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-ethanol (ABE) fermentation from diverse feedstocks.
Butanol could occupy a significant portion of advanced biofuel 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 solvents 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
vii
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 Biofuels ...... 13
viii
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 Clostridium 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 Lignocellulosic biomass ...... 30
2.3.2 Glycerol 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 Carbon (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 fuels ...... 71
Table 2.2 Comparison of properties of n-butanol with its isomers ...... 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
xix
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
xx
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 alcohol) is a four carbon straight chain primary alcohol 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 solvent 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 fuel 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 carbon dioxide, 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.
4
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, renewable energy 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 fossil fuel 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 alcohols, 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 fermentations, 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, Hydrogen)
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
transesterification 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 methanol 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
(hydroformylation) 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.
21
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-carbons 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 oxygen 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 isomer 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. Butyraldehyde 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 cellulose 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 oleyl alcohol/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
50
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
54
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 carbonyl group 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
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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
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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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