS extraction medium (Table 4.3) shows that the maximum ethanol produced was 2.14 g/L (which gave an ethanol:butanol ratio of 1:2) which was more than the acetone produced (1.64 g/L) in this medium. Similarly, the pure inulin P2 medium produced 1.67 g/L ethanol whereas the acetone was 1.71 g/L.

However, such increase in ethanol:acetone ratio was not observed in the chicory extract inulin medium. It is unclear whether the difference in inulin DP between the media caused this difference in ethanol production. In that case, inulin or any other factors associated with it might have increased level of enzymes responsible for ethanol production in ABE pathway. However, such a proposition warrants detailed study.

4.3.4 Sugar consumption pattern of C. saccharobutylicum P262 from unhydrolyzed inulin extract media

C. saccharobutylicum consumed a total of 28.96 g/L inulin from the pure inulin

P2 medium. The rate of substrate consumption was higher in the first 48 h of fermentation, which correlates with the lag phase observed in the inulin P2 medium. The

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inherent inulinase enzyme producing ability of this organism was apparent from the inulin utilization. Interestingly, analysis of residual sugars at every 12 h time interval by

HPLC showed only inulin and fructose peaks (Table 4.2; Appendix E). Therefore, it appears that C. saccharobutylicum P262 produced exo-inulinase enzyme, which cleaved inulin molecule at its terminal unit, resulting in monomer production. Glucose is the preferred substrate in the metabolism of many Clostridia sp (Ezeji & Blaschek, 2008), and the residual sugar analysis of Chicory extract and pure inulin P2 medium showed no residual glucose after 12 h of fermentation. In the presence of glucose, inulin consumption by C. saccharobutylicum P262 was severely limited, and the organism began inulin consumption only after the exhaustion of glucose in the medium i.e. after 12 h. Based on percentage of sugars utilized, after glucose exhaustion, the next preferred sugar choice was fructose. The order of sugar preference by C. saccharobutylicum P262 is glucose > fructose > sucrose > inulin > kestose. This sugar consumption pattern of the organism was demonstrated clearly in the chicory inulin extract media (Figure 4.16b).

In the pure inulin P2 medium, it can be assumed that C. saccharobutylicum P262 consumed the released glucose immediately after hydrolysis. The amount of glucose released upon hydrolysis of inulin depends on inulin chain length. Since reports suggest that commercial inulins have DP between 12 and 22, it can be presumed that glucose: fructose availability ratio in the medium could be in the range of 12 to 22. The pH profile presented in Figure 4.15 showed that the lowest pH was 5.17 from the pure inulin

P2 medium. Inulin undergoes hydrolysis by concerted action of pH below 4.0 and temperatures above 60°C (Glibowski & Bukowska, 2011). Medium pH of more than

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5.00 at temperature of 35±1°C nullified any possibility for self-hydrolysis of inulin.

Therefore, C. saccharobutylicum P262 did enzymatically hydrolyze inulin for ABE production.

In addition to exo-inulinase activity, C. saccharobutylicum P262 also exhibited invertase or sucrose activity which can be inferred from the 100% sucrose utilization displayed clearly in the chicory extract medium (Table 4.3, Figure 4.16b). Exo-inulinase activity from Clostridia sp. was earlier reported in C. acetobutylicum IFP 911 by Looten et al (1987). They found that the enzyme was less active with short oligomeric compounds such as kestose and sucrose. Our experimental results showed that kestose was not consumed very well by C. saccharobutylicum P262, however, sucrose was utilized immediately by this organism. This observation was buttressed from the findings of Efstathiou et al (1986) who found inulinase and sucrase activity in a newly isolated strain C. acetobutylicum ABKn8. Further, they concluded that inulinase production can be induced only by inulin but not by xylose, fructose or sucrose (i.e. inulinase production was inducible). Addition of glucose was found to repress inulinase activity (Efstathiou et al., 1986), and such repression of enzyme activity in C. saccharobutylicum P262 was observed especially in the chicory inulin medium. Bacterial inulinases reportedly have high affinity for inulin substrate as compared to sucrose (Efstathiou et al., 1986; Looten et al., 1987). HPLC chromatograms of TKS and chicory extract showed two unknown peaks, one at retention time of 8.15 min and the other one at 8.79 min (Appendix I)

Based on this observation, it can be inferred that the molecule eluted at 8.15 min was bigger than the molecule appearing at 8.79 min in the chromatogram. It should be

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noted, that kestose elutes at 7.70 min and sucrose appears at 8.45 min. Based on this reference, it is highly likely that the unknown molecule which elutes at 8.15 min is be inulotriose (made up of three molecules of fructose) and the other unknown molecule eluting at 8.79 min could be a disaccharide molecule since it elutes right after sucrose in the column. The two possible disaccharide molecules that might present in the inulin extract medium are inulobiose (a di-fructose) and difructose anhydride (DFA) (cyclic disaccharide molecule made up of two fructose units). Inulobiose (C12H22O11) has a molecular weight of 342.3 g/mol whereas difructose anhydride (C12H20O10) is 324.28 g/mol56. Since the molecular weight of inulobiose is identical to sucrose, it is very likely that inulobiose elutes very close to sucrose. On the other hand, the molecular weight of difructose anhydride (324.28 g/mol) suggests that this molecule would elute after sucrose molecule (Appendix D).

Comparison of HPLC chromatograms of TKS and Inulin extract media before and after fermentation (Appendix D & E) suggested that there was no decrease in the amount of inulotriose present in the extracts after fermentation. This finding corroborated our earlier observation that C. saccharobutylicum P262 had poor ability to consume trisaccharide molecules i.e. kestose and inulotriose. Difructose anhydride (DFA) can exist in four different forms depending on the linkage (Saito & Tomita, 2000) and the presence of appreciable amounts of DFA in raw inulin extract was possible because of the steps involved in inulin extraction. At low pH and at high temperatures (80-100°C), fructose degraded easily into difructose anhydride (Arand et al., 2002), which often

56 http://www.guidechem.com/reference/dic-295125.html

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colors inulin hydrolyzate (Dilipkumar et al., 2011; Kim & Lim, 2002). Interestingly, lower DFA levels were observed after fermentation of TKS and chicory extracts

(Appendix D & E). These results suggest that C. saccharobutylicum P262 did consume

DFA, although in small amounts. However, this hypothesis needs further study. No reports on Clostridia utilizing DFA as carbon source are available currently.

Carbon catabolite repression (CCR) is defined as the inhibition of an enzyme active site by a preferred carbon source such as glucose (Tracy et al., 2011; Ren et al.,

2010). In C. acetobutylicum ATCC 824, it was found CCR is regulated by a catabolite control protein A (CcpA) encoded by gene CAC3037 (Ren et al., 2010). Such a repression mechanism may have repressed inulinase activity in the presence of glucose since upon glucose exhaustion no repression was observed and C. saccharobutylicum

P262 was able to consume multiple sugars. The presence or absence of CCR mechanism differs among Clostridial strains, Heluane et al (2011) did not find evidence of CCR in C. beijerinckii SA-1 when grown on medium containing mixtures of glucose and xylose.

Regulation of the CCR mechanism in C. saccharobutylicum P262 in mixed-sugar media was not clear since this organism exhibited two different patterns depending on glucose availability. (Table 4.3; Figure 4.16b). From the chicory extract medium the organism utilized 13.17 g/L inulin, 1.81 g/L kestose, 6.21 g/L sucrose, 3.53 g/L glucose and 15.80 g/L fructose from the medium. It should be noted that this organism preferred

DP3 molecules the least. The chicory extract medium showed better ABE production and sugar utilization than TKS extract medium which was least successful possibly

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because of the acid-load (acetic and butyric acid concentration) and inhibition associated with it.

The ability of C. saccharobutylicum P262 to utilize multiple-sugars in a medium makes it more suitable for ABE fermentation of plant inulin extracts since these extracts likely contain a mixture of oligomers and monomers. Also, from the results of this study, mixed sugar media produced more solvents than pure inulin medium. These results suggest that low-cost complex substrates, such as inulin extracts, can be used directly for

ABE fermentation without additional cost-intensive enzymes. Future research focusing on identification of relevant genes responsible for inulinase production, genetic manipulation, enzyme characterization and optimization of fermentation parameters may help better understand the ABE production from raw inulin extracts using this organism.

4.3.5 Butanol production from hydrolyzed inulin media

Since fructose is the major catabolite of inulin hydrolysis, we employed fructose

P2 medium (60 g/L fructose) to determine the best strain for utilizing hydrolyzed inulin.

C. beijerinckii NCIMB 8052, C. beijerinckii NRRL B592, C. acetobutylicum ATCC 824, and C. saccharobutylicum P262 were screened. From the fructose P2 medium, C. beijerinckii NCIMB 8052 and C. acetobutylicum ATCC 824 produced total ABE of

20.18 g/L (14.29 g/L butanol) and 20.15 g/L ABE (14.13 g/L), respectively. On the other hand, C. beijerinckii NRRL B592 and C. saccharobutylicum P262 gave maximum total

ABE of 16.36 g/L (11.80 g/L butanol) and 10.47 g/L (7.35 g/L butanol), respectively

(Figure 4.17). From these results, it was evident that C. beijerinckii NCIMB 8052 and C. saccharobutylicum ATCC 824 produced equivalent amounts of total ABE from the

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fructose P2 medium. Therefore, the capability of these strains to use inulin-derived oligomers in the presence of monomers (glucose and fructose) was investigated.

Commercial inulin (~55 g/L) was hydrolyzed using endo-inulinase (Novozyme 960) at

200 IU/g of inulin for 1 h and 48 h to prepare media with different amounts of oligosaccharides (Tables 4.5. and 4.6, respectively). An unknown compound appears in the HPLC chromatogram of 1h pure inulin hydrolyzate at the retention time of approximately 7.40 minutes, suggesting inulotetrose (a tetra-fructose), but this was not quantified (Appendix F) and so the estimate of total sugars after 1 h (Table 4.5 & 4.6) is an underestimate. (From the preliminary experiments we found that nystose, which is a molecule containing three fructoses with a terminal glucose unit, eluted around 7.1 minutes which overlapped with the inulin peak, so we didn’t include it in the HPLC standard calibration).

Intriguingly, in the 1 h pure-inulin hydrolyzate medium, both C. beijerinckii

NCIMB 8052 and C. acetobutylicum ATCC 824 displayed carbon catabolite repression for sucrose but not for kestose or the 4-mer. It should be noted that ~10 g/L of monomers present in the medium was enough to induce sucrose catabolite repression. On the other hand, kestose was hydrolyzed into sucrose and fructose, from which the organism consumed only fructose leaving the sucrose molecule in the medium (Figure 4.6). The decrease in HPLC peak height of the unknown compound (Appendix F), indicates that that the organism was able to hydrolyze it. The residual sucrose molecule would have also contributed to the total sucrose quantified at the end of the fermentation.

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Tangney et al (2003) found that the regulation of sucrose metabolism was not the same in C. acetobutylicum ATCC 824 and C. beijerinckii NCIMB 8052 due to different scr operons in each organism. Despite these differences, they found that sucrose metabolism was repressed in the presence of glucose in both, and also possibly that fructose might repress sucrose metabolism (Tangney et al 1998; Tangney et al., 2000).

This might be why catabolite repression was observed for sucrose in the 1 h pure-inulin hydrolyzate medium. However, no such repression was found for kestose and the short- oligomeric compound. C. beijerinckii NCIMB 8052 and C. acetobutylicum ATCC 824 produced total ABE of 5.62 g/L (4.47 g/L butanol) and 4.96 g/L (3.79 g/L butanol), respectively, from the 1 h pure-inulin hydrolyzate (Figure 4.22).

The 48 h pure-inulin hydrolyzate contained ~ 45 g/L of monomers (mostly fructose) and ~15 g/L of short-oligomers such as sucrose and kestose (Table 4.7 & 4.8).

As expected, both organisms showed severe catabolite repression, and the oligomers were not consumed. This behavior implies that both organisms should have a critical limit in the amount of monomers available in the medium in order to express complete carbon catabolite repression. The highest total ABE produced from the 48 h pure-inulin hydrolyzate was 14.99 g/L (9.21 g/L butanol) by C. beijerinckii NCIMB 8052, whereas

15.28 g/L (9.12 g/L butanol) was produced by C. acetobutylicum ATCC 824 (Figure

4.22). It was apparent that both the organisms displayed similar behavior in fructose and mixed-substrate medium (monomers and oligosaccharides). Owing to the similarity between the two organisms in terms of substrate utilization and solvent production, we

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selected C. beijerinckii NCIMB 8052 for our further experiments with inulin hydrolyzate media.

TKS and chicory medium were hydrolyzed for 48 h with endo-inulinase enzyme

(Novozyme 960) at 200 IU/g of inulin (further increase in hydrolyzing time was found to have no influence on monomer production) (Appendix H). The hydrolyzate contained a significant amount of inulin and other short-oligomers along with monomers (i.e. it was not completely hydrolyzed) (Table 4.4). Fermentation of chicory and TKS hydrolyzate using C. beijerinckii NCIMB 8052 produced total ABE of 12.60 g/L and 10.00 g/L, respectively, which was less than the total ABE production from 48 h pure-inulin hydrolyzate medium (total ABE 14.99 g/L) and the pure fructose medium (total ABE

20.18 g/L) (Figure 4.18). In comparison with our results, Chen at al (2010) obtained

11.21 g/L butanol from the acid hydrolyzate of Jerusalem artichoke juice using C. acetobutylicum L7. In general, both chicory and TKS hydrolyzate media produced very little solvent in the first 24 hours which was reflected in the slow cell growth compared to the pure-inulin hydrolyzate and the fructose P2 control employed (Figure 4.21). Notably, solvent production from the 48 h pure-inulin hydrolyzate was faster than the fructose P2 medium in the first 24 h of fermentation, due to the presence of glucose in the pure-inulin hydrolyzate medium. Since glucose was the preferred substrate for Clostridia, C. beijerinckii NCIMB 8052 produced solvents faster than in fructose P2 medium (Figure

4.18). However, due to catabolite repression, the solvent production from the pure-inulin hydrolyzate medium was limited, whereas there was no such repression in the fructose P2 medium (fructose was the only carbon source) and it produced maximum total ABE of

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20.18 g/L (Table 4.4). Lee & Blaschek (2001) found that fructose PTS activity was inducible by more than 10-fold for C. beijerinckii NCIMB 8052 when fructose was used in a growth medium instead of glucose. Presence of glucose and other oligomers might have repressed fructose PTS (Phosphotransferase system) activity, and that would have also contributed to the limited ABE production from the pure-inulin hydrolyzate medium.

The carbon catabolite repression and high-salt load (natural presence of salts in the inulin extract and addition of excessive base during pH adjustment) might have inhibited the solvent production from the chicory and TKS inulin hydrolyzate since both of the hydrolyzates contained less solvents. This observation was further supported by the reduced capacity for uptake and reassimilation of the acids produced by the microorganism from the inulin extract hydrolyzate media. On the other hand, low levels of acids were observed in the pure-inulin hydrolyzate and fructose P2 medium, which indicates a more efficient fermentation (Figure 4.19).

Since C. beijerinckii NCIMB 8052 solvent production was dependent on the amount of monomers available in the medium, inulin extracts completely hydrolyzed with a mixture of endo-and exo-inulinases would increase monomers in the medium and thereby improve the solvent yield and overall performance of the inulin extract hydrolyzate fermentation.

4.3.6 Applications of residual biomass of TKS and guayule

Less than 15% of TKS and guayule plant materials are rubber, and fractionation yields a significant amount of co-products–inulin and residual biomass in TKS, and resin and biomass in guayule. Utilization of inulin for butanol production was demonstrated in

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this study. Economical production of rubber from these plants is largely dependent on successful valorization of residual biomass such as bagasse, leaves, and stems into income derived co-products. The energy density (through bomb calorimeter) of the residual biomass is presented in Table 4.2. Notably, the energy density of TKS and guayule biomass rank above the high energy woods and the lower energy grasses

(Boateng et al., 2009).

U.S. industrial processes generate solid and gasesous by-products that amount to more than two quads (one Quad = 1015 BTU, or 1.055×1018 J) of energy per year

(Boateng et al., 2010). These extensive energy sources are under-utilized and if efficiently used, could supply enormous amounts of energy needed for various industrial processes and alleviate the energy demands on fossil fuels (Boateng et al., 2010). A description of bagasse components, structure and possible applications is presented in

Chapter 2 (section 2.3.1) of this thesis.

Reportedly, TKS leaves contain 12-15% inulin (based on fresh weight) (Ricca et al., 2007). The energy density of TKS leaves (15.70 MJ/kg) substantiate that this could also be a good source of biomass for renewable fuel production. Regarding guayule leaves (which account for 30-70 wt% of plant materials depending on season), Boateng et al. (2010) observed that guayule leaf fractions have little fuel value owing to higher concentration of protein and mineral ash present in them. Therefore, biofuel production from guayule leaf did not appear to be an efficient application.

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4.4 Conclusion

Fermentation of unhydrolyzed inulin media with C. saccharobutylicum P262 produced fair amount of solvents (TKS, 8.48 g/L ABE; chicory, 12.50 g/L ABE), and the strain was able to utilize inulin, sucrose, glucose, and fructose concurrently during fermentation, although the rate of sugar utilization varied between sugar molecules. DP3 molecules are poor substrate for this organism. However, the simultaneous consumption and metabolism of these sugar molecules for solvent production by C. saccharobutylicum

P262 is a positive trait since the sugar composition of raw inulin extract derived from plants such as TKS is diverse. Hence, further optimization of the fermentation could enhance solvent production from raw inulin extract.

Carbon catabolite repression was observed during the fermentation of inulin extract hydrolyzate primarily in the presence of monomers. It should be noted that C. beijerinckii NCIMB 8052 can effectively utilize the monomers, if the medium contained only monomers. Therefore, upon complete enzymatic hydrolysis of inulin, successful fermentation of inulin extract hydrolyzate was possible using this strain. The kinetics and performance of inulin extract hydrolyzate media by C. beijerinckii NCIMB 8052 can be related to C. acetobutylicum ATCC 824, which exhibited similar behavior in substrate utilization and solvent production similar to the former.

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Sample Inulin (DP 4+) Kestose Sucrose Glucose Fructose (g/L) (g/L) (g/L) (g/L) (g/L)

Guayule extract 2.56 0.00 0.00 0.27 0.36 TKS extract (Latex process) 11.38 0.068 0.248 0.74 1.01

TKS extract (Eskew process) 35.98 2.65 0 2.09 28.32

Chicory extract 41.83 5.95 6.21 3.53 19.45

Table 4.1 Amount of sugars present in raw inulin extract from different sources

Residual biomass of TKS and guayule Estimated Calorific Value (MJ/kg) (dry basis) TKS leaves 15.70 ± 0.12 TKS bagasse (Eskew process) 22.89 ± 0.16 TKS bagasse (Latex process) 18.13 ± 0.08 Guayule bagasse (Latex process) 22.06 ± 0.36 Whole guayule shrub 13.81 (Boateng et al., 2010) (after resin & rubber extraction) Guayule resin 37.90 (Coffelt et al., 2009) Bio-oil ~30.00 (Boateng et al., 2009)

Table 4.2 Energy density of residual TKS and guayule biomass

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Parameters Pure inulin P2 Raw Inulin Raw Chicory medium Eskew P2 P2 medium medium (control) Acetone (g/L) 1.71 ± 0.41 1.64 ± 0.06 2.80 ± 0.05 Ethanol (g/L) 1.67 ± 0.44 2.14 ± 0.04 1.36 ± 0.15 Butanol (g/L) 6.56 ± 0.50 4.88 ± 0.28 8.58 ± 0.11 Total ABE (g/L) 9.71 ± 0.57 8.48 ± 0.38 12.50 ± 0.13 Initial inulin (g/L) 55.78 ± 0.34 35.98 ± 0.01 41.83 ± 0.34 Initial kestose (g/L) 0.00 2.65 ± 0.07 5.96 ± 0.03

Initial sucrose (g/L) 0.00 0.00 6.21 ± 0.02 Initial glucose (g/L) 0.00 2.09 ± 0.11 3.53 ± 0.00 Initial fructose (g/L) 0.00 28.32 ± 0.29 19.45 ± 0.08

Initial amount of total sugars (g/L) 55.78 ± 0.34 69.03 ± 0.47 76.97 ± 0.47 Final inulin (g/L) 21.86 ± 4.20 31.55 ± 4.15 28.66 ± 3.22 Final kestose (g/L) 0.00 2.57 ± 0.13 4.14 ± 0.00

Final sucrose (g/L) 0.00 0.00 0.00 Final glucose (g/L) 0.00 1.97 ± 0.09 0.00 Final fructose (g/L) 4.80 ± 1.86 7.97 ± 0.25 3.65 ± 0.01 Final amount of total sugars (g/L) 26.82 ± 2.22 44.07 ± 2.36 36.45 ± 3.21 Total sugars utilized (g) 28.96 ± 2.22 25.20 ± 0.17 40.52 ± 3.67 ABE yield (g/g of substrate) 0.33 ± 0.04 0.33 ± 0.01 0.32 ± 0.03 ABE productivity (g/L/hour) 0.10 ± 0.01 0.12 ± 0.01 0.21 ± 0.01 Fermentation time (h) 108 84 84

Table 4.3 Performance and kinetic parameters of ABE production from unhydrolyzed inulin media by C. saccharobutylicum P262

209

Parameters Pure fructose P2 Hydrolyzed pure Hydrolyzed TKS Hydrolyzed medium inulin medium Eskew extract Chicory extract (control) Acetone (g/L) 5.46 ± 0.01 4.07 ± 0.17 2.01 ± 0.07 3.36 ± 0.54

Ethanol (g/L) 0.56 ± 0.13 2.02 ± 0.17 1.53 ± 0.08 1.55 ± 0.02 Butanol (g/L) 14.26 ± 0.20 9.21 ± 0.20 7.46 ± 0.15 8.93 ± 0.47

Total ABE (g/L) 20.18 ± 0.22 14.99 ± 0.41 10.00 ± 0.17 12.60 ± 0.99 Inulin (g/L) 0.00 0.00 13.49 ± 0.37 1.35 ± 0.17 Initial kestose (g/L) 0.00 4.48 ± 0.18 2.37 ± 0.08 5.90 ± 0.02 Initial sucrose (g/L) 0.00 9.80 ± 0.67 0.00 ± 0.00 0.00 Initial glucose (g/L) 0.00 5.70 ± 0.33 9.47 ± 0.13 12.84 ± 0.01

210 Initial fructose (g/L) 61.25 ± 0.98 40.50 ± 3.78 41.01 ± 0.17 41.84 ± 0.01 Initial amount of total sugars (g/L) 61.25 ± 0.98 60.48 ± 2.97 66.34 ± 0.75 61.93 ± 0.17 Final Inulin (g/L) 0.00 0.00 13.36 ± 0.35 1.36 ± 0.38 Final kestose (g/L) 0.00 4.45 ± 0.02 2.31 ± 0.20 5.26 ± 0.04 Final sucrose (g/L) 0.00 8.53 ± 0.07 0.00 0.00 Final glucose (g/L) 0.00 3.36 ± 0.23 1.67 ± 0.17 0.39 ± 0.11 Final fructose (g/L) 9.79 ± 0.58 6.32 ± 0.03 9.46 ± 1.27 9.56 ± 0.90 Final amount of total sugars (g/L) 9.79 ± 0.58 22.66 ± 0.29 26.80 ± 1.50 16.57 ± 1.14 Total sugars utilized (g) 51.46 ± 1.56 37.83 ± 1.68 40.58 ± 0.42 45.52 ± 1.54 ABE yield (g/g of substrate) 0.42 ± 0.01 0.41 ± 0.01 0.26 ± 0.01 0.30 ± 0.02 ABE productivity (g/L/hour) 0.42 ± 0.01 0.31 ± 0.01 0.17 ± 0.01 0.21 ± 0.02

Table 4.4 Performance and kinetic parameters of ABE production from 48 h enzymatically-hydrolyzed inulin extract media after 72 h fermentation by C. beijerinckii NCIMB 8052

Samples Inulin Kestose Sucrose Glucose Fructose

Before fermentation 0.00 16.08 ± 0.69 6.93 ± 1.38 3.78 ± 0.89 7.63 ± 2.62

After fermentation 0.00 10.11 ± 0.30 15.27 ± 0.42 0.86 ± 0.16 6.97 ± 0.22

Table 4.5 Amount of sugars present in 1 h enzymatically-hydrolyzed pure inulin medium before and after fermentation by Clostridium beijerinckii NCIMB 8052

Samples Inulin Kestose Sucrose Glucose Fructose

Before fermentation 0.00 19.00 ± 0.13 8.59 ± 0.01 4.87 ± 0.18 9.61 ± 1.61

After fermentation 0.00 12.04 ± 0.14 14.95 ± 0.08 1.78 ± 0.01 6.18 ± 0.02

Table 4.6 Amount of sugars present in 1 h enzymatically-hydrolyzed pure inulin medium before and after fermentation by Clostridium acetobutylicum ATCC 824

211

Samples Inulin Kestose Sucrose Glucose Fructose

Before fermentation 0.00 4.48 ± 0.18 9.80 ± 0.67 5.70 ± 0.33 40.50 ± 3.78

After fermentation 0.00 4.45 ± 0.02 8.53 ± 0.07 3.36 ± 0.23 6.32 ± 0.03

Amount of sugars 0.00 0.06 ± 0.12 1.27 ± 0.59 2.34 ± 0.11 34.18 ± 3.81 consumed

Table 4.7 Amount of sugars present in 48 h enzymatically-hydrolyzed inulin medium before and after fermentation by Clostridium beijerinckii NCIMB 8052

Samples Inulin Kestose Sucrose Glucose Fructose

Before 0.00 4.45 ± 0.04 9.89 ± 0.06 6.02 ± 0.02 41.42 ± 0.09 fermentation After 0.00 4.47 ± 0.06 8.57 ± 0.10 3.41 ± 0.02 6.44 ± 0.14 fermentation Amount of sugars 0.00 0.00 1.33 ± 0.05 2.61 ± 0.05 34.97 ± 0.05 consumed

Table 4.8 Amount of sugars present in 48 h enzymatically-hydrolyzed inulin medium before and after fermentation by Clostridium beijerinckii NCIMB 8052

212

Figure 4.1 Flow schemes of TKS processing and rubber extraction

213

Figure 4.2 Overview of inulin fermentation

214

TKS (Eskew) extract Chicory extract

Figure 4.3 Appearance of chicory and TKS extract

215

Figure 4.4 Effect of pH on endoinulinase, Novozyme 960 (Adapted from Novozyme)

216

217

Figure 4.5 Overview of inulinase production (Modified from Sangeetha et al., 2005)

Figure 4.6 Kestose hydrolysis pattern of C. beijerinckii NCIMB 8052

218

(a) 45

Inulin (50 IU/g) Inulin (100 IU/g) Inulin (150 IU/g) Glucose (50 IU/g) 30 Glucose (100 IU/g) Glucose (150 IU/g) Fructose (50 IU/g) Fructose (100 IU/g) Fructose (150 IU/g) 15 Concentration (g/L)

0 0 6 12 18 24 30 36 42 Time (h)

(b) 60

Inulin (50 IU/g) Inulin (100 IU/g) Inulin (150 IU/g) 40 Glucose (50 IU/g) Glucose (100 IU/g) Glucose (150 IU/g) Fructose (50 IU/g) Fructose (100 IU/g) Fructose (150 IU/g) 20 Concentration(g/L)

0 0 6 12 18 24 30 36 42 Time (h)

Figure 4.7 Inulin hydrolysis pattern at different enzyme concentrations using endo- inulinase (Novozyme 960) (a) TKS Eskew inulin, (b) Chicory inulin, (c) Commercial standard inulin

219

Figure 4.7 continued

(c) 60

Inulin (50 IU/g) 45 Inulin (100 IU/g) Inulin (150 IU/g) Glucose (50 IU/g) Glucose (100 IU/g) 30 Glucose (150 IU/g) Fructose (50 IU/g) Fructose (100 IU/g) Fructose (150 IU/g) Concentration(g/L) 15

0 0 6 12 18 24 30 36 42 Time (h)

220

45

Inulin (+ DP4) Kestose 30 TKS inulinTKS inulin Sucrose Glucose Fructose 60 15

Chicory inulin (g/L) Concentration

0 40 0 6 12 18 24 30 36 42 Time (h) 221 60 20 Commercial inulin Commercial inulin Concentration (g/L) Concentration

40 0 0 6 12 18 24 30 36 42 20 Time (h)

(g/L) Concentration

0 0 6 12 18 24 30 36 42 Time (h) Figure 4.8 Inulin hydrolysis and sugar production pattern at maximum enzyme concentration (150 IU/g of inulin) from different inulin source

(a)

5.10

5.05

5.00 TKS extract (50 IU/g) TKS extract (100 IU/g) 4.95 pH TKS extract (150 IU/g) Chicory extract (50 IU/g) Chicory extract (100 IU/g) 4.90 Chicory extract (150 IU/g)

4.85

4.80 0 12 24 36 Time (h)

5.2 (b) Pure inulin (50 IU/g) Pure inulin (100 IU/g) Pure inulin (150 IU/g)

4.8 pH

4.4

4.0 0 12 24 36

Time (h) Figure 4.9 Change in pH of (a) TKS and chicory inulin extract, (b) standard commercial inulin during hydrolysis

222

(a) 15

12

9

6 OD @ OD 600nm

3 10 g/L inulin 20 g/L inulin 30 g/L inulin 0 0 20 40 60 80 100

Time (h)

3 (b)

2

OD @ OD 600nm 1

10 g/L inulin 20 g/L inulin 30 g/L inulin 0 0 20 40 60 80 100 Time (h) Figure 4.10 Effect of substrate concentration on biomass concentration of (a) Kluyveromyces marxianus ATCC 16045, (b) Kluyveromyces marxianus ATCC 52466

223

8

(a) 10 g/L inulin 20 g/L inulin 30 g/L inulin 6

4

Inulinase activity (IU/ml) activity Inulinase 2

0 0 20 40 60 80 100 Time (h)

12

(b) 9

6

Inulinase activity (IU/ml) activity Inulinase 3 10 g/L inulin 20 g/L inulin 30 g/L inulin 0 0 20 40 60 80 100 Time (h) Figure 4.11 Effect of substrate concentration on production of (a) endoinulinase from Kluyveromyces marxianus ATCC 16045, (b) exo-inulinase from Kluyveromyces marxianus ATCC 52466

224

10 16 4 C. bei NCIMB 8052 C. bei NRRL B592 8 C. ace ATCC 824 C. sac P262 12

6 8 2 4 Butanol Butanol (g/L) Total ABE (g/L) ABE Total 4 (g/L) acid Acetic 2

0 0 0 0 24 48 72 96 120 0 24 48 72 96 120 0 24 48 72 96 120 Time (h) Time (h) Time (h) 225 5 3 6 4

2 3 4

2

Acetone (g/L) Acetone 1 Butyric acid (g/L) acid Butyric 2 1 @600 O.D nm

0 0 0 0 24 48 72 96 120 0 24 48 72 96 120 0 24 48 72 96 120 Time (h) Time (h) Time (h) Figure 4.12 Pure inulin fermentation using different strains of Clostridia species

12 Pure inulin (control) (a) Raw TKS extract Raw Chicory extract 9

6 Butanol (g/L)Butanol

3

0 0 24 48 72 96 Time (h)

(b) 12

9

6 Total Total (g/L)ABE

3

0 0 24 48 72 96 Time (h)

Figure 4.13 Fermentation of unhydrolyzed inulin extract media using C. saccharobutylicum P262 (a) butanol, (b) total ABE production

226

(a) 10 Pure inulin (control) Raw TKS extract 8 Raw Chicory extract

6

4 Aceticacid (g/L)

2

0 0 24 48 72 96 Time (h)

(b) 8

6

4 Butyric acid (g/L)Butyric 2

0 0 24 48 72 96 Time (h)

Figure 4.14 Fermentation of unhydrolyzed inulin extract media using C. saccharobutylicum P262 (a) acetic acid, (b) butyric acid

227

4 (a) Pure inulin (control) Raw TKS extract Raw Chicory extract 3

2 Acetone(g/L)

1

0 0 24 48 72 96 Time (h)

6.8 (b)

6.4

6.0 pH 5.6

5.2

4.8 0 24 48 72 96 Time (h)

Figure 4.15 Fermentation of unhydrolyzed inulin extract media using C. saccharobutylicum P262 (a) acetone production, (b) change in pH

228

(a) Inulin consumption pattern by C. saccharobutylicum P262 in pure inulin medium 60 Fructose Glucose Sucrose 45 Kestose Inulin (DP 4+ ) Inulin consumption pattern 30

15 Concentration (g/L)

0 0 12 24 36 48 60 72 96 108

Time (h)

Inulin consumption pattern by C. saccharobutylicum (b) P262 in raw chicory inulin medium 90 Fructose Glucose 75 Sucrose Kestose 60 Inulin (DP 4+) Inulin consumption pattern 45

30

Concentration (g/L) 15

0 0 12 24 36 48 60 72 84 Time (h)

Figure 4.16 Inulin consumption pattern by C. saccharobutylicum P262 (a) pure inulin P2 medium, (b) unhydrolyzed chicory inulin medium, (c) unhydrolyzed TKS Eskew inulin medium 229

Figure 4.16 continued

( c)

Inulin consumption pattern by C. saccharobutylicum P262 in raw TKS inulin medium Fructose 75 Glucose Sucrose ) 60 Kestose Inulin (DP 4+)

45

30

Concentration (g/L 15

0 0 12 24 36 48 60 72 84 Time (h)

230

20 25 6 C. bei NICMB 8052 C. bei NRRL B592 16 C. ace ATCC 824 20 C. sac P262 4 12 15

8 10

Butanol (g/L) Butanol 2 Total ABE (g/L) Total Acetic acid (g/L) Acetic

4 5

0 0 0 0 24 48 72 0 24 48 72 0 24 48 72 231 Time (h) Time (h) Time (h)

4 6 9

3

4 6

2

Acetone (g/L) Acetone 2 O.D @ 600 600 nm O.D @ 3 Butyric(g/L) acid 1

0 0 0 0 24 48 72 0 24 48 72 0 24 48 72 Time (h) Time (h) Time (h) Figure 4.17: Pure fructose fermentation using different strains of Clostridia species

20 Pure fructose (control) (a) Hydrolyzed Chicory extract Hydrolyzed TKS extract 16 Hydrolyzed pure inulin medium

12

8 Butanol (g/L)

4

0 0 24 48 72

Time (h)

(b) 25 Pure fructose (control) Hydrolyzed Chicory extract Hydrolyzed TKS extract 20 Hydrolyzed pure inulin medium

15

10 Total ABE (g/L)

5

0 0 24 48 72

Time (h)

Figure 4.18 Fermentation of 48 h enzymatically-hydrolyzed inulin extract media using C. beijerinckii NCIMB 8052 (a) butanol production, (b) total ABE production 232

6 (a)

5

4

3 cetic acid cetic (g/L) acid

A 2

Pure fructose (control) 1 Hydrolyzed Chicory extract Hydrolyzed TKS extract Hydrolyzed pure inulin medium 0 0 20 40 60

Time (h)

(b) 6

4

2 Butyric acid (g/L) acid Butyric

0 0 24 48 72 Time (h)

Figure 4.19 Fermentation of 48 h enzymatically-hydrolyzed inulin media using C. beijerinckii NCIMB 8052 (a) acetic acid, (b) butyric acid, (c) acetone production 233

Figure 4.19 continued

(c) 8 Pure fructose (control) Hydrolyzed TKS extract Hydrolyzed Chicory extract 6 Hydrolyzed pure inulin medium

4 Acetone(g/L)

2

0 0 24 48 72

Time (h)

234

6

4

O.D @nm600O.D 2

Pure inulin (control) Raw Chicory extract Raw TKS extract 0 0 20 40 60 80 100 Time (h)

Figure 4.20 Cell growth of C. saccharobutylicum P262 in unhydrolyzed inulin media

9 Pure fructose (control) Hydrolyzed TKS extract Hydrolyzed Chicory extract Hydrolyzed pure inulin medium

6

O.D @nm600O.D 3

0 0 24 48 72 Time (h)

Figure 4.21 Cell growth of C. beijerinckii NCIMB 8052 in 48 h enzymatically- hydrolyzed inulin media 235

48 h hydrolyzed pure inulin - C. bei NCIMB 8052 48 h hydrolyzed pure inulin - C. ace ATCC 824 1 h hydrolyzed pure inulin - C. bei NCIMB 8052 1 h hydrolyzed pure inulin - C. ace ATCC 824 10 20 6 8 16

6 4 12

4 8 Butanol Butanol (g/L)

Acetone (g/L) Acetone 2 Total ABE (g/L)Total ABE 2 4 236 0 0 0 0 24 48 72 0 24 48 72 0 24 48 72 Time (h) Time (h) Time (h)

6 4

4

4

2 2 2 O.Dnm 600 @ Acetic acid (g/L) acid Acetic Butyric acid (g/L)acid Butyric

0 0 0 0 24 48 72 0 24 48 72 0 24 48 72 Time (h) Time (h) Time (h)

Figure 4.22 Fermentation of 48 h and 1 h enzymatically-hydrolyzed pure inulin media

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282

APPENDIX A: DESCRIPTION OF FOOD PROCESSING WASTES

AND MODES OF UTILIZATION BY FOOD PROCESSING

INDUSTRIES

283

Industry Type of waste Current disposal methods Cabbage trimmings are returned to the farm with the grower delivering cabbage. It is then spread on the field The Fremont 1. Cabbage trimmings for composting. Though it is little Company 2. Sauerkraut juice inconvenient for the grower, but it is a 802 N Front street, relatively an inexpensive option. Also, Fremont, cabbage trimmings are supplied to OH 43420-1917 local cattle producers.

Sauerkraut juice is disposed through the city water treatment facility. Since this juice has very high BOD level, disposing them to the specific requirements of water treatment plant makes this as a very expensive disposal method. Potato waste is sent to a farm and Ballreich Brothers composted into fertilizer. Inc. 1. Potato waste products 186 Ohio Avenue, 2. Soybean Oil shortening Soybean oil shortening waste is used Tiffin, OH 44883 wastes to enrich animal feed in Wapakoneta.

Sensus LLC Waste tea leaves It is sent to a composting company. 7255 Hamilton Enterprise park Dr. Hamilton, OH 45011

Table A1 Types of food processing wastes and modes of utilization by food processing industries

284

Table A1 continued

Industry Type of waste Current disposal methods

Earlier Attempt to use spent wastes for land application (soil remediation) was not 1. Wet celery spent successful because of some of the inherent 2. Dry celery spent compounds present in them. Currently, these Joseph Adams Corp. 3. Parsley spent wastes are being shipped to another county 5740 Grafton Road, 4. Oregano spent for disposal (means of disposal is not known). Valley City, 5. All spice spent OH- 44280-9327 6. Fennel spent Setting up of gas generator to use these wastes was considered not cost effective by the company.

Possibility of using the wastes as feed supplement necessitates few changes in the manufacturing processes. Breading is applied directly to a landfill. Lake Erie Frozen foods CO (Inc.) 1. Breading Batter liquid is being shipped to Endres 1830 Orange Road, 2. Batter liquid processing, LLC to mix with other food waste Ashland, 3. Fryer siftings streams and used to prepare livestock feed. OH 44805-1335 Disposal of fryer sifting is not provided. Animal feed and whey streams are used as The Dannon Company feed for hogs. Inc. 1. Animal feed 216 Southgate Drive, 2. Whey liquid waste WWTP sludge is land filled as fertilizer. Minster, OH 45865 3. WWTP sludge Using it in an offsite anaerobic digester is proposed but has not yet been commissioned.

285

Table A1 continued

Industry Type of waste Current disposal methods

Cardboard, Dough, cheese, pizza for used for mulching 1. Uncooked meat 2. Cooked meat Bone and fat meat wastes are 3. Inedible cheese used for making animal feed 4. Inedible dough ConAgra Foods Inc 5. Inedible pizza Wood & metal remnants are 801 Dye Mill Road, 6. Waste Water solids used to prepare Pallets Troy, OH 45373-4223 7. Waste water 8. Inedible sauce Waste Water & Sludge is 9. Card board waste incinerated and land applied 10. Metal & wood remnants Metal and plastic leftover are recycled by a local recycler

The rest of the wastes are compacted and sent to landfills. Bellisio foods 1. Liquid waste All these sludge wastes are 100 E Broadway street, 2. Digester bottom sludge land applied and used as Jackson, 3. Digester floating sludge fertilizer OH 45640-1347 4. Landfill food waste

286

Table A1 continued

Industry Type of waste Current disposal methods

1. Tomato peel These wastes are locally 2. Tomato pulp disposed of in compost pile Pilot plant, Parker food 3. Chipped raw potato located in the campus. science & technology 4. Coriander lab 5. Minced pepper 110 Parker Food Science 6. Raw pepper and Technology, 7. Onion peel 2015 Fyffe Ct, 8. Fried donut waste Columbus, OH 43210 9. Donut wet mix 10. Donut dry mix 11. Raw potato 12. Beans 13. Canned broccoli waste 14. Canned tomato juice waste Cargill salt 1. Calcium sulfate waste stream -Information not provided- 2065 Manchester Road 2. Coal boiler ash Akron, OH 44314-1770 International dairy Milk dust powder -Information not available- ingredients, Inc. 625 Commerce Road, Wapakoneta, OH 45895-8265

287

Type of waste Description of waste

Cabbage trimmings These solid and liquid wastes are gathered during sauerkraut Sauerkraut juice preparation from cabbage.

This primarily contains peelings and potato, dirt from washing Potato waste products process, and unsatisfactory potato chips.

Soybean Oil shortening wastes These are remnants after oil extraction from soybean that mostly contains small chip pieces and large grease trap.

Waste tea leaves This waste occurs during pulverization and packaging of tea leaves. Wet celery spent These are residues after extraction of flavor and aroma through Dry celery spent steam distillation from the ground spices. Parsley spent Oregano spent All spice spent Fennel spent

 Batter liquid is basically liquid dough made up of starch Batter liquid flour and seasonings. This waste is gathered as a thick

sludge at the bottom of the batter machine.

 Breading is dry coarse flour along with seasonings and Breading flavors (used as coatings in fried foods). This breading

material is gathered as waste from machine and floors.

 Siftings are small-sized solid particles which are leftover Fryer siftings materials from frying. This is an oily and fat-rich waste.

Table A2 Description of food processing wastes 288

Table A2 continued

Type of waste Description of waste

It is a liquid waste remnant from animal feed Animal feed processing plant.

Whey liquid is generated during cheese Whey liquid waste making.

WWTP sludge (biosolids) is a semisolid WWTP sludge produced from the treatment of wastewater.

Uncooked meat Meat and cheese wastes are produced during Cooked meat processing and packaging of products. Inedible cheese Inedible dough represents over-fermented, Inedible dough unusable and leftover dough from the processing line. Inedible pizza Inedible sauce Inedible pizza and sauce are leftover or spoiled products.

Waste water and solids comprises liquid and Waste Water solids solid wastes generated in a food processing Waste water plant. These wastes are collected from a digester tank.

Card board waste Card board wastes are produced in the packaging section.

289

Table A2 continued

Type of waste Description of waste

Liquid waste Liquid waste is obtained from wastewater treatment plant.

Landfill food waste primarily contains solid food Landfill food waste wastes generated in a food processing facility.

Digester bottom and floating sludge are wet semi- solid wastes generated during anaerobic digestion Digester bottom sludge of food processing wastes Digester floating sludge Tomato peel These vegetable wastes are produced during pre- Tomato pulp processing of vegetables. Chipped raw potato Coriander Minced pepper Raw pepper Onion peel Raw potato Beans Canned broccoli waste Canned tomato juice waste

Donut dry mix These are formulated donut mix flour waste Donut wet mix collected from a plant.

Fried donut waste It is mostly oil substances which includes siftings and unfinished or overcooked fried donuts.

290

Table A2 continued

Type of waste Description of waste

Calcium sulfate waste stream Calcium sulfate is used as seed to create salt crystals during salt production from saturated brine. This waste is

accumulated in the evaporator.

Coal boiler ash This bottom ash is produced in large quantities and this is the most common type of ash produced in coal boilers. Nearly 12% of coal is burned up as bottom ash. Milk dust powder This powdered milk waste is collected in spray driers during

manufacturing of dried milk products.

291

APPENDIX B: CALCULATIONS ON AMOUNT OF SUBSTRATE

REQUIRED TO MAKE 50 G/L STARCH MEDIUM

292

Substrate % TS Starch Amount of substrate required (in Starch content in the g) to make 50 g/L starch medium (g) = ((% (% of dry weight) TS/100)×(% starch/100)×Amount of substrate) Breading 90.47 60.97 92.00 50.75

293 Batter liquid 32.96 62.08 248.00 50.74

Inedible dough 63.05 60.38 133.00 50.63

Table B1 Calculations on amount of substrate required to make 50 g/L starch medium

APPENDIX C: CELL GROWTH (COLONY FORMING UNIT PER ML)

OF CLOSTDRIA IN FOOD PROCESSING WASTES FERMENTATION

MEDIA

294

Glucose control Inedible dough Breading Batter liquid Milk Dust Powder Time (h) 0 5.10×107 ± 0.47 6.70×107 ± 0.93 7.20×107 ± 0.30 5.20×107 ± 0.50 1.17×107 ± 0.35 12 2.00×109 ± 0.3 2.50×109 ± 0.20 3.10×109 ± 0.49 2.80×109 ±0.72 1.27×108 ± 0.15 24 3.30×109 ± 0.49 5.30×109 ± 0.61 7 00×109 ± 0.7 7.00×109 ±0.7 4.67×108 ± 2.52 10 10 10 10 9 36 1.10×10 ±0.1 3.50×10 ±0.26 3.90 × 10 ±0.6 1.70×10 ± 0.15 3.67×10 ± 1.53 48 6.00×109 ± 0.15 2.70×1010 ±0.5 2.20 × 1010 ±0.30 2.30×1010 ± 0.36 2.33×109 ± 0.71 60 8.60×109 ± 0.11 2.10×1010 ± 0.30 1.60 × 1010 ±0.25 2.80×1010 ±1.15 1.30×108 ± 0.58 72 1.20×109 ± 0.2 1.30×109 ± 0.20 2.60× 109 ±0.45 1.9 0×109 ±0.55 1.60×108 ± 0.58

295 Table C1Growth of C. beijerinckii NCIMB 8052 (CFU/ml) in food processing wastes media

Glucose control Inedible dough Breading Batter liquid Milk Dust Powder Time (h) 0 2.86×107 ± 0.61 4.36×107 ± 1.35 4.13×107 ± 0.61 5.40×107 ± 0.62 1.80×107 ± 0.26 12 8.06×109 ±0.49 1.00×1010 ± 1.3 6.53×109 ±0.81 7.30×109 ±0.75 3.10×108 ± 0.21 24 4.46×1010 ±0.30 3.10×1010 ±1.2 5.53×1010 ±0.47 1.66×1010 ±0.42 4.00×109 ± 2.0 36 1.39×1011 ±1.69 7.10×1010 ±0.47 6.50×1010 ±0.81 8.86×1010 ±1.51 7.67×109 ± 4.04 48 8.30×1010 ±0.40 3.50×1010 ±0.95 5.20×1010 ±1.4 8.23×1010 ±0.51 1.03×109 ± 0.21 60 6.10×1010 ±0.3 5.53×1010 ±0.57 6.23×1010 ±0.49 5.63×1010 ±0.97 1.13×109 ± 0.30 72 6.90×109 ±0.89 3.66×109 ±0.42 3.30×109 ±0.89 4.56×109 ±0.50 7.67×108 ± 4.36

Table C2 Growth of C. acetobutylicum ATCC 824 (CFU/ml) in food processing wastes media

APPENDIX D: HPLC CHROMATOGRAMS OF UNHYDROLYZED

INULIN MEDIUM BEFORE FERMENTATION

296

RID1 A, Refractive Index Signal (003-0302.D) nRIU 50000

40000 6.949 - Inulin6.949 - 30000 10.828 - Fructose 10.828 -

20000 8.796 13.696 10000 8.176 7.707 - Kestose - 7.707 9.879 9.188 14.071 10.085 - Glucose 10.085 - 11.606 3.524 11.902 0.907 14.944 1.854 13.000 0 5.299 0 2 4 6 8 10 12 14 min

Figure D1 Raw TKS extract before fermentation

RID1 A, Refractive Index Signal (004-0401.D) nRIU

50000

40000 6.966 - Inulin6.966 - 8.794

30000 Fructose 10.807 -

20000 8.452 - Sucrose -8.452

10000 Kestose - 7.636 13.688 8.145 9.877 - Glucose - 9.877 9.163 6.743 3.509 0.876 11.945 1.818 0 5.904 14.913 0 2 4 6 8 10 12 14 min

Figure D2 Raw chicory extract before fermentation

RID1 A, Refractive Index Signal (001-0101.D) nRIU 120000

100000

80000 Inulin6.932 -

60000

40000

20000 8.081 8.384 Sucrose - 9.580 9.810 Glucose - Fructose 10.811 - 13.712 0 0.284 0.439 0.673 0.756 0.932 1.091 1.372 1.487 1.721 1.915 2.146 2.293 2.470 2.732 2.869 3.087 3.251 3.419 3.784 3.882 4.061 4.388 4.536 4.676 4.766 5.022 5.188 5.317 5.400 5.753 5.839 11.970 12.107 12.239 12.704 13.039 13.187 14.252 14.483 14.951 0 2 4 6 8 10 12 14 min

Figure D3 Pure inulin medium before fermentation by C. saccharobutylicum P262

297

APPENDIX E: HPLC CHROMATOGRAMS OF UNHYDROLYZED

INULIN MEDIUM AFTER FERMENTATION

298

RID1 A, Refractive Index Signal (030-3001.D) nRIU

40000

30000 Inulin6.948 -

20000 8.795

10000 Fructose 10.885 - 8.175 13.694 7.707 - Kestose - 7.707 9.187 14.071 10.114 - Glucose 10.114 - 1.761 3.610 9.679 14.945 0.867 11.915 0 4.993 5.418 5.828 0 2 4 6 8 10 12 14 min

Figure E1 Raw TKS extract after 72 h fermentation by C. saccharobutylicum P262

RID1 A, Refractive Index Signal (031-3101.D) nRIU

35000 30000 8.796

25000 Inulin6.961 - 20000 15000 10000 7.622 - Kestose - 7.622 10.806 - Fructose 10.806 - 8.133

5000 9.157 1.826 3.678 10.106 - Glucose -10.106 9.603 0.928 11.637 13.295 13.735 12.407 14.906 14.350 0 5.342 5.512 0 2 4 6 8 10 12 14 min

Figure E2 Raw Chicory extract after 72 h fermentation by C. saccharobutylicum P262

RID1 A, Refractive Index Signal (003-0301.D) nRIU 100000

80000 6.935 - Inulin6.935 - 60000

40000

20000 10.811 - Fructose 10.811 - 8.102 8.411 - Sucrose 8.411 - 3.770 9.151 9.587 0 1.015 2.308 2.513 2.606 2.862 4.594 4.686 4.820 5.612 5.821 11.754 12.773 13.036 13.134 13.717 14.466 14.946 0 2 4 6 8 10 12 14 min

Figure E3 Pure inulin medium after fermentation by C. saccharobutylicum P262 299

APPENDIX F: HPLC CHROMATOGRAMS OF 1 H

ENZYMATICALLY-HYDROLYZED PURE INULIN MEDIUM BEFORE

AND AFTER FERMENTATION

300

RID1 A, Refractive Index Signal (003-0301.D) nRIU 25000

20000 7.363 15000 7.766 - Kestose 7.766- 10.887 Fructose -

10000 Sucrose 8.665-

5000 Glucose 9.961- 14.229 8.157 16.478 0.580 0.788 0.997 1.354 2.236 2.992 3.287 3.517 3.733 4.061 4.352 4.511 6.791 13.100 15.005 0 4.705 4.846 4.993 5.231 5.377 5.591 5.882 5.985 6.361 6.480 18.140 19.519 19.676

0 5 10 15 min

Figure F1 1 h enzymatically-hydrolyzed pure inulin medium before fermentation by C. beijerinckii NCIMB 8052

RID1 A, Refractive Index Signal (019-1901.D) nRIU

15000 7.839 - Kestose 7.839- 10000 Sucrose 8.761- 7.432

5000 2.267 14.415 8.248 10.987 - Fructose - 10.987 9.760 0.873 16.659 11.869 12.604 13.319 4.270 4.453 4.802 4.943 5.177 5.562 5.850 6.052 6.865 0 6.397 18.256 19.199 19.429 19.602

0 5 10 15 min

Figure F2 1 h enzymatically-hydrolyzed pure inulin medium after 72 h fermentation by C. beijerinckii NCIMB 8052

301

APPENDIX G: HPLC CHROMATOGRAMS OF 48 H

ENZYMATICALLY-HYDROLYZED PURE INULIN MEDIUM BEFORE

AND AFTER FERMENTATION

302

RID1 A, Refractive Index Signal (005-0501.D) nRIU 50000

40000

30000 10.785 - Fructose 10.785 - 20000

10000 8.574 - Sucrose -8.574 7.683 - Kestose - 7.683 9.867 - Glucose - 9.867 7.283 8.140 16.310 14.088 6.732 13.664 0.594 0.706 0.846 1.012 1.159 1.871 2.909 3.042 3.139 3.337 3.510 3.672 3.823 3.978 4.122 4.230 4.550 4.666 4.763 5.036 5.234 5.504 5.759 5.864 6.030 6.174 17.980 19.163 19.480 19.595 19.876 0 4.936

0 5 10 15 min

Figure G1 48 h enzymatically-hydrolyzed pure inulin medium before fermentation by C. beijerinckii NCIMB 8052

Figure G2 48 h enzymatically-hydrolyzed pure inulin medium after 72 h fermentation by C. beijerinckii NCIMB 8052

303

APPENDIX H HPLC CHROMATOGRAMS OF ENZYMATICALLY

HYDROLYZED INULIN EXTRACTS

304

RID1 A, Refractive Index Signal (001-0102.D) nRIU

50000

40000

30000 -Fructose 10.818 6.910Inulin - 20000 Sucrose - 8.776 13.691 14.076

10000 Glucose - 9.868 8.167 9.184 7.726 - Kestose - 7.726 11.601 3.742 11.896 1.013 14.933 1.958 2.840 0 5.562 5.814 0 2 4 6 8 10 12 14 min

Figure H1 48 h enzymatically-hydrolyzed TKS Eskew Extract using endo-inulinase (Novozyme 960)

RID1 A, Refractive Index Signal (002-0201.D) nRIU

60000 50000 40000 10.810 - Fructose 10.810 - 30000 8.778 Sucrose - 20000 9.881 Glucose -

10000 Kestose 7.721 - 14.084 13.692 8.176 9.165 7.273 6.941 - Inulin - 6.941 3.665 6.750 11.924 0.936 1.883 0 2.639 2.822 14.926 0 2 4 6 8 10 12 14 min

Figure H2 48 h enzymatically-hydrolyzed Chicory Extract using endo-inulinase (Novozyme 960)

305

APPENDIX I HPLC CHROMATOGRAM CALIBRATION

STANDARDS

306

RID1 A, Refractive Index Signal (ASHOK\INU KES SUC FRUGLU 071512 2012-07-15 20-38-43\001-0101.D) nRIU 250000

200000 7.049 - Inulin - 7.049 150000 10.020 - Glucose 10.020 - 10.950 - Fructose 10.950 -

100000 Sucrose -8.567

50000 Kestose - 7.718

0 0.277 0.450 0.630 0.731 1.026 1.220 1.343 1.444 1.577 1.890 1.984 2.245 2.362 2.552 2.725 2.948 3.256 3.398 3.503 3.607 3.769 3.856 4.158 4.324 4.568 4.705 4.835 4.986 5.144 5.334 5.425 5.609 5.749 6.286 13.607 16.240 16.355 16.456 18.227 19.343 19.566 19.670 19.904 0 5 10 15 min

Figure I1 HPLC chromatogram of standard sample containing inulin, kestose, sucrose, glucose and fructose (25g/L inulin; 5 g/L kestose; 10 g/L sucrose; 20 g/L glucose; 25 g/L fructose )

(a) Inulin standard 4000000

3000000 R² = 0.9995

2000000

Area 1000000

0 0 5 10 15 20 25 30 Amount (g/L)

Figure I2 HPLC calibration standard curves (a) inulin, (b) kestose, (c) sucrose, (d) glucose, (e) fructose

307

Figure I2 continued

(b) Kestose standard 800000

600000 R² = 0.9981 400000 Area 200000

0 0 1 2 3 4 5 Amount (g/L)

(c) Sucrose standard

1600000

1200000

800000 Area Area R² = 0.9966 400000

0 0 2 4 6 8 10 12 Amount (g/L)

308

Figure I2 continued

Glucose standard (d) 4000000

3000000

Area Area 2000000 R² = 0.9986 1000000

0 0 5 10 15 20 Amount (g/L)

( e) Fructose standard 5000000

4000000

3000000 Area Area 2000000 R² = 0.9981

1000000

0 0 5 10 15 20 25 Amount (g/L)

309