Iowa State University Capstones, Theses and Retrospective Theses and Dissertations Dissertations

2006 Recovery of monosaccharides via catalytic hydrolysis Jason Alan Bootsma Iowa State University

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This Dissertation is brought to you for free and open access by the Iowa State University Capstones, Theses and Dissertations at Iowa State University Digital Repository. It has been accepted for inclusion in Retrospective Theses and Dissertations by an authorized administrator of Iowa State University Digital Repository. For more information, please contact [email protected]. Recovery of monosaccharides via catalytic hydrolysis

by

Jason Alan Bootsma

A dissertation submitted to the graduate faculty

in partial fulfillment of the requirements for the degree of

DOCTOR OF PHILOSOPHY

Major: Chemical Engineering

Program of Study Committee: Brent H. Shanks, Major Professor Peter J. Reilly R. Dennis Vigil Lawrence A. Johnson D. Raj Raman

Iowa State University

Ames, Iowa

2006 UMI Number: 3229053

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or the M Pr r iii

TABLE OF CONTENTS

ACKNOWLEDGEMENTS v

ABSTRACT vi

CHAPTER 1: General Introduction 1

1.1 Introduction 1

1.2 Lignocellulose Structure 1

1.3 Utilization 3

1.4 Hydrolysis 4

1.5 Feedstock Supply 7

1.6 Motivation 9

References 10

CHAPTER 2: Hydrolysis Characteristics of Tissue Fractions 13 Resulting From Mechanical Separation of Corn Stover

2.1 Introduction 13

2.2 Materials and Methods 16

2.3 Results and Discussion 18

2.4 Conclusion 25

References 25

CHAPTER 3: Disaccharide Hydrolysis Using Organic-Inorganic 38 Hybrid Mesoporous Silica Catalysts

3.1 Introduction 38

3.2 Experimental 41 iv

3.3 Results and Discussion 45

3.4 Conclusion 51

Acknowledgements 52

References 52

CHAPTER 4: Monosaccharide Degradation Kinetics during Organic-Inorganic 64 Mesoporous Silica Catalyzed Hydrolysis Conditions

4.1 Introduction 64

4.2 Experimental 66

4.3 Results and Discussion 68

4.4 Conclusion 74

References 75

CHAPTER 5: Hydrolysis of Oligosaccharides from Distiller's Grain Using Organic- 89 Inorganic Hybrid Mesoporous Silica Catalysts

5.1 Introduction 89

5.2 Experimental 90

5.3 Results and Discussion 92

5.4 Conclusion 95

References 96

CHAPTER 6 Summary and Future Investigations 108

6.1 Summary 108

6.2 Future Investigation 110

References 112 V

ACKNOWLEDGEMENTS

I would like to thank my major professor, Brent Shanks, who has been a source of support, guidance, and encouragement. Without his direction this work would not have been possible. I wish to express my thanks to my committee members Peter Reilly, Dennis Vigil,

Larry Johnson, and Raj Raman for their suggestions and advice. I would also like to thank

Rob Anex and Tom Richard who also served as members on my committee. I would like to thank the members of my research group for their friendship and support during my time at

Iowa State.

I would like to thank the United States Department of Agriculture through the Iowa

Biotechnology By-products Consortium, the Centers for Crops Utilization Research, the

Corn Refiners' Association, The National Science Foundation, and the United States

Department of Energy through the Midwest Consortium for their generous financial support of this research.

Most importantly, I would like to thank my family for their love and support. I would

like to especially offer a thank you to my wife, Holly, who agreed to become a graduate

student's wife and make many sacrifices to help me reach my goals. vi

ABSTRACT

This research has provided a fundamental analysis of the use of novel separation and catalyst techniques for saccharification of lignocellulosic tissues. Separation techniques focused on providing a feedstock that was more amenable to hydrolysis. Catalyst development focused on solid acid materials that would provide processing advantages, cost reductions, and waste reduction. A mechanical separation technique was used on corn stover to separate the parent material into two streams with improved processing characteristics.

The desire was to produce a material that was easier to hydrolyze in addition to producing a material with enhanced fiber character. The separation technique was effective; however, the material streams did not have significantly different hydrolysis characteristics, which made further development of this technique unattractive. A second separation process that was evaluated was the hydrothermal treatment of distiller's dry grain (performed at Purdue

University). This treatment was effective in producing a stream of soluble oligosaccharides that were amenable to hydrolysis to monosaccharides. Acid-functionalized mesoporous silica was synthesized and tested in a model system as a cellobiose hydrolysis catalyst, as well as a catalyst for the hydrolysis of solubilized oligosaccharides generated from distiller's dry grain hydrothermal treatment. These materials were also evaluated with monosaccharides to determine the catalyst activity towards undesirable degradations reactions. The acid-functionalized silica materials were effective catalyst materials with performance similar to homogeneous catalyst materials. These materials were robust enough to be utilized in real systems without severe reduction in catalyst activity. Acid- functionalized silicas were particularly attractive due to low degradation rates of desirable vii products. Distinct pH controlled kinetic regimes were shown to be responsible for the low degradation rates and these materials produced ideal pH for desirable degradation kinetics.

The chemistry of aqueous phase systems was discussed and several key attributes (leveling, proton mobility, and the ion product) were identified as critical considerations for future catalyst development for use in aqueous systems. 1

CHAPTER 1

General Introduction

1.1. Introduction

Every day plants are converting the energy of the sun into materials collectively referred to as biomass. Biomass has been defined as organic material of recent biological origin. Biomass is important because it can be used to provide food, fuel and goods while being the only truly renewable source of organic fuels and chemicals [1,2]. Fossil fuels are differentiated from biomass because these resources require extremely long time periods to develop and therefore are not renewable or sustainable.

Some types of biomass have been easier to use than others. Biomass consists of three major classes of materials: starches, oils and lignocellulose. Starches and oils are readily digested by people and animals, and directly or indirectly provide almost all dietary calories.

These materials are also used to produce a variety of chemicals and fuels. However, lignocellulose makes up an estimated 50% of the biomass in the world and only a small fraction of it is used for materials or fuels [2, 3]. The limited utilization of lignocellulose is due to the challenges its complex structure presents.

1.2. Lignocellulose Structure

Lignocellulose is a composite of three classes of polymers: cellulose, hemicellulose, and lignin [3], Cellulose is the most abundant polymer on earth. It is a linear homopolymer of anhydroglucopyranose. The anhydroglucopyranose units are connected through (3-(l-4) linkages. When two glucose units are bonded, a molecule of water is liberated. This bond requires the rotation of the second glucose molecule about the C1-C4 axis. The resulting 2

H OH

HO

OH

H H

Figure 1: Cellobiose structure.

disaccharide is the repeating unit of cellulose referred to as cellobiose (Figure 1). The linear

nature of the (3-(l-4) glycosidic bond, the stable nature of the pyranose conformation, and the

large degree of hydrogen bonding, result in the very stable secondary and tertiary structure of

cellulose. These features in addition to its close association with the other plant polymers

make cellulose very resistant to hydrolysis [4], Hemicellulose is a complex branched-chain

polymer of a variety of hexoses and pentoses linked by a variety of glycosidic bonds. The

principal components of hemicellulose are xylose and arabinose, but there are a number of

minor components. The composition and complexity of hemicellulose varies with its source.

Due to its heteropolymeric character and lack of crystallinity hemicellulose is easier to

hydrolyze than cellulose. Lignin is a complex phenolic polymer that has a waxy consistency

and coats the cellulosic and hemicellulosic strands. This complex structure makes

lignocellulose very resistant to degradation by microbes as well as chemical or physical

disruption [2, 5-7].

Lignocellulosic biomass originates from a wide variety of sources, resulting in a

broad spectrum of characteristics [3,8]. The materials that currently are under utilized are

wastes, crop residues and processing by-products. These materials do not have desirable

fiber characteristics to be used for paper products or building materials [9]. They typically 3

. have very low animal feed value and have too high a moisture content to provide energy

through combustion.

1.3. Utilization

Attempts have been made to convert lignocellulose into fuels by using thermal

processing techniques such as gasification, pyrolysis, liquefaction, and supercritical fluid

extraction. These techniques are effective disposal strategies, but they currently do not

provide fuels that are cost competitive with fossil sources. Fuels resulting from these

processes are typically of low quality and unstable [10].

A popular approach to the utilization of these materials is saccharification [1, 2, 4, 8,

9, 11-19], Saccharification is the hydrolysis of the polysaccharides into monosaccharides.

The resulting monosaccharides can be used in a variety of applications; the most common is

fermentation to ethanol. Saccharification can be done with enzymes or acids. The acid-

catalyzed process has a much longer history operating since the early 1900's on industrial

scale [20]. These plants used sulfuric acid-catalyzed saccharification of southern pine to

make 95% ethanol, but eventually closed down due to economic conditions and competition

from the cheaper blackstrap molasses process. Interest and investment in these processes has

peaked and fallen several times in the last century, resulting in sporadic research, but many

of the same issues remain. A variety of techniques have been applied to this problem, but

none have provided the compelling economics required for continued industrial application

[8,18].

Several approaches have been developed to deal with the complex structure of

lignocellulose. Several reviews of techniques have been conducted. Three recent reviews by

Mosier et al. [19], Lin and Tanaka [21], and Hsu [19, 21, 22] summarized the more 4 promising technologies. I will briefly mention some of the techniques that have been used.

Most of these strategies involve a multi-step approach. The first step is typically size reduction through a grinding technique [5, 6, 8, 13, 15, 23]. The next step is typically referred to as a pretreatment. This can be cooking in steam[8, 24, 25], hot water[12, 24-26], sulfuric acid[5, 12, 13, 15, 17, 23, 27-29], phosphoric acid[13, 15], hydrochloric acid [23,

30], ammonia [7, 8, 31, 32], or calcium hydroxide [12, 25, 33]. These pretreatment steps result in a variety of products including delignified cellulose [7, 31], monosaccharides [1,5,

15, 27], oligosaccharides[24, 26], and a variety of degradation products[34]. The pretreatment steps that leave polysaccharides and oligosaccharides are typically followed by enzymatic hydrolysis to reduce the remaining polysaccharides to monosaccharides[12, 13,

16, 25, 26]. Other methods for treating biomass have focused on unique reactor designs.

These designs include plug-flow reactors, percolation reactors, progressive batch reactors, counter-current reactors, co-current reactors, and shrinking bed reactors [18].

1.4. Hydrolysis

The most widely accepted mechanism for the acid-catalyzed hydrolysis of the glycosidic bond is a three step process. The reaction begins with protonation of the glycosidic oxygen, also referred to as an A-1 mechanism [35]. This protonation is followed by the breaking of the bond, causing the formation of a non-reducing end-group and an unstable carbonium ion. The second step is believed to be slow and rate limiting [36]. The final step is the carbonium ion reacting with water to form a reducing end-group, the new hemiacetalic hydroxyl. The reason the water adds on the hemiacetal end of bond is because the hemiacetal has greater electron density, leading to a more stable carbonium ion [4], The

A1 mechanism is favored because the rate of reaction correlates with the acid concentration 5 of the solution [37]. Therefore the limiting step is the protonation of the glycosidic oxygen and not the reaction with water [35, 37].

There is some disagreement over the strength of acidity needed to accomplish hydrolysis. Abatzogluo [4] states that "organic acids are not strong enough to give useful commercial hydrolysis of cellulose." Mosier et al. [38] has found that carboxylic acids hydrolyze cellobiose while offering a much better selectivity due to the lower amount of

glucose degradation. They also Cat.

OH HO found that maleic acid would OH

HO hydrolyze cellulose achieving a HO OH HO vOH greater glucose yield due to lack of

degradation [38]. The use of

JO carboxylic acids is justified by the

Cat. study of the reactive domain of

cellulase enzymes. These enzymes

OH use glutamic and aspartic acid OH

HO residues as the catalysts for the HO HO OH HO OH hydrolysis reaction [39]. These OH OH HO .0 residues perform the reaction by one

of two mechanisms: retaining and Figure 2: Enzymatic hydrolysis of cellobiose via the inverting mechanism. inverting. The inverting method uses the charged site to activate a water molecule for nucleophilic attack on the glycosidic oxygen. The opposite amino acid provides the needed proton. The retaining mechanism uses a direct nucleophilic attack by the amino acid to form a covalently bound intermediate. The opposite amino acid activates a

water molecule for a nucleophilic

attack that frees the hydrolysis

product and recharges the proton

donor [39]. Cellulase uses the

close proximity of a proton donor

and a proton acceptor to achieve

the reaction with limited acid

strength.

The starting point in

modeling acid-catalyzed

saccharification of lignocellulosic

biomass is to determine what

material will be used as the model

system. Models have been done Figure 3: Enzymatic hydrolysis of cellobiose via the retaining mechanism. starting with everything from wood

[4, 40, 41] to cellobiose [35, 42]. The reactivity of the glycosidic bond in wood will be much different than the reactivity of the glycosidic bond in cellobiose, even if the bonds are chemically identical. Accessibility differences occur due to the tertiary structure of cellulose and the associated polymers of hemicellulose and lignin compared to the soluble nature of cellobiose. Models that begin with wood or other lignocellulosic biomass have been developed dating back to the beginnings of the process [41]. Models that start with complex structures are typically relegated to empirical fits and correlations due to lack of 7 understanding of several steps that occur with homogeneous hydrolysis. These models do provide some useful kinetic information that can be used for scale-up and prediction of hydrolysis conditions even without precise knowledge of the mechanism. This has led to what is referred to as pseudokinetic modeling [4] of these systems. The use of severity factors and other correlations allow for more widespread application of these models and provide a method for dealing with the complexity of these systems [43]. The alternative route is to use simple model compounds as a basis for the model [35,38]. This method simplifies the model to a single type of bond and a single type of molecule, and allows investigations into the hydrolysis reaction without the added complexity of a heterogeneous system. The use of this model would have little direct application to more heterogeneous materials, but the goal is to help focus the study at higher levels of complexity [38].

1.5. Feedstock Supply

Production of corn (Zea mays) is the mainstay of the agricultural economy in the

Midwestern region of the United States. Corn was originally grown exclusively as a food or feed source, but recently corn has been increasingly used as an energy crop. It will take nearly 15% of the annual corn crop to feed the ethanol facilities that are in production or under construction. Over 98% of US ethanol production is currently derived from corn [44].

The United States annually produces about 250 million metric tons of corn grain per year.

The overall corn yield has an upward trend over its entire production history. The planted acreage has stabilized, but advances in production techniques and seed genetics have continued to fuel the increases in yield. 8

Ethanol production from corn currently only uses the starch fraction of the grain.

This leaves over one-half of the corn plant unused. Starch composes about 71% of the corn grain [11], but the com grain only composes about 45-48% of the com plant by weight [45].

The promise of com as an energy crop is obviously hindered by the practice of only using a relatively small fraction of the com plant. The reason that the rest of the plant goes largely unharvested is because it is composed of mostly lignocellulose. The ability to process the rest of the com plant would allow com to compare favorably to other dedicated energy crops that are being considered [10]. Production of ethanol from com is favored over that from other potential energy crops, because it is a relatively mature technology and because com can be produced, stored, and transported efficiently.

Com stover is the most abundant agricultural residue in the United States. The majority of com stover either decomposes in the field or is buried with tillage. The use of com stover as an energy source would greatly reduce COa emissions because the carbon in com stover is released as it degrades in the field. Replacing fossil sources with stover would lead to a net CO2 reduction [10]. Some of the stover must remain on the fields to help with erosion control and some is used as low quality animal feed. After these considerations there remains an estimated 80-100 million dry tons available each year [9]. There is a potential to use a portion of this feedstock in non-fermentative applications such as fiberboard and paper, but a majority of the material could go toward bioethanol production [1 ].

The remaining fraction of the com grain after ethanol production can have several names. The characteristics of these co-products depend on the milling method used to extract the starch. The wet-milling process uses a steeping step to help with the separation of the com kernel into its constituents. The kernel is separated in a series of steps into germ, 9 fiber (hulls), gluten, and starch. The starch is used for ethanol production. The germ is extracted for oil. The remaining parts are sold as feed products[10]. The dry-grind process is much simpler. The entire kernel is ground up and travels through the fermentation process.

The solids are separated after fermentation and are called distillers' dried grains and solubles.

The dry-grind process is the area where most of the growth in ethanol production is occurring[28]. This has led to several variations to this process that have some effect on the co-products. Examples of these modifications are the "quick germ"[46] and the "quick fiber"[47] processes developed at the University of Illinois. These techniques are focused on the separation of the germ and fiber so the oil can be recovered from these fractions. The oil is considered to be a higher value product that can be separately marketed. The overall effect on composition of the remaining co-products is limited to nutritive changes, and would not likely have a large effect on saccharification. The use of modified procedures abounds in the industry, but these techniques are held as proprietary. Regardless of the process a "rule of thumb" that holds is production of ethanol, feed products, and CO2 each account for 1/3 of the dry mass of the com kernel [10].

1.6. Motivation

There is a large supply of biomass available to be used to meet needs of energy and materials. There is a need to improve the understanding of lignocellulosic structure and improve the methods used to hydrolyze it. The ability to economically release the sugars tied up in lignocellulose will open the door to energy sustainability.

The second chapter will evaluate the use of mechanical separation of com stover to produce a stream with improved fiber characteristics and a stream that would allow for a facile saccharification treatment. The third chapter will address the potential use of solid 10 acid-catalyzed saccharification of corn fiber and distillers' grain. The fourth chapter will investigate the degradation behavior of sugars in reaction conditions. The fifth chapter discusses the actual use of pretreated distiller's grain in the catalyst system. The sixth chapter suggests future investigations into improving the acid-catalyzed saccharification procedure.

References

1. Wyman, C.E., Potential Synergies and Challenges in Refining Cellulosic Biomass to Fuels, Chemicals, and Power. Biotechnology Progress, 2003. 19: p. 254-262. 2. Claasen, P.A.M., et al., Utilization of Biomass for the Supply of Energy Carriers. Applied Microbiology and Biotechnology, 1999. 52: p. 741-755. 3. Zaldivar, J., J. Nielsen, and L. Olsson, Fuel Ethanol Production from Lignocellulose: A Challenge for Metabolic Engineering and Process Integration. Applied Microbiology and Biotechnology, 2001. 56: p. 17-34. 4. Abatzoglou, N. and E. Chornet, Acid Hydrolysis of Hemicellulose and Cellulose: Theory and Applications, in Polysaccharides: Structural Diversity and Functional Versatility, S. Dumitriu, Editor. 1998, Marcel Dekker: New York. p. 1007-1046. 5. Dadach, Z.-E. and S. Kaliaguine, Acid Hydrolysis of Cellulose: Part I. Experimental Kinetic Analysis. The Canadian Journal of Chemical Engineering, 1993. 71: p. 880- 891. 6. Jung, H.-J.G., et al., Impact of Accessibility and Chemical Composition on Cell Wall Polysaccharide Degradability of Maize and Lucerne Stems. Journal of the Science of Food and Agriculture, 2000. 80: p. 419-427. 7. Kim, T.H., et al., Pretreatment of Corn Stover by Aqueous Ammonia. Bioresource Technology, 2003. 90: p. 39-47. 8. Sun, Y., Hydrolysis of lignocellulosic Materials for Ethanol Production: A Review. Bioresource Technology, 2002. 83: p. 1-11. 9. Kadam, K.L. and J.D. McMillian, Availability of Corn Stover as a Sustainable Feedstock for Bioethanol Production. Bioresource Technology, 2003. 88: p. 17-25. 10. Brown, R.C., Biorenewable Resources: Engineering New Products from Agriculture. 1st ed. 2003, Ames, Iowa: Iowa State Press. 11. Gulati, M., et al., Assessment of Ethanol Production Options for Corn Products. Bioresource Technology, 1996. 58: p. 253-264. 12. S aha, B.C. and R.J. Bothast, Pretreatment and Enzymatic Saccharification of Corn Fiber. Applied Biochemistry and Biotechnology, 1999. 76: p. 65-77. 11

13. Um, B.-H., M.N. Karim, and L.L. Henk, Effect of Sulfuric and Phosphoric Acid Pretreatments on Enzymatic Hydrolysis of Corn Stover. Applied Biochemistry and Biotechnology, 2003. 105-108: p. 115-125. 14. Weil, J.R., et al., Removal of Fermentation Inhibitors Formed during Pretreatment of Biomass by Polymeric Adsorbents. Industrial and Engineering Chemistry Research, 2002. 41: p. 6132-6138. 15. Bergeron, P., C. Benham, and P. Werdene, Dilute Sulfuric Acid Hydrolysis of Biomass for Ethanol Production. Applied Biochemistry and Biotechnology, 1989. 20- 21: p. 119-134. 16. Saha, B.C., B.S. Dien, and R.J. Bothast, Fuel Ethanol Production from Corn Fiber. Applied Biochemistry and Biotechnology, 1998. 70-72: p. 115-125. 17. Esteghlalian, A., et al., Modeling and Optimization of the Dilute-Sulfuric-Acid Pretreatment of Corn Stover, Poplar and Switchgrass. Bioresource Technology, 1997. 59: p. 129-136. 18. Lee, Y.Y., P. Iyer, and R.W. Torget, Dilute-Acid Hydrolysis of Lignocellulosic Biomass. Advances in Biochemical Engineering / Biotechnology, 1999. 65: p. 93- 115. 19. Mosier, N., et al., Features of Promising Technologies for Pretreatment of Lignocellulosic Biomass. Bioresource Technology, 2005. 96: p. 673-686. 20. Sherrard, EC. and F.W. Kressman, Review of Processes in the United States Prior to World War II. Industrial and Engineering Chemistry Research, 1945. 37: p. 5-8. 21. Lin, Y. and S. Tanaka, Ethanol Fermentation from Biomass Resources: Current State and Prospects. Applied Microbiology and Biotechnology, 2006. 69: p. 627-642. 22. Hsu, T.-A., Pretreatment of Biomass, in Handbook of Bioethanol: Production and Utilization, C.E. Wyman, Editor. 1996, Taylor and Francis: Washington, p. 424. 23. Choi, C.H. and A.P. Mathews, Two-Step Acid Hydrolysis Process Kinetics in the Saccharification of Low-Grade Biomass: 1. Experimental Studies on the Formation and Degradation of Sugars. Bioresource Technology, 1996. 58: p. 101-106. 24. Allen, S.G., et al., A Comparison between Hot Liquid Water and Steam Fractionation of Corn Fiber. Industrial and Engineering Chemistry Research, 2001. 40: p. 2934- 2941. 25. Belkacemi, K., G. Turcotte, and P. Savoie, Aqueous/Steam-Fractionated Agricultural Residues as Substrates for Ethanol Production. Industrial and Engineering Chemistry Research, 2002. 41: p. 173-179. 26. Weil, J.R., et al., Pretreatment of Corn Fiber by Pressure Cooking in Water. Applied Biochemistry and Biotechnology, 1998. 73: p. 1-17. 27. Xiang, Q., Y.Y. Lee, and R.W. Torget, Kinetics of Glucose Decomposition during Dilute-Acid Hydrolysis of Lignocellulosic Biomass. Applied Biochemistry and Biotechnology, 2004.113-116: p. 1127-1138. 28. Tucker, M.P., et al., Conversion of Distiller's Grain into Fuel Alcohol and a Higher- Value Animal Feed by Dilute-Acid Pretreatment. Applied Biochemistry and Biotechnology, 2004. 113-116: p. 1139-1159. 29. Kim, S.B. and Y.Y. Lee, Diffusion of Sulfuric Acid Within Lignocellulosic Biomass Particles and its Impact on Dilute-Acid Pretreatment. Bioresource Technology, 2002. 83: p. 165-171. 12

30. Bustos, G., et al., Modeling of the Hydrolysis of Sugar Can Bagasse with Hydrochloric Acid. Applied Biochemistry and Biotechnology, 2003. 104: p. 51-68. 31. Gould, J.M., Studies on the Mechanism of Alkaline Peroxide Delignification of Agricultural Residues. Biotechnology and Bioengineering, 1985. 27: p. 225-231. 32. Iyer, P.V., et al., Ammonia Recycled Percolation Process of Pretreatment of Herbaceous Biomass. Applied Biochemistry and Biotechnology, 1996. 57-58: p. 121- 132. 33. Kaar, W.E. and M.T. Holzapple, Using Lime Pretreatment to Facilitate the Enzymic Hydrolysis of Corn Stover. Biomass and Bioenergy, 2000.18: p. 189-199. 34. Lopez, M.J., et al., Isolation of Microorganisms for Biological Detoxification of Lignocellulosic Hydrolysates. Applied Microbiology and Biotechnology, 2004. 64: p. 125-131. 35. Moiseev, Y.V., N.A. Khalturinskii, and G.E. Zaikov, The Mechanism of the Acid- Catalyzed Hydrolysis of Glucosides. Carbohydrate Research, 1976. 51: p. 23-37. 36. Smidsrod, O., A. Haug, and B. Larsen, The Influence ofpH on the Rate of Hydrolysis of Acidic Polysaccharides. Acta Chemica Scandinavica, 1966. 20: p. 1026-1034. 37. Lai, Y. Z., Chemical Degradation, in Wood and Cellulosic Chemistry, D.N.-S. Hon and N. Shiraishi, Editors. 2001, Marcel Dekker: New York. p. 443-512. 38. Mosier, N.S., et al., Characterization of Dicarboxylic Acids for Cellulose Hydrolysis. Biotechnology Progress, 2001. 17: p. 474-480. 39. Mosier, N.S., et al., Reaction Kinetics, Molecular Action, and Mechanisms of Cellulolytic Proteins. Advances in Biochemical Engineering / Biotechnology, 1999. 65: p. 23-40. 40. Conner, A.H., et al., Kinetic Modeling of the Saccharification of Prehydrolyzed Southern Red Oak, in Cellulose: Structure, Modification and Hydrolysis, R.A. Young and R.M. Rowell, Editors. 1986, Wiley: New York. p. 281-296. 41. Saeman, J.F., Kinetics of Wood Saccarification: Hydrolysis of Cellulose and Decomposition of Sugars in Dilute Acid at High Temperature. Industrial Engineering Chemistry Research, 1945. 37(1): p. 43-52. 42. Mosier, N.S., C.M. Ladisch, and M.R. Ladisch, Characterization of Acid Catalytic Domaines for Cellulose Hydrolysis and Glucose Degradation. Biotechnology and Bioengineering, 2002. 79(6): p. 610-618. 43. Abatzoglou, N., et al., Phenomenological Kinetics of Complex Systems: The Development of a Generalized Severity Parameter and its Application to Lignocellulsic Fractionation. Chemical Engineering Science, 1992. 47: p. 1109. 44. U.S. Fuel Ethanol Production Capacity, [world wide web] 2005 Feb 2005 [cited 2005 2/28]; Available from: www.ethanolrfa.org/eth prod fac.html. 45. Hedin, P. A., W.P. Williams, and P.M. Buckley, Caloric Analyses of the Distribution of Energy in Corn Plants Zea mays L. Journal of Agricultural and Food Chemistry, 1998. 46: p. 4754-4758. 46. Singh, V. and S.R. Eckoff, Effects of Soak Time, Soak Temperature, and Lactic Acid on the Germ Recovery Parameter. Cereal Chemistry, 1996. 73: p. 716-720. 47. Wahjudi, J., et al., Quick Fiber Process: Effect of Mash Temperature, Dry Solids, and Residual Germ on Fiber Yield and Purity. Cereal Chemistry, 2000. 77: p. 640-644. 13

CHAPTER 2

Hydrolysis Characteristics of the Tissue Fractions Resulting from Mechanical

Separation of Corn Stover

A paper published in Applied Biochemistry and Biotechnology1

2.1. Introduction

Corn stover continues to receive a great deal of attention as a potential source of valuable co- products from corn production. Because it is high volume, low cost, and renewable, stover has desirable attributes as a feedstock for energy, chemicals, and materials. Although most com stover is currently left in the field, collection strategies with improved economics are being developed and demonstrated that hold promise for delivering stover economically to processing sites [1], Therefore, the limiting factor in the utilization of com stover resides primarily with conversion technologies. Despite nearly a century of research into the utilization of com stover in paper manufacturing, building materials, and ethanol and energy production, no economically viable utilization strategy has yet been demonstrated.

The most active area of research in com stover utilization is its hydrolysis into monomeric sugars, which can be subsequently converted to a number of possible chemicals including ethanol. Quite a number of different approaches have been explored relative to the production of monomeric sugars from stover including concentrated-acid hydrolysis [2], dilute-acid hydrolysis [3-5], and enzymatic hydrolysis [5-11]. In the case of enzymatic hydrolysis, a pretreatment step is required such as steam pretreatment [6, 12, 13], ammonia

1 Reprinted with permission of Applied Biochemistry and Biotechnology, 2005.125: p. 27-39. fiber explosion [14-16], lime pretreatment [7,17], alkaline pretreatment [8, 18, 19], dilute- acid pretreatment [9, 10, 20], and alkaline peroxide pretreatment [11]. Many of these methods as well as combinations of the methods show promise; however the economics have not been sufficiently compelling to lead to industrial use.

The challenge of hydrolyzing com stover can be viewed from both the molecular and cellular level. At the molecular level, stover is a lignocellulosic biomass that is composed of cellulose interwoven with hemicellulose and coated with lignin. Lignin, which is a complex phenolic polymer, creates a barrier to the hydrolysis of the hemicellulose and cellulose.

Cellulose, which forms relatively stiff fibrils, is a linear polymer of glucose units, and hemicellulose is a branched polymer of a variety of pentoses and hexoses that surrounds the cellulose fibrils. Due to its structure, hemicellulose is more susceptible to hydrolysis than cellulose.

Com stover also has a heterogeneous structure at the cellular level. Mature com stover can be classified as consisting of two main types of tissue: fiber and pith. The pith fraction, which comprises 40-50% of the stalk by weight and about 75% by volume [21], consists of the remains of the parenchyma and collenchyma cells from the growing plant.

This tissue is a relatively soft, spongy amorphous material. Parenchyma cells are thin- walled metabolically active cells whose function in stems is mostly storage. Collenchyma cells are thick walled; however, they are not lignified, which allows them to stretch as the stalk elongates. These cells provide structural support and are typically arranged in layers or bundles. The fiber fraction is the result of the sclerenchyma cells from the growing plant.

These cells have very thick walls, with generally over one-half of the cell volume within the 15 wall [21], and are morphologically long and narrow. Sclerenchyma cells are typically dead at maturity and have the function of providing mechanical support to the plant.

The hydrolysis methods discussed above are largely driven by molecular level considerations. For example, a goal of many pretreatment procedures is to solubilize the lignin and hydrolyze the hemicellulose leaving primarily cellulose for the subsequent enzymatic hydrolysis step. The heterogeneity of the distinct tissue types lends them to a potential alternative processing approach. The differences in the functions of the types of tissue create tissue fractions that can be mechanically separated. Grinding corn stover yields morphologically distinct fractions, with the fiber portion having a high-aspect-ratio rodlike shape and the pith portion having irregular low-aspect-ratio shapes. A simple mechanical separation can then exploit the morphological differences between the two types of tissue.

Once the fiber and pith portions are separated, the fiber portion could have potential application in traditional fiber utilization whereas the pith fraction could be hydrolyzed to its constituent monomeric sugars. This mechanical separation according to type of tissue could be an attractive processing option if the pith fraction is hydrolyzed more readily than the fiber portion and, therefore, the overall stover.

Presented here are results concerning the relative hydrolysis efficacy for the two tissue types to determine whether mechanical separation can be used to improve the subsequent processing of corn stover. The hydrolysis experiments were performed under both acidic and alkaline conditions to evaluate the relative hydrolysis behavior of the tissue fractions that result from mechanical separation. Hemicellulose is readily hydrolyzed under acidic conditions, whereas alkaline hydrolysis more readily targets the lignin. Saddler et al.

[12] compared the results of ammonia recycled percolation with those of dilute-acid 16 hydrolysis. They found that both treatments solublized about 45% of the stover. Whereas the ammonia-recycled percolation process removed about 85% of the lignin and 60% of the hemicellulose, the dilute-acid treatment removed only 33% of the lignin and nearly 95% of the hemicellulose. Using both types of hydrolysis approaches would provide additional information about the structural and chemical differences of the two tissue fractions.

The experiments were designed to determine the hydrolysis characteristics of these tissue fractions rather than to optimize the hydrolysis reaction. The results from the hydrolysis reactions with the tissue fractions will be discussed within the context of evaluating the usefulness of physical separation prior to hydrolysis.

2.2 Materials and Methods

Materials

Sulfuric acid and sodium hydroxide used were from Fisher Scientific (Fair Lawn,

NJ). The corn stover fractions, which were harvested from the 2001 growth year near

Harlan, IA, were provided by the former BioMass Agri-Products of Harlan. The stover was washed and chopped at its facility. The mechanical approach used to separate the tissue fractions in the current work was similar to that used in pulping processes. The chopped corn stover was slurried with water and the slurry was then fed through a Tornado underwater pulper for size reduction. To further separate the fibers, the pulped material was sent through a rotor-stator refiner. The slurry was then dewatered with a screw press and dried in a tubular flash drier. The dried material was separated using air classification with selective removal of the lighter pith fraction. Shown in Figure 1 is the dried material prior to air classification. Hydrolysis Reactor

The hydrolysis experiments were conducted in a Parr 300-mL stainless-steel batch reactor, which was equipped with a glass liner and an impeller-type mixer. The reactor temperature was controlled using a programmable logic controller (PLC) connected to both a heating jacket and an internal cooling loop. In the hydrolysis experiments, pith and fiber materials were individually slurried in a 1:30 weight ratio (stover: liquid).

Analytical Methods

The hydrolysis products were analyzed with an HPLC system (Agilent 1050) using a

Hewlett-Packard refractive index detector. The mobile phase was ultrapure water with a pump rate of 0.4 mL/min, and the column was a Supelcogel Pb (Supelco). The column temperature was set at 85°C. Data were collected and analyzed with Chemstation software.

The Branauer-Emmett-Teller (BET) surface area of the materials was determined with nitrogen adsorption at 77 K with an ASAP 2000 Surface Area and Porosity Analyzer

(Micromeritics). Scanning electron microscopy (SEM) micrographs were obtained using a

JEOL JSM-840 microscope operating at a vacuum of 10"6 torr. Samples were sputter coated with gold for analysis. The micrographs used were taken at x200 magnification.

Acid Hydrolysis

From trials at several dilute sulfuric acid concentrations, it was determined that 0.8 wt% was a reasonable condition for the dilute-acid hydrolysis experiments. The hydrolysis experiments were conducted over a temperature range of 140 to 180°C. Below 140°C, very little hydrolysis occurred for either material, and above 180°C, excessive sugar degradation made accurate mass balances impossible. These conditions were consistent with others reported in the literature [9]. For the acid-hydrolysis experiments the temperature was 18 ramped from 140 to 180°C. The reactor temperature was increased in 10°C increments with the temperature change requiring about 5 min. The reactor was then allowed to maintain this temperature for 5 min before the sample was taken. This procedure was used throughout the specified temperature range.

Base Hydrolysis

Base-catalyzed hydrolysis is most often used as a pretreatment to remove lignin and swell the cellulosic strands. Alkaline hydrolysis was used in this study to determine to what extent any lignification differences affected the response of the tissue fractions to hydrolysis.

A 1 wt% sodium hydroxide solution was used for the experiments. Although the alkaline- hydrolysis experiments were performed over a range of temperatures, the results did not exhibit strong temperature dependence. Therefore, the base-hydrolysis experiments included here are for varying hydrolysis times at 140°C.

2.3 Results and Discussion

Shown in Fig. 2A, B are representative SEM micrographs for the separated pith and fiber fractions. As can be seen, the pith fraction consisted of irregularly sized and shaped particles, whereas the primary particles in the fiber fraction were of more uniform dimension.

The fiber particles exist individually or with several bundled together. Review of several

SEM micrographs as well as visual inspection revealed that mechanical separation resulted in a fiber fraction with little pith cell content. By contrast, the pith fraction contained about

20% of particles that had fiber-type morphology and could be described as short fibers.

Morphology and surface area of the corn stover fractions are important because hydrolysis is affected by mass transfer considerations [22, 23]. SEM micrographs were used to estimate surface-to-volume ratios for the cell fractions. Although the fiber cells were bundled together into larger particles as seen in Fig. 2B, the fiber cell bundles appeared to have adequate porosity such that the surface-to-volume ratio of fiber particles was taken to be that of the individual fibers. With this assumption, the surface-to-volume ratio of the fiber fraction fell within the range of 28:1 to 32:1 m2/m3. The pith fraction was significantly more heterogeneous than the fiber fraction and had surface-to-volume ratios from 5:1 to 22:1 m2/m3. The surface areas of the two fractions were determined using BET analysis with adsorption. The surface area of the pith fraction was 2.85 m2/g and that of the fiber fraction was 2.93 m2/g, indicating that the fractions would be equally accessible to acid hydrolysis.

It has been demonstrated under similar acid-hydrolysis conditions to those used in the current study that degradation of xylose and glucose will occur. To insure closure of the mass balances for the hydrolysis experiments, the degradation rates of the free sugars were determined. Therefore, experiments were conducted with xylose and glucose using the same protocol as that used to hydrolyze the corn stover fractions to quantify their degradation rates under the hydrolysis conditions. Figure 3 presents the results from the degradation experiments. The quantitative values for glucose degradation agreed closely with published results [24]. Comparable quantitative degradation rate data for xylose were not available, but consistent with prior work, the xylose degraded more rapidly and at a lower temperature than the glucose.

Shown in Fig. 4 are results from acid hydrolysis of the pith cell fraction in which the mass fraction of pith cells hydrolyzed is given as a function of temperature during the experimental run. The lower curve on the figure corresponds to the apparent mass fraction of the pith cells hydrolyzed as determined by the amount of sugars measured by HPLC. As 20 demonstrated in Fig. 3, sugar degradation was simultaneously occurring with generation of sugars by hydrolysis. Therefore, the hydrolysis curve was corrected for the amount of sugar degraded based on the pure sugar controls run under similar reaction conditions. For the correction, the degradation rates for the hexoses and pentoses were assumed to be characterized by those of glucose and xylose, respectively, and to be zero order in sugar concentration. By invoking these assumptions, the HPLC results were corrected for the sugar degradation rate to yield the actual hydrolysis rate. The top curve in Fig. 4 corresponds to the corrected hydrolysis rate.

No solubilized oligosaccharides were observed from HPLC analysis in the pith cell hydrolyzate, which is consistent with previous results that have demonstrated that the concentration of oligomers are typically very low under comparable hydrolysis conditions

[25]. For further validation, a secondary hydrolysis was performed using the method of Kim et al. [26] to check for the presence of oligomers. No change in sugar concentrations were detected in the solution following the secondary hydrolysis, so the oligomer content of the hydrolyzate had to be sufficiently low as to be under the detection limit of the HPLC analysis.

Table 1 gives a representative mass balance taken at the 180°C sample point.

Included in Table 1 are the initial pith cell mass as well as the solid unhydrolyzed mass remaining at the conclusion of the experiment, as well as the measured quantity of free sugars and the amount of sugars that were lost to degradation as determined from the glucose and xylose degradation experiments. As can be seen, the closure of the mass balance is quite good when only the solids and free and degraded sugars are considered, indicating that the sugar degradation estimate used to find the actual hydrolysis rate was reasonable. The closure within 2-3% also indicated that the lignin, which was present in corn stover at about

20 wt% [27], must have been present in the solids fraction at the conclusion of the experiment. Therefore, any lignin that was solubilized during hydrolysis must primarily have been re-precipitated on the solids. This lignin precipitation behavior has been previously demonstrated in the hydrothermal treatment of com stover [28].

A comparison of the acid hydrolysis of the pith and fiber fractions is shown in Fig. 5, with the mass hydrolyzed given as a function of reaction temperature. Both curves were determined by HPLC analysis of dissolved sugars corrected for sugar degradation, as already described. A mass balance for the fiber cell hydrolysis experiment is given in Table 1 corresponding to the 180°C sample point. As with the pith cell hydrolysis, good mass balance closure was achieved for the fiber cell hydrolysis. From Fig. 5 it is clear that the acid-hydrolysis behaviors of the two corn stover cell fractions were indistinguishable.

The commonality of acid hydrolysis results for the two corn stover fractions can also be seen when the individual sugar measurements are compared. Shown in Fig. 6 are the hexose and pentose mass fractions measured in solution over the course of the hydrolysis experiments for the pith and fiber fractions. The mass fraction represents the amount of free sugar measured relative to the initial mass of pith or fiber cells. The curves shown in Fig. 6 were not corrected for sugar degradation as can be seen most clearly from the pentose curves.

As expected, the pentoses were the first sugars to be detected because the hemicellulose fraction of the material was easier to hydrolyze. Higher temperatures were required to hydrolyze the cellulose fraction to glucose, which was consistent with cellulose being more recalcitrant towards hydrolysis.

Because the goal of the current study was to compare the hydrolysis characteristics of the pith and fiber fractions, hydrolysis experiments were also performed under alkaline conditions. It has been demonstrated that alkaline pretreatment in the hydrolysis of lignocellulose, given sufficiently high base concentration and reaction temperature, can solubilize most of the lignin and hemicellulose [12]. In contrast, the glycosidic bonds in cellulose are known to be stable under alkaline conditions [12], so the cellulose will remain intact even after most of the lignin and hemicellulose are solubilized [14]. For the growing corn plant, the fiber fraction of corn stover is known to be slightly more lignified [26].

Therefore, the alkaline hydrolysis experiments should provide some additional information on the hydrolysis characteristics of the two cell fractions.

Figure 7 presents the results for the alkaline hydrolysis in which the loss of mass of the solids was determined as a function of reaction time. Unlike acid-hydrolysis experiments, mass balance closure was immpossible to achieve under the alkaline hydrolysis, because the hydrolysis resulted in few monomers that could be detected by the HPLC method. Solubilized oligomers were detected but were not separated into molecular species, making quantification difficult. Therefore, the reported mass loss was determined exclusively by direct measurement of the mass of the solids portion left after hydrolysis. As can be seen from Fig. 7, the fiber portion had a more rapid initial mass loss under the alkaline conditions, but by 90 min both fractions had similar amounts of solubilized mass. Because the loss of mass in these reactions could be attributed to solubilization of lignin and hemicellulose, the overall mass losses for the cell fractions would be consistent with the two tissue types having similar mass composition of the two polymeric types. The slight difference in time dependence of the mass suggested a difference in the architecture and/or accessibility of the type of tissue to the alkaline catalyst. The current results suggested that despite the difference in cell function between the pith and fiber tissue fractions, the tissue types when taken from the mature corn stover have similar hydrolysis characteristics. These results can be compared against reports that have attempted to quantify the molecular content of corn stover. Morrison et al. [29] conducted studies on the cell-wall composition of maize that compares the pith and rind tissues. The tissue that is referred to as rind in their study would be similar to the fiber tissue used in the current study. In that work, stalks of varying maturity were separated into outer rind and central pith portions. Shown in the first two rows of Table 2 are neutral sugar compositions for the two tissue fractions taken from the most mature stover used in their study. The neutral sugar compositions as determined by HPLC analysis of the acid-hydrolyzed starch- free residue were found to be very similar. The glucose contents of both fractions, which correlates with the cellulose content, were nearly identical. Although the distribution of the four major neutral sugars that comprise the hemicellulose fraction were distributed differently, the total amount of sugars was the same.

In another study, Jones et al. [30] examined neutral sugar contents from mature com stover that was separated into three segments: pith, fibrovascular bundles, and fiber from the rind. The pith tissue in that study was similar to the pith tissue in the current study and the combination of the fibrovascular bundles and fiber from the rind correlates with the fiber tissue. Results from the study are given in the lower three rows of Table 2. In contrast to the other study, their results show a higher percentage of glucose in each tissue fraction. Their work included an attempt to quantify the hemicellulose and cellulose portions of the tissues by using alkaline extraction of hemicellulose from delignified tissues. Although the authors acknowledged that their method overestimated the glucose content in the hemicellulose, and therefore underestimated the cellulose content, the results provided some insight into the

composition of the tissues examined. They found that the fibrovascular bundles had the

highest cellulose content (32.3 wt%), followed by pith (27.8 wt%), with the rind being

lowest (20.3 wt%).

Although these two studies gave slightly different results for the composition of

neutral sugars in the pith and fiber tissue fractions as defined in the current study. The

overall conclusion is that both fractions appear to have similar distributions of neutral sugars, suggesting that the cellulose and hemicellulose composition in each fraction was comparable.

The current results were consistent with this conclusion because the overall mass hydrolyzed for the pith and fiber fractions was similar.

Because the amount of cellulose and hemicellulose present in the pith and fiber

fractions were comparable, the other factor that could have affected the hydrolysis

characteristics of the fractions was their respective lignification. Lignification of cell walls is

widely regarded as the primary factor limiting acidic and enzymatic hydrolysis. This

conclusion has been drawn from the correlation between degradability and lignin content.

The degree of lignification between the pith and fiber tissue does differ slightly [29], but for

corn stover the lignin content in general increases as the plant matures [31]. Degradability studies based on mature tissue have suggested that there is another factor that affects the degradability of the tissues as they age, because lignin content was not predictive of degradability with the tissue maturity [32]. Jung et al. [33] performed experiments in which

corn stover was ball milled to assess whether substrate accessibility was as a factor in

degradation rates. They were unable however to explain differences in polysaccharide

degradation that remained even when accessibility limitations were removed, and they 25 concluded that the most important factor limiting cell wall degradation was currently not being measured [33].

2.4 Conclusions

Hydrolysis of corn stover is a promising source of monomelic sugars if an economical hydrolysis process could be developed. Mechanical separation of the pith and fiber tissue fractions provides an inexpensive approach to separation of corn stover by cellular function. However, hydrolysis experiments performed on the separated tissue fractions showed no significant difference in hydrolysis characteristics despite the significant difference in gross cell structure of the two tissue fractions. The comparable hydrolysis characteristics for the tissue types was likely related to the fact that mature corn stover was used in the study, because the lignin content in the stover increases with maturation. Because mature com stover is the best target for a hydrolysis feedstock, the use of mechanical separation prior to hydrolysis does not appear to present an attractive alternative to the current hydrolysis work that emphasizes hydrolysis processing of the entire com stover.

References

1. Glassner, D.A., J.R. Hettenhaus, and T.M. Schechinger. Corn Stover Collection Project, in Bioenergy 98. 1998. Madison. 2. Goldstein, I.S., et al., The Hydrolysis of Cellulose with Superconcentrated Hydrochloric Acid. Biotechnology and Bioengineering, 1983.13: p. 17-25. 3. McParland, J. J., H.E. Grethlein, and A.O. Converse, Kinetics of Acid Hydrolysis of Corn Stover. Solar Energy, 1982. 28: p. 55-63. 4. Bhandari, N., D.G. Macdonald, and N.N. Bakhshi, Kinetic Studies of Corn Stover Saccharification Using Sulphuric Acid. Biotechnology and Bioengineering, 1984. 26: p. 320-327. 26

5. Chen, R.F., Z.-W. Wu, and Y.Y. Lee, Shrinking-bed Model for Percolation Process Applied to Dilute-Acid Pretreatment Hydrolysis of Cellulosic Biomass. Applied Biochemistry and Biotechnology, 1998. 70-72: p. 37-50. 6. Belkacemi, K., G. Turcotte, and P. Savoie, Aqueous/Steam-Fractionated Agricultural Residues as Substrates for Ethanol Production. Industrial and Engineering Chemistry Research, 2002. 41: p. 173-179. 7. Kaar, W.E. and M.T. Holzapple, Using Lime Pretreatment to Facilitate the Enzymic Hydrolysis of Corn Stover. Biomass and Bioenergy, 2000. 18: p. 189-199. 8. Varga, E., et al., Pretreatment of Corn Stover Using Wet Oxidation to Enhance Enzymatic Digestibility. Applied Biochemistry and Biotechnology, 2003. 104: p. 37- 50. 9. Esteghlalian, A., et al., Modeling and Optimization of the Dilute-Sulfuric-Acid Pretreatment of Corn Stover, Poplar and Switchgrass. Bioresource Technology, 1997. 59: p. 129-136. 10. Um, B.-H., M.N. Karim, and L.L. Henk, Effect of Sulfuric and Phosphoric Acid Pretreatments on Enzymatic Hydrolysis of Corn Stover. Applied Biochemistry and Biotechnology, 2003.105-108: p. 115-125. 11. Gould, J.M., Studies on the Mechanism of Alkaline Peroxide Delignification of Agricultural Residues. Biotechnology and Bioengineering, 1985. 27: p. 225-231. 12. Saddler, J.N., L.P. Ramos, and C. Breuil, Steam Pretreatment of Lignocellulosic Residues, in Bioconversion of Forest and Agricultural Plant Wastes, J.N. Saddler, Editor. 1993, C.A.B. International: Wallingford. 13. Nevell, T.P. and S.H. Zeronian, Cellulose Chemistry and its Applications. 1985, New York: John Wiley & Sons. 14. Iyer, P.V., et al., Ammonia Recycled Percolation Process of Pretreatment of Herbaceous Biomass. Applied Biochemistry and Biotechnology, 1996. 57-58: p. 121- 132. 15. Holtzapple, M., et al., The Ammonia Freeze Explosion Process-A Practical Lignocellulose Pretreatment. Applied Biochemistry and Biotechnology, 1991. 28-29: p. 59-74. 16. Moniruzzaman, M., et al., Enzymatic Hydrolysis of High-Moisture Corn Fiber Pretreated by AREX and Recovery and Recycling of the Enzyme Complex. Applied Biochemistry and Biotechnology, 1997. 67: p. 113-126. 17. Chang, V.S., M. Nagwani, and M. Holtzapple, Lime Pretreatment of Crop Residues Bagasse and Wheat Straw. Applied Biochemistry and Biotechnology, 1998. 74: p. 135-159. 18. Kim, T.H., et al., Pretreatment of Corn Stover by Aqueous Ammonia. Bioresource Technology, 2003. 90: p. 39-47. 19. Hsu, T.-A., Pretreatment of Biomass, in Handbook of Bioethanol: Production and Utilization, C.E. Wyman, Editor. 1996, Taylor and Francis: Washington, p. 424. 20. Grohmann, K., R.W. Torget, and M. Himmel, Dilute Acid Pretreatment of Biomass at High Solids Concentrations. Biotechnology and Bioengineering, 1985. 15: p. 59-80. 21. Brett, C. and K. Waldron, eds. Physiology and Biochemistry of Plant Cell Walls. 1990, Unwin Hyman: London. 27

22. Mosier, N.S., et al., Reaction Kinetics, Molecular Action, and Mechanisms of Cellulolytic Proteins. Advances in Biochemical Engineering / Biotechnology, 1999. 65: p. 23-40. 23. Kim, S.B. and Y.Y. Lee, Diffusion of Sulfuric Acid Within Lignocellulosic Biomass Particles and its Impact on Dilute-Acid Pretreatment. Bioresource Technology, 2002. 83: p. 165-171. 24. Bienkowski, PR., et al., Correlation of Glucose Degradation at 90 to 190°C in 0.4 to 20% Acid. Chemical Engineering Communications, 1987. 51: p. 179-192. 25. Lee, Y.Y., P. Iyer, and R.W. Torget, Dilute-Acid Hydrolysis of Lignocellulosic Biomass. Advances in Biochemical Engineering / Biotechnology, 1999. 65: p. 93- 115. 26. Kim, J.S., Y.Y. Lee, and R.W. Torget, Cellulose Hydrolysis Under extremely Low Sulfuric Acid and High-Temperature Conditions. Applied Biochemistry and Biotechnology, 2001. 91-93: p. 331-340. 27. Hames, B.R., et al., Rapid Biomass Analysis. Applied Biochemistry and Biotechnology, 2003.105-108: p. 5-16. 28. Rubio, M., et al., Fractionation ofLignocellulosics. Solubilization of Corn Stalk Hemicelluloses by Autohydrolysis in Aqueous Medium. Biomass and Bioenergy, 1998.15: p. 483-491. 29. Morrison, T.A., et al., Cell-Wall Composition of Maize Internodes of Varying Maturity. Crop Science, 1998. 38: p. 455-460. 30. Jones, R.W., et al., Neutral Sugars of Hemicellulose Fractions of Pith from Stalks of Selected Plants. Cereal Chemistry, 1979. 56: p. 441-442. 31. Hedin, P. A., W.P. Williams, and P.M. Buckley, Caloric Analyses of the Distribution of Energy in Corn Plants Zea mays L. Journal of Agricultural and Food Chemistry, 1998. 46: p. 4754-4758. 32. Jung, H.-J.G., D R. Buxton, and R.D. Hatfield, Forage Cell Wall Structure and Digestibility, ed. J. Ralph. 1993, Madison: American Society of Agronomy. 33. Jung, H.-J.G., et al., Impact of Accessibility and Chemical Composition on Cell Wall Polysaccharide Degradability of Maize and Lucerne Stems. Journal of the Science of Food and Agriculture, 2000. 80: p. 419-427. 28

Table 1 Representative mass balances for the pith and fiber fractions under 0.8 wt% acid hydrolysis at 180°C.

PITH FIBER Initial weight (g) 5.0 5.3 Mass after hydrolysis (g) 2.6 2.5 Mass of detected sugars (g) 1.04 1.20 Mass of degraded sugars (g) 1.27 1.46 Total difference (g) 0.09 0.14 29

Table 2 Sugar distribution reported previously for the cell-wall matrix of pith and rind tissues.

Glucose Xylose Arabinose Galactose Mannose Pith a (%) 60.6 25.4 4.9 4.4 4.7 Rind a (%) 60.8 31.8 2.9 2.5 2.0 Pith b (%) 65.5 29.5 4.6 Trace Trace Fibrovascular 67.5 29.5 3.0 Trace Trace Bundles b(%) Fiber-Rind b(%) 69.0 26.8 4.6 Trace Trace a Morrison, T. A., H. G. Jung, et a . (1998). Crop Sci. 38,455-460. b Jones, R. W., L. H. Krull, et al. ( 1979). Cereal Chem. 56, 441^442. Fig. 1 Ground corn stover prior to air classification. Fig. 2a) SEM Micrograph (200X) of mechanically separated pith fraction Fig. 2b) SEM Micrograph (200X) of mechanically separated fiber fraction 33

70 i

60

50 -à- xylose -•-glucose w 40 0) o —i %30 nj S 20

10

0 1 » 130 140 150 160 Temperature (°C)

Fig. 3 Mass of glucose and xylose from degradation. 34

0.5 0.45

0.35 !" £<>.25 (/> : o.2 -•-sugars wI 55 0.15 degraded fraction 0.1 •sugars 0.05

0 130 140 150 160 170 180 Temperature (°C)

Fig. 4 Sugar released during acid hydrolysis with correction for sugar loss due to degradation. 35

•pith •fiber

130 140 150 160 170 180 190 Temperature (°C) Fig. 5 Acid hydrolysis results for the pith and fiber fractions. 36

0.35 -4-hexoses - pith -t-pentoses -pith 0.3 -à-hexoses - fiber pentoses -fiber .0.25 -F

S 0.2

*0.15 t/> CO s 0.1

0.05

130 140 150 160 170 180 190 Temperature (°C) Fig. 6 Distribution of hexoses and pentoses released by acid hydrolysis of the tissue fractions. 37

30 60 90 Time (min)

Fig. 7 Mass loss of the pith and fiber fractions under alkaline conditions. 38

CHAPTER 3

Disaccharide Hydrolysis Using Organic-Inorganic Hybrid

Mesoporous Silica Catalysts

3.1. Introduction

The production of ethanol in the United States has increased 10-fold over the past 30

years with the production capacity expected to reach 5 billion gallons in 2005 [1]. This

ethanol production is primarily from the starch fraction of corn, so with the increase in

production has come a concomitant large increase in the co-product streams. The most significant co-product streams are com fiber from the wet-milling process and distillers' dry grains from the dry milling process. These streams are currently sold as animal feed and the revenue from them is important to the economics of ethanol production. With the large increase in ethanol production, however, there is a significant danger of market saturation for the co-products [2, 3]. A possible alternative would be to hydrolyze the co-product streams to produce fermentable sugars thereby providing three major benefits: a boost in the yield of ethanol per bushel, a substantial decrease in the volume of the co-product streams, and an increase in the value of the remaining co-product due to the increase protein content and reduced fiber content [2, 3],

Several previous studies on the conversion of distillers' dry grains and com fiber to ethanol have been reported [4-7], with most of the proposed processes utilizing multi-step approaches. Generally, a key step is the hydrolytic release of fermentable sugars from the co-products. The hydrolysis has commonly been performed on oligosaccharides released from pretreatment of the fiber streams, with the oligosaccharides possibly in a solubilized 39 form. This hydrolysis step has typically been done with enzymes or a homogeneous acid catalyst.

Tucker et al. conducted a study on distillers' dry grain and found that they could achieve solubilization of 77% of the total carbohydrates in the distillers' grain by using sulfuric acid. Saha and Bothast performed studies on corn fiber using hot water, acid, and alkaline pretreatments [5], They found that few sugar monomers were released by the pretreatments, but the pretreatment assisted in subsequent enzymatic treatments. Their study gave no indication of the extent to which solubilized oligomers were released by pretreatment of the material. More recently, the Ladisch group has performed tests on com fiber and found that using a simple hydrothermal treatment with pressure cooking of the material at 220 °C solubilized up to 70% of the original mass with this solubilized fraction consisting of mostly oligosaccharides [4],

Enzymes were used in each of these previous studies to subsequently hydrolyze the oligosaccharides. While enzymes are very selective and produce no degradation products, they may be too expensive for the production of fuel ethanol [7, 8]. Also, the enzymes typically required long reaction times in these studies using hydrolysis times of two to four days [2, 5,6], Due to the variety of hexoses and pentoses present in the oligosaccharides, the enzymatic hydrolysis reaction required a cocktail of enzymes to be effective. Finally, enzymes are typically difficult to separate and reuse [9],

Alternatively, homogeneous acids offer the possibility of higher hydrolysis reaction rates from higher temperature operation, but this will also lead to degradation products. This degradation reaction not only reduces yield, but it forms compounds that are toxic to the yeasts used to ferment the released sugars. The homogenous acids must be neutralized 40 following reaction, and this can form salts that adversely affect feed value to livestock [2], requiring expensive separation and disposal steps [10, 11]. Solid acid catalysts, however, offer the potential to replace homogeneous acids yielding fast reaction rates, but with simple catalyst separation.

In designing a solid acid catalyst for hydrolysis, a material with a large number of accessible acid sites that have sufficient acidity to catalyze the hydrolysis is desirable.

Sulfonic acid resins potentially meet these criteria as long as the maximum temperature to be used does not exceed about 130 °C. However, another possibility with higher temperature operability as well as improved accessibility to the acidic groups is organic-inorganic hybrid mesoporous materials as well as improved accessibility to the acidic groups. These types of materials with organosulfonic acid groups have been previously used for the conversion of biorenewable molecules [12-15]. Mbaraka et al. [14] compared the catalytic performance of organosulfonic acid-functionalized mesoporous silicas to a sulfonated resin in the esterification of fatty acids and found that the improved internal transport properties of the mesoporous materials gave better overall reaction rates. Organosulfonic acid-functionalized mesoporous silica was used by Dhepe et al. [12] to hydrolyze the predominantly a-(l ,4) glycosidic bonds involved in the conversion of starch to glucose. They report better starch hydrolytic activity with an ethylsulfonic acid-functionalized mesoporous silica than with a sulfonated resin.

The oligosaccharides released in the hydrothermal treatment of corn fiber or distillers' dry grains would have primarily P-(1,4) glycosidic bonds rather than a-(l,4) bonds.

A simple model system for these soluble species would be cellobiose, which is a disaccharide with (3-(l ,4) linked glucose units. Cellobiose is the subunit of cellulose and has been used as 41 the model system in several hydrolysis studies [16-20], The current work investigates the use of organo-functionalized mesoporous silicas as catalysts for cellobiose hydrolysis. In addition to examining the hydrolysis reactivity of the organic-inorganic hybrid materials, the selectivity to glucose relative to its further degradation was determined.

3.2. Experimental

Organic-inorganic hybrid mesoporous silicas with three different types of organic acid groups were synthesized for the study. Shown in Figure 1 are schematics of the propylsulfonic, arenesulfonic, and butylcarboxylic acid moieties used in the catalytic materials. The precursor silane species employed in the synthesis of the catalytic materials, which were tetraethoxysilane (TEOS) (98%, Aldrich), (3-mercaptopropyl)trimethoxysilane

(MPTMS) (85%, Acros), 2-(4-chlorosulfonylphenyl)ethyltrimethoxysilane (CSPTMS) solution in dichloromethane (50%, Gelest), and 3-cyanopropyltriethoxysilane (CPTES)

(98%, Aldrich), were used as received. The surfactant used in the supramolecular assembly of the organic-inorganic hybrid materials was Pluronic 123 (BASF), a tri-block copolymer of polyethylene oxide-polypropylene oxide-polyethylene oxide with the molecular structure

PEO20-PPO70-PEO20 (MW = 5800).

3.2.1 Propylsulfonic acid-functionalized mesoporous silica

The method used to synthesize the propylsulfonic acid-functionalized silica was based on the procedure reported by Stucky and co-workers [21], in which the TEOS and

MPTMS were co-condensed in the presence of a tri-block copolymer and hydrogen peroxide under acidic conditions. In a typical synthesis, 8 g of Pluronic 123 was dissolved in a mixture of 260 ml of de-ionized water and 40 ml of concentrated hydrochloric acid. The mixture was heated to 50 °C before the addition of 14.9 ml of TEOS. A prehydrolysis time of 3 h was allowed before the MPTMS and 30% hydrogen peroxide were added. The mixture was then stirred for 20 h at 50 °C and subsequently aged in a 100 °C oven for 24 h without stirring. The solid product was recovered by filtration and air dried followed by ethanol reflux for 24 hours to remove the template. The material was made with two different concentrations of the acidic groups corresponding tot MPTMS/TEOS molar ratios of 0.075 and 0.15.

The method used to synthesize the HMS materials was based on a method developed by Bosseart and co-workers [22] using n-dodecylamine as the templating agent while co- condensing MPTMS and TEOS. The amine was dissolved in 40% ethanol at room temperature before adding the MPTMS and TEOS. The solution was stirred at room temperature for 24 h. The solids were then filtered and air dried. The template was removed by refluxing in ethanol for 24 h. Thiol oxidation was done with hydrogen peroxide (2 g/g solid) stirring in a methanol water mixture for 24 h at room temperature. The solids were then filtered and washed with ethanol followed by water. The wet cake was then acidified with 0.1 M sulfuric acid for 4 h and washed thoroughly with water. The material was then dried at 100° C. The HMS material was made with a MPTMS/TEOS molar ratio of 0.2.

3.2.2 Arenesulfonic acid-functionalized mesoporous silica

The method used to synthesize the arenesulfonic acid fiinctionalized materials was also based on a method developed by Stucky and co-workers [23] with co-condensation of the TEOS and CSPTMS. Unlike with the propylsulfonic acid-functionalized material no oxidant was needed since the transformation of the chlorosulfonyl groups to sulfonic groups was acid catalyzed during hydrolysis. The typical synthesis procedure was similar to that given above except that following the TEOS prehydrolysis CSPTMS was added. The mixture was stirred for 20 h at 50 °C and then aged for an additional 24 h in a 100 °C oven with no stirring. The solid product was then recovered and extracted as previously described.

Only a material with a CSPTMS/TEOS molar ratio of 0.1 was made for the study.

3.2.3 Butylcarboxylic acid-functionalized mesoporous silica

The synthesis of the carboxylic acid-functionalized silica was based on the procedure published by Yang et al. [24] in which TEOS and CPTMS were co-condensed in the presence of Pluronic 123. In a typical synthesis, 8 g of Pluronic 123 was dissolved in a mixture of 260 ml de-ionized water with 40 ml concentrated HC1. CPTMS was added following dissolution of the Pluronic 123. After a 30-min delay, 14.9 ml of TEOS was added to the mixture, which was then stirred for 20 h at 50 °C. The mixture was then aged for 24 h in a 90 °C oven with the resulting solid material recovered by filtration and air dried. The material was then refluxed in 50 wt% sulfuric acid to remove the template and hydrolyze the cyano groups to carboxylic acid groups. The carboxylic acid group concentration then resulted from a CPTMS/TEOS molar ratio of 0.2.

3.2.4 Catalyst characterization

The total acid capacity of the materials was determined by titration with NaOH. In a typical titration, 50 mg of the catalyst material was added to a 0.1 M NaCl solution after an initial pH measurement was recorded. Then, a 0.01 M NaOH solution was titrated into the stirred solution until the initial pH was achieved. The NaOH amount required in this titration was used to calculate the total number of acid sites.

The pKa of the acidic materials was determined using the Gran Plot method with the same NaOH solutions as above [25], In this method, the pH was monitored after the addition 44 of 0.1 ml NaOH solution aliquots. The addition was repeated until the equivalence point was reached with the resulting titration volume and pH data used to construct a Gran Plot. The slope of this plot allowed the calculation of the pKa of the acid being analyzed. The amount of catalyst used in each titration was adjusted to a uniform low level in an effort to limit the error introduced by the concentration dependence of the solution activity coefficients.

Adsorption-desorption isotherms for the materials were determined using a

Micrometrics ASAP 2020 Porosimetry Analyzer. Elemental analysis was conducted on the catalyst samples using a Perkin-Elmer Series II 2400 CHNS analyzer to determine the carbon, nitrogen, and sulfur content. X-ray diffraction measurements were taken with a

Scintag XDS 2000 Powder X-ray diffractometer, and the materials exhibit all the typical

XRD patterns for SBA-15 type materials [26].

3.2.5 Reaction studies

The overall reaction system to be probed in the study (Figure 2) was the hydrolysis of cellobiose as well as the sequential degradation of glucose to degradation products such as hydroxymethyl furfural. The reaction sequence was studied by performing both cellobiose hydrolysis reaction studies and glucose degradation reaction studies.

The cellobiose hydrolysis reactions were conducted in a 300-ml Parr stainless-steel batch reactor. The reactor was equipped with a stirrer and had temperature maintained by the

PLC control of an electric heating mantle and a cold water loop. A 1 wt% solution of cellobiose was placed in the reactor with 0.2 wt% catalyst. This mixture was then heated to the desired temperature and held constant for a prescribed reaction time. Upon reaching it, the reactor was quickly cooled to stop the reaction. Therefore, multiple reaction runs were performed to determine the cellobiose conversion as a function of reaction time. The time 45 zero point was determined by simply bringing the reactor up to the reaction temperature and then quickly cooling the reactor. The reaction results were evaluated using a Hewlett-

Packard 1050 HPLC system with a Hewlett-Packard 1100 refractive index detector. The column used in the HPLC was a Supelcogel Pb+ run at 85 °C with a water mobile phase.

Chemstation software was used to capture the chromatograms and quantify the cellobiose concentrations.

Glucose degradation experiments were performed in independent reaction studies using the same reactor and conditions as for the hydrolysis reactions. A 1 wt% glucose solution was placed in the reactor with 0.2% wt of catalyst. The reaction results were determined by using the same procedure described for the hydrolysis reaction. The glucose concentrations in the samples were analyzed using an YSI2700 Biochemistry Analyzer, This analyzer provides for better measurement of the glucose concentration than could be readily achieved with HPLC since the degradation products has similar retention times to the glucose in the HPLC procedure that was used.

3.3. Results and Discussion

The results from characterization of the organic-inorganic hybrid materials are summarized in Table 1. The textural properties for the materials synthesized with Pluronic

123 were generally similar as seen from the adsorption-desorption isotherms in Figure 2.

While the materials synthesized for the study had similar pore size distributions, the 7.5% propylsulfonic acid and 20% butylcarboxylic acid materials did have slightly lower median pore diameters and the surface area of the 20% butylcarboxylic acid materials was also lower than for the other samples. 46

There are two phenomena occurring in the aqueous phase that have a dramatic impact on the effectiveness of catalytic materials. They are the result of the formation of the hydrated proton. Any acid site that is strong enough to ionize in water will form a hydrated proton, and the strength of the acid will effectively be reduced to that of the hydrated proton.

This is referred to as the leveling effect because it "levels" the strength of any acid strong enough to ionize in the aqueous phase to the strength of the hydrated proton [27]. The second phenomenon is the resulting mobility of the hydrated proton. Protons have abnormally high mobility in aqueous solution [28, 29]. The cause of this unusual transport speed is attributed to a "jumping" mechanism. The details of this mechanism are still being debated [29-31]. When the acid is ionized it produces a hydrated proton that has the ability to be an active site in the solution phase away from the catalyst surface. Simulations of ionization conducted by Paddison [32] indicated that six water molecules are required to shield the proton (as a hydronium ion) from direct electrostatic charges with the anion. The shielding of the proton allows it to move away from the anion and the hydrated proton is free to catalyze reactions in the bulk away from the support surface.

Knowledge of the important attributes of the aqueous phase assists in the selection of the catalyst material. Organically functionalized mesoporous silica materials are a good choice to study aqueous phase reactions because they have excellent hydrothermal stability provided by the combination of the stable silica support and the organic functional group that is resistant to leaching problems. The silica support also offers the ability to control pore sizes in addition to recyclability. The use of organic functional sites provides well defined chemistries [33, 34]. The pKa values in Table 1 show that all three of the sulfonic acids have similar acid strengths. These values indicate that all of these materials have strong enough acidity to completely ionize in solution and will effectively be leveled. Butylcarboxylic acid has a much higher pKa and will not completely ionize in solution. The hydrolysis results shown in

Figure 4 confirm these ideas. The three sulfonic acid materials had excellent and similar catalytic activity. All three of the materials tested were able to hydrolyze greater than 98% of the cellobiose in 1 h at 175° C. The butylcarboxylic acid was not strong enough to ionize in solution and was able to hydrolyze only about 15% of the cellobiose in Ih at 175° C. This performance is the same as water alone at this temperature (hydrothermolysis). Others have found that acid catalysis appears to require a pH below 4 [18] and the butylcarboxylic acid was not able to attain that pH.

The effect of active site concentration was investigated by using materials that had different concentrations of propylsulfonic sites. Table 1 shows the pKa values of the 7.5% and the 15% (molar) propylsulfonic acid silica. The 15% material has a lower pKa than the

7.5% material indicating that the acid strength of the sites is strengthened by closer positioning. The close proximity allows interaction between the sites, resulting in a stronger acidity and lower pKa. This effect of spatial arrangement on the catalyst properties has previously been investigated by Dufaud and Davis [35].

The formation of the hydrated proton and the resulting high mobility prevents some of the control over the active site that was thought to have been achieved through pore size selection. The effect of proton mobility in this reaction system was investigated by synthesizing a material with a much smaller pore diameter than the SBA type materials and comparing the resulting kinetics to those of the SBA materials. An HMS templating method 48 was used to synthesize propylsulfonic functionalized silica with 19 À pores. Changing pore size from 19 to 54 Â did not change the reaction rate or the activation energy. These materials were able to catalyze the hydrolysis reaction at the same rate and the same activation energy. The smaller pore size did not limit the reaction rate, indicating that the proton is sufficiently mobile to travel out of the pore and catalyze the reaction in the bulk phase. These results indicate that the interaction with the support material is likely limited, as the active site is effectively in solution and not limited to the interior of the pore. A recent paper by Dhepe et al. [12] postulated that differences in pore morphology caused a much lower turnover frequency (tof) for resin catalysts. This explanation is suspect due to the sulfonic acid groups completely dissociating in water and not being limited to the pore interior. An alternative reason for the low calculated turnover frequency is the difficulty in accounting for the large mass of water that is associated with the resin (50-60% of the total).

The calculation of kinetic parameters was conducted to allow greater comparison to homogenous acids. The calculations used a first-order expression of the rate law consistent with literature. The hydrolysis of disaccharides and oligosaccharides has been treated as a first order reaction by others [20, 36, 37]. Table 2 lists the rate constants for the hydrolysis and degradation reactions. The activation energy was 133 ± 13 kJ/mol for the 7.5% propylsulfonic functionalized silica and 137 ± 20 kJ/mol for the arenesulfonic functionalized silica. The use of 15% propylsulfonic silica resulted in a slightly lower activation energy of

113 ±17. This change was not significant and was attributed to an increase in the number of active sites. These values were similar to values in the literature for mineral and organic acids [18-20, 38]. These papers reported activation energies for the hydrolysis of cellobiose that range from 110 to 138 kJ/mol. 49

The proton concentration versus the kinetic rate constant was plotted to investigate the influence of pH on hydrolysis rate (Figure 5). This plot demonstrated a linear relationship between hydronium ion concentration and rate constant. This indicated that the hydronium ion was the active site of the catalysis. These results were expected due to results published in the literature that have established hydrogen ions are responsible for cellobiose hydrolysis and they agree with the previous discussion of proton mobility. The study conducted by Mosier et al. [20] used homogenous acid systems that covered a much larger pH range than what was attainable with the acid-functionalized mesoporous silica system.

This study found that the rate constant corresponded with the pH regardless of the proton source[20]. This was attributed to the leveling effect of water. The use of the higher temperature regime (175 °C) was of interest because it allowed direct comparison to many acid hydrolysis studies that have been done on biomass at similar temperatures [16, 20, 36].

The high temperature regime resulted in cellobiose hydrolysis occuring relatively quickly and brought concerns of substrate limitations at longer time scales. The lower temperature runs were valuable by allowing for investigation of the kinetics without these concerns.

Reports in the literature have indicated arenesulphonic groups in solvents without strong leveling effect have greater acidity and greater catalytic activity in a Fries rearrangement

[23].

The glucose degradation experiments were done in separate, but identical conditions. This allowed the hydrolysis and degradation reactions to be decoupled and studied separately.

All of the catalysts performed similarly in regards to glucose degradation. All had degraded about 20% of the glucose after 2 h at 175 °C. Figure 6 shows the degradation 50 percents for the catalysts tested. The 2-h time period is most likely much longer than was needed to hydrolyze the oligosaccharides at this temperature. The carboxylic acid- functionalized silica had a low rate of degradation, however, it was not a very effective hydrolysis catalyst. The 7.5% propylsulfonic acid catalyst was the best overall catalyst in this study due to its low degradation and high hydrolysis rates.

Carbohydrate degradation is a complex process. It involves a number of reactions that can occur on any of the active sites of the carbohydrate. By definition, a carbohydrate is a polyhydroxy aldehyde or ketone. All of the hydroxy! groups are reactive, but the most reactive site of the molecule is at the anomeric center. Examples of possible reactions include mutarotation, isomerization, dimerization, and dehydration. Some of these reactions are readily reversible but it is important to note because the alternate species can also participate in irreversible degradation reactions. These reactions are typically modeled as pseudo-first order [20, 36, 37]. Conner [37] developed a more detailed hydrolysis model that created three separate pathways for levoglucosan, disaccharides, and decomposition products. Mosier et al. [20] suggest a mechanism that involves two separate pathways.

They separated the degradation into a ubiquitous baseline pathway that occurs even in pure water and a second pathway that is acid catalyzed and is a function of the chemistry of the acid anion. Xiang et al. [38] acknowledge the complexity of the mechanism and Conner's model. They proceeded to condense the model into three components: glucose, reversible components, and decomposition products. This condensation required several assumptions that were not rationalized. It is apparent that more investigation needs to be done into the mechanisms of degradation. 51

Glucose degradation rates dependence on proton concentration was investigated by charting their relationship in Figure 7. The correlation to acid concentration was not as strong as with the hydrolysis rate constants. The slopes of the lines at both temperatures were similar and relatively flat. The pH range that these data covers is quite limited. Mosier et al. [20] conducted a study over a much broader pH range with homogeneous acid and found interesting effects that were suggested to be related to anion character. The use of supported acids allows the potential to sequester the anion away from the active site. The activation energy for glucose degradation ranged from 67 kJ/mol for propylsulfonic acid to

84 kJ/mol for arenesulfonic acid. These values are similar to the results of Mosier et al. [20], for maleic acid (72 kJ/ mol) and lower than the value found for sulfuric acid (118 kJ/mol).

The lower activation energy means that the reaction had lower dependence on temperature.

The low activation energy for degradation in relation to hydrolysis indicated that yield could be improved at higher temperatures.

3.4. Conclusion

Organically functionalized mesoporous silica materials can give similar performance to homogenous acids. This is due to the complete ionization of strong acids in water and the high mobility of the hydronium ion. The pH ranges attained with these materials were sufficient to prove adequate hydrolysis while limiting degradation reactions. The kinetics indicated that increased temperatures and decreased reaction times were desirable to improve the selectivity to glucose and limit degradation reactions. The excellent thermo-stabilities of mesoporous silicas and the known functionality of the organic groups confer suitable characteristics to these materials, making them attractive catalysts for hydrolysis reactions. 52

Acknowledgments

This material is based upon work partially supported by the Cooperative State Research,

Education, and Extension Service, U.S. Department of Agriculture, under Agreement Nos.

2002-34188-12035 and 2004-34188-15067, as well as from National Science Foundation grant CTS-0455965.

References

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14. Mbaraka, I.K., et al., Organosulfonic acid-functionalized mesoporous silicas for the esterification of fatty acid Journal of Catalysis, 2003. 219: p. 329-336. 15. Mbaraka, I.K. and B.S. Shanks, Design of multifunctionalized mesoporous silicas for esterification offatty acid Journal of Catalysis, 2005. 229: p. 365-373. 16. Mosier, N.S., et al., Characterization ofDicarboxylic Acids for Cellulose Hydrolysis. Biotechnology Progress, 2001.17: p. 474-480. 17. Sasaki, M., et al., Kinetics and Mechanism of Cellobiose Hydrolysis and Retro-Aldol Condensation in Subcritical and Supercritical Water. Industrial Engineering Chemistry Research, 2002. 41: p. 6642-6649. 18. Bobletter, O., et al., Hydrolysis of Cellobiose in Dilute Sulfuric Acid and Under Hydrothermal Conditions. Journal of Carbohydrate Chemistry, 1986. 5: p. 387-399. 19. Dadach, Z.-E. and S. Kaliaguine, Acid Hydrolysis of Cellulose: Part I. Experimental Kinetic Analysis. The Canadian Journal of Chemical Engineering, 1993. 71: p. 880- 891. 20. Mosier, N.S., CM. Ladisch, and MR. Ladisch, Characterization of Acid Catalytic Domains for Cellulose Hydrolysis and Glucose Degradation. Biotechnology and Bioengineering, 2002. 79: p. 610-618. 21. Margolese, D., et al., Direct Syntheses of Ordered SBA-15 Mesoporous Silica Containing Sulfonic Acid Groups. Chemistry of Materials, 2000. 12: p. 2448-2459. 22. Bossaert, W.D., et al., Mesoporous Sulfonic Acids as Selective Heterogeneous Catalysts for the Synthesis ofMonoglycerides. Journal of Catalysis, 1999. 182: p. 156-164. 23. Melero, J.A., et al., Direct Syntheses of Ordered SBA-15 Mesoporous Materials Containing Arenesulfonic Acid Groups. Journal of Materials Chemistry, 2002. 12: p. 1664-1670. 24. Yang, C.-M., et al., Formation of Cyanide-Functionalized SBA-15 and its Transformation to Carboxylate-Functionalized SBA-15. Physical Chemistry Chemical Physics, 2004. 6: p. 2461-2467. 25. Harris, D C., Quantitative Chemical Analysis. 4th ed. 1995, New York: W. H. Freeman. 26. Kruk, M. and M. Jaroniec, Characterization of the Porous Structure of SBA-15. Chemistry of Materials, 2000.12: p. 1961-1968. 27. Reichardt, C., Solvents and Solvent Effects in Organic Chemistry. Second ed. 1988, New York: VCH Publishers. 28. Levine, I.N., Physical Chemistry. 4th ed. 1995, New York: McGraw-Hill. 29. Agmon, N., The Grotthuss Mechanism. Chemical Physics Letters, 1995. 244: p. 456- 462. 30. Tuckerman, M.E., D. Marx, and M. Parrinello, The Nature and Transport Mechanism of Hydrated Hydroxide Ions in Aqueous Solution. Nature, 2002. 417: p. 925-929. 31. Day, T.J., U.W. Schmitt, and G.A. Voth, The Mechanism of Hydrated Proton Transport in Water Journal of American Chemical Society, 200. 258: p. 12027- 12028. 32. Paddison, S.J., Proton Conduction Mechanisms at Low Degrees of Hydration in Sulfonic Acid-Based Polymer Electrolyte Membranes. Annual Review of Materials Research, 2003. 33: p. 289-319. 54

33. Wight, A.P. and M.E. Davis, Design and Preparation of Organic-Inorganic Hybrid Catalysts. Chemical Reviews, 2002.102: p. 3589-3614. 34. Stein, A., B.J. Melde, and R.C. Schroden, Hybrid Inorganic-Organic Mesoporous Silicates — Nanoscopic Reactors Coming of Age. Advanced Materials, 2000.12: p. 1403-1419. 35. Dufaud, V. and M.E. Davis, Design of Heterogeneous Catalysts via Multiple Active Site Positioning in Organic-Inorganic Hybrid Materials. Journal of American Chemical Society, 2003. 125: p. 9403-9413. 36. Saeman, J.F., Kinetics of Wood Saccarification: Hydrolysis of Cellulose and Decomposition of Sugars in Dilute Acid at High Temperature. Industrial Engineering Chemistry Research, 1945. 37: p. 43-52. 37. Conner, A.H., et al., Kinetic Modeling of the Saccharification of Prehydrolyzed Southern Red Oak, in Cellulose: Structure, Modification and Hydrolysis, R.A. Young and R.M. Rowell, Editors. 1986, Wiley: New York. p. 281-296. 38. Xiang, Q., Y.Y. Lee, and R.W. Torget, Kinetics of Glucose Decomposition during Dilute-Acid Hydrolysis of Lignocellulosic Biomass. Applied Biochemistry and Biotechnology, 2004. 113-116: p. 1127-1138. 55

Table 1 Characteristics of acid-functionalized mesoporous silicas

20% Butyl­ 10% 7.5% 15% 20% carboxylic Arenesulfonic Propylsulfonic Propylsulfonic Propylsulfonic Acid Acid Acid Acid Acid (HMS)

Median Pore 54 56 54 62 19 Diameter (A) Surface Area 560 805 727 765 860 (m2/g)

Pore Volume 0.78 1.14 0.98 1.27 0.40 (cm3/g)

[H+] 1.40 0.86 0.73 1.10 0.69 (mequil/g)

pKa (gran 4.65 2.84 2.86 2.67 2.89 plot)

pH in 4.90 3.00 3.19 2.77 3.20 Solution

Carbon 14.05 10.9 7.67 7.73 (%)

Sulfur 2.50 2.27 4.65 (%)

Nitrogen 0.45 (%) 56

Table 2 Activation energies for cellobiose hydrolysis reaction and glucose degradation reaction (a) from [20] Catalyst k1(kJ/mol) k2 (kJ/mol)

7.5% Propylsulfonic Acid 133 ±13 67± 6

15% Propylsulfonic Acid 113 ± 17 65± 18

10% Arenesulfonic Acid 137 ±20 73± 11

20% Butylcarboxylic Acid 118 ±31 76±20

Sulfuric Acid3 110 ±29.6 118 ±37.5

Maleic Acid3 114 ±9.3 72.6 ± 22.5 Fig. 1 Organic acid-functionalized 58

Fig. 2 Model system: reactions of cellobiose 59

0 0.2 0.4 0.6 0.8 Relative Pressure (P/P0) Fig. 3 BET adsorption-desorption isotherms of mesoporous silcas (+ 7.5% propylsulfonic acid, • 15% propylsulfonic acid, • 10% arenesulfonic acid, A 20% butylcarboxylic acid) 60

100%

E 80%

60%

o 20%

o 0% 0 120 Time (minutes) Fig. 4 Hydrolysis of cellobiose at 175 °C with acid-functionalized mesoporous silicas (+ 7.5% propylsulfonic acid, • 15% propylsulfonic acid, • 10% arenesulfonic acid, À20% butylcarboxylic acid, X hydrothermolysis) 61

0.08

0.07

0.06 145

c 0.05 160 175 "50.04

0.02

0.01

0 0.5 1 1.5 2 H+ Concentration (mM) Fig. 5 Hydrolysis rate constants as a function of hydrated proton (+ 20% butylcarboxylic acid, A 7.5% propylsulfonic acid, • 10% arenesulfonic acid, • 15% propylsulfonic acid) 62

25 120% Butylcarboxylic Acid S 7.5% Propylsulfonic Acid 5-20 DD15% Propylsulfonic Acid c o ES 10% Arenesulfonic Acid •m ' c o ° 10 Q> (/) O O 3 5 O

0 30 60 120 Time (min) Fig. 6 Glucose degradation at 175 °C with acid-functionalized mesoporous silicas 63

2.5 - - 145 C - 160 C —175 C -j

* 1.5 c

Ë • o 1 E, JCCM • 0.5 • »

I I I 0.2 0.4 0.6 0.8 1 1.2 1.4 1.6 1.8 H+ Concentration (mM)

+ Fig. 7 Degradation rate constants as a function of [H30 ] (+ 20% butylcarboxylic acid, A 7.5% propylsulfonic acid, • 10% arenesulfonic acid, • 15% propylsulfonic acid) 64

CHAPTER 4

Monosaccharide Degradation Kinetics during Organic-Inorganic Mesoporous

Silica Catalyzed Hydrolysis Conditions

4.1. Introduction

The use of homogeneous acid catalysts for the hydrolysis of oligosaccharides is limited by the lack of ability to separate it from the products and its propensity to form degradation products. Heterogeneous catalysts such as organically functionalized mesoporous silica are effective for the hydrolysis of disaccharides and starches [1,2]. The use of these materials provides a solution to the separation problems; however, product degradation issues remain. This paper will investigate the kinetics of degradation in the heterogeneous catalyst system using the three most common sugar residues found in corn products: glucose, xylose, and arabinose [3], There have been several studies done on degradation of glucose with a variety of homogeneous acids [4-14]. There have been no studies on degradation kinetics with solid acid catalyst systems and only a few studies have included the pentose sugars [9, 15, 16].

The need for understanding the acid-catalyzed hydrolysis system is driven by the ethanol industry. Ethanol production capacity in the United States is growing at an impressive rate. New plants and plant expansions currently under construction will increase capacity by 2 billion gallons[17]. The growth of the ethanol industry is fueled by high petroleum prices and the desire to replace petroleum fuels with a renewable source. The rapid growth of this industry is also presenting a major challenge, as its co-product streams 65 are increasing proportionally. Dry-grind plants comprise the majority of the new plants being constructed in the United States due to their lower capital requirements [18]. Dry grind plants produce distiller's dry grain as a co-product in equal proportion to ethanol. The sudden increase in production of co-product streams is threatening to overwhelm the market and is putting downward pressure on prices. The price of the distiller's dry grain co-product has a significant impact on the profitability of ethanol production [3].

Distiller's dry grain is composed of mostly lignocellulosic material, which can be converted into fermentable sugars with saccharification [3], Converting these materials to sugars and fermenting to alcohol would improve ethanol yield and simultaneously reduce and upgrade the by-product stream by increasing protein concentration [19]. Distiller's dry grain has a significant content of hemicellulose similar to other lignocellulosic feedstocks. The hemicellulosic content of most lignocellulosic feedstocks ranges from 16 to 50% [10, 20].

The large amount of hemicellulose in distiller's grain translates to a pentose sugars content of

39% of the total carbohydrates present [21]. The large fraction of total carbohydrates present as pentose sugars is important, and these materials must be utilized to make the economics feasible [10, 22].

The complex structure of hemicellulose is difficult to hydrolyze enzymatically.

Hemicellulose is a highly branched heteropolymer having several distinct linkages, requiring a cocktail of at least seven types of enzymes to completely hydrolyze hemicellulose into monosaccharides[23 ]. The assortment of enzymes required drives up the cost of the enzyme system. Acid catalysts can quickly hydrolyze oligosaccharides from a variety of sources[l,

2, 7, 24-29], Acid catalysts are not specific and can catalyze the hydrolysis of all the 66 glycosidic and ester linkages present. The versatility of chemical catalysis is a significant advantage in the hydrolysis of hemicellulosic materials.

4.2. Experimental

Organic-inorganic hybrid mesoporous silicas with three different types of organic acid groups were synthesized for the study. Shown in Figure 1 are schematics of the propylsulfonic, phosphonic, and butylcarboxylic acid moieties used in the catalytic materials.

The precursor silane species employed in the synthesis of the catalytic materials, tetraethoxysilane (TEOS) (98%, Aldrich), (3-mercaptopropyl)trimethoxysilane (MPTMS)

(85%, Acros), diethylphosphatoethyltriethoxysilane (DEPTES) (95%, Gelest), and 3- cyanopropyltriethoxysilane (CPTES) (98%, Aldrich), were used as received. The surfactant used in the supramolecular assembly of the organic-inorganic hybrid materials was Pluronic

123 (BASF), a tri-block copolymer of polyethylene oxide-polypropylene oxide-polyethylene oxide with the molecular structure PEO20-PPO70-PEO20 (MW = 5800). Glucose (99%,

Acros), xylose (Acros), and arabinose (Sigma) were used in this study without modification.

Mesoporous silica materials synthesis methods were modifications of those of

Margolese (propylsulfonic acid) [30] and Yang (butylcarboxylic acid) [31]. The phosphonic acid functionalized material was synthesized by dissolving PI23 into a 1.9M hydrochloric acid solution. TEOS was added and allowed to hydrolyze for 1 h at 50° C. DEPTES was added and the solution was stirred for 20 h at 50° C. The solution was then aged at 100° C for 24 h in static conditions. Solids were removed from solution by filtration and refiuxed in ethanol for 24 h to remove the template. The hydrolysis experiments were conducted in a Parr 300-ml stainless-steel batch reactor, equipped with a glass liner and an impeller-type mixer. The reactor temperature was controlled using a programmable logic controller (PLC) connected to both a heating jacket and an internal cooling loop. In the degradation experiments, glucose, arabinose, and xylose were individually dissolved in deionized water in 0.01, 0.02, and 0.05 weight ratios (sugar: liquid). The initial condition was established by extraction of a sample after reaction temperature was reached to minimize the effect of heat up time. Samples were taken every

30 min over a period of 120 min. Temperatures of 145,160, and 175°C were used for degradation experiments.

The remaining sugars and degradation products were analyzed with an HPLC system

(Agilent 1100) using a Hewlett-Packard refractive index detector. The mobile phase was ultrapure water with a flow rate of 0.4 ml/min, and the column was a Supelcogel Ca+

(Supelco). The column temperature was set at 85°C Data were collected and analyzed with

Chemstation software (Agilent).

The total acid capacities and pka of the materials were determined by titration with

NaOH. Titrations were conducted with a 798 MPT Titrino automatic titrator (Metrohm). In a typical titration, 50 mg of the catalyst material was added to a 0.001 M NaCl solution then

0.01 M NaOH solution was titrated into the stirred solution until the endpoint was detected.

The NaOH required in this titration was used to calculate the total number of acid sites.

Analysis of the data was done with Tiamo software (Metrohm). The pKa of the material was determined with the detection of the equivalence point. These calculations were checked using the Gran Plot method [32]. The slope of this plot allowed the calculation of the pKa of the acid being analyzed. The amount of catalyst used in each titration was adjusted to a 68 uniform low level in an effort to limit the error introduced by the concentration dependence of the solution activity coefficients.

Adsorption-desorption isotherms for the materials were determined by using a

Micrometrics ASAP 2020 porosimetry analyzer.

4.3. Results and Discussion

The results from characterization of the organic-inorganic hybrid materials are summarized in Table 1. These materials were selected to span a range of acid strengths in an effort to determine the effect of acid strength on these reactions. The kinetics of degradation were determined for glucose, xylose, and arabinose at conditions known to be effective for hydrolysis of disaccharides[l]. The calculation of kinetic constants was done assuming a first-order reaction [4, 6, 11, 13, 14].

All three monosaccharides demonstrate a reaction rate minimum that occurs with very low acid concentration. Figures 2, 3, and 4 show the rate constants at 175 °C as a function of proton concentration for arabinose, xylose, and glucose, respectively. The plots suggested the existence of pH-controlled kinetic regimes. The values from 0.01 to 2 mM were from the current study using acid-functionalized mesoporous-silica; the remaining values were from literature studies of hydrothermolysis or dilute acid hydrolysis degradation studies[9, 13, 16].

The arabinose degradation rate data in Figure 2 only shows the use of mesoporous silica and hydrothermolysis data, because no dilute-acid degradation data was found in the 69 literature. The highest rate in this figure is for the hydrothermolysis condition, however the rate was showing a clear upward trend with increasing acidity.

The xylose degradation rate data in Figure 3 shows the widest range of acidities available, from hydrothermolysis to 1.5% sulfuric acid. This figure clearly shows a minimum in the range obtained by the acid-functionalized mesoporous-silica. The rate increased from this minimum with increased proton concentration.

The glucose degradation data in Figure 4 shows the increase of degradation with increasing proton concentration. Rate data for the degradation in hydrothermolysis conditions were not available in the literature. There are accounts of higher rates of degradation with hydrothermolysis conditions than with dilute acid hydrolysis conditions^,

8]. It is expected that a rate profile for glucose degradation would be similar to that of xylose if the data for hydrothermolysis were included in the plot.

The existence of pH-controlled kinetic regimes can be discussed in the context of water chemistry and specifically with regard to the ion product. The ion product of water

(ATJ is temperature dependant and increased in value as the temperature was raised. For example, the ion product increased from -13.995 at 25 °C to -11.432 at 175 °C [33]. The

+ increased concentrations of H30 and OH caused the increase in amphoteric character of water at high temperature [34]. The dual role of water at high temperatures can be useful in many reaction systems, but in the degradation system it increased the overall degradation rate by catalyzing both acid- and base-catalyzed degradation reactions[8]. The addition of small amounts of acid limits the occurrence of base-catalyzed reactions[8], effectively lowering the observed degradation rate. This regime was a very narrow one in terms of pH and ended when the heat-generated hydroxide ions were titrated with the added acid. Addition of acid after titration of hydroxide ions increased the degradation rate proportional to increases in hydrated proton concentration.

The existence of distinct regimes of pH dependence in similar systems has been noted by other authors. Xiang et al. [6] studied glucose degradation at 200 °C and found degradation rates of glucose were proportional to hydrated proton concentration in the pH range of 1.5 to 2.2, but discovered a "surprising" leveling off of the rate between pH 2.2 and

7.0. They did not provide data in this region but noted that the rate of degradation at pH 7 was higher than the rate at pH 2.2. Watanabe et al.[8] studied glucose degradation reactions at 200 °C and found addition of acid or base catalyst changed the product slate. Mosier et al.

[13] studied the use of organic acids for cellulose hydrolysis and found distinct regimes of degradation reactions and suggested that there are two different mechanisms for glucose degradation. Conner et al. [11] reported non-acid-catalyzed reactions become significant below 0.05 M and correlations between proton concentrations and degradation rates were not valid, because non-acid-catalyzed reactions dominated.

Figure 5 shows the degradation rate constants at 175 °C for all three monosaccharides and all three catalyst materials used in this study. Arabinose had the highest rate of degradation of the three monosaccharides in this study. The lower stability of arabinose relative to xylose was expected because xylose can assume the highly stable 4C, conformation with all bulky groups equatorial [35]. Comparison of arabinose degradation kinetic data was available from only one author [15, 16]. The frequency factors and activation energies of arabinose degradation from the current study as well as literature values taken from studies on the autohydrolysis of corn cobs and almond shells are shown in

Table 2. The values calculated in the current study were slightly lower than the values 71 calculated for autohydrolysis for reasons discussed earlier. The range of pH covered by the hybrid-silica catalyst was quite narrow and data from higher acid concentrations would assist in drawing conclusions concerning at which concentration pH-dependant degradation begins.

The overall trend of kinetic behavior for arabinose degradation has been observed in the degradation of glucose [6, 13].

Table 3 shows the frequency factor and activation energy values for the current study as well as autohydrolysis and sulfuric acid hydrolysis values from the literature [9, 15, 16].

The range of data covering xylose degradation is the most complete for the purposes of this comparison; however, the data are from a few authors. The previous studies covered a relatively wide range of acidity with a small variation in rates. The relatively wide variation in kinetics of degradation of arabinose and xylose originating from corn cobs compared to almond shells with the studies being performed by the same group of authors [15, 16], indicates the difficulty of regulating hydrothermolysis reactions as well as the difficulty in decoupling degradation kinetics from the other reactions occurring during lignocellulosic feed hydrolysis.

Xylose and arabinose have higher activation energies for degradation than has been reported for glucose. Their lower stability relative to glucose is expected based on observations in the literature [9,15, 16, 36]. There were no explanations given in the literature and Qian et al. [36] even noted that glucose would be more reactive than xylose according to their simulation work. The lower stability and particularly the higher activation energy are problematic when attempting to optimizing lignocellulosic treatment conditions because the activation energies of xylose and arabinose degradation are similar to the 72 hydrolysis activation energy of cellobiose[13] and xylo-oligosaccharides [37], eliminating the ability to improve selectivity by increasing reaction temperature.

Glucose degradation has been studied by a number of authors under a variety of conditions [4, 6-8, 11, 13, 14]. Table 4 shows the frequency factor and activation energy of this reaction calculated in this study as well as values from other authors [9, 13]. Glucose degradation exhibited the pH-dependent degradation regimes that have been discussed previously. The mechanism of glucose degradation has received attention from other authors, but the details of the mechanism are being debated [36, 38]. It is important to note the difference between degradation data taken from model and "real" systems. The model systems starting with glucose [6, 13] tend to have lower degradation rates than systems that started with cellulosic or lignocellulosic materials [9, 14]. The difference in reported values is perhaps due to the presence of other reactants, such as solublized lignin [6], in the "real" system or from error in decoupling the kinetics in sequential reactions. The presence of acid- soluble lignin adds other potential reactions to the system; however, these reactions are beyond the scope of this study and will not be evaluated here.

Glucose degradation is actually a collection of several reactions including those catalyzed by both base acid, as previously discussed. These reactions include reversible and irreversible reactions. The reversible reactions occur much faster than the irreversible reactions and include isomerization, dehydration, and reversion. Figure 6 shows a model developed by Conner et al. [11] showing the interplay of the reactions involved in cellulose hydrolysis and glucose decomposition. This model was simplified by Xiang et al. [6] into groupings of glucose and reversible components both contributing to degradation products. Isomerization is a base-catalyzed reaction that occurs in hydrothermolysis or low acid situations. Isomerization is considered an important reaction because it generates ketoses, which are more suseptible to dehydration reactions. Isomerization is not included in

Conner's model, but is a constituent of the reversible components of Xiang's model.

Dehydration is the most prevalent acid-catalyzed reaction and is responsible for the generation of furfurals during hydrolysis conditions. The furfural formed can undergo further reactions that can lead to fragmentation into formic acid and levulinic acid [35,39].

Reversion reactions refer to the formation of disaccharides and oligosaccharide from monsaccharides. 1,6-linked glucosides such as gentiobiose and isomaltose are preferentially formed due to steric hindrances limiting other products. Xiang et al. [6] found that products of these reversible reactions are susceptible to decomposition at rates approximately equal to glucose.

The most prevalent irreversible base-catalyzed reaction is retro-aldol condensation, which generates products including glyceraldehyde, dihydroxyacetone, and erythrose. This reaction is typically not observed in acid hydrolysis conditions.

Dehydration of glucose to 5-hydroxymethylfurfuraldehyde is considered the primary irreversible degradation reaction; however, this product accounts for only a portion of the observed sugar loss during pretreatment applications. Qian et al. [36] simulated acid degradation in glucose and xylose, and found the rate-limiting step in sugar degradation was the protonation of the hydroxy! groups on the sugar ring. They found in simulations of degradation reactions the structure of the water molecule plays a significant role in the pathways. Xylose was only reactive at the OH-2 and OH-3 groups and glucose was found to be reactive at the OH-2, OH-3, and OH-4 groups[36]. They acknowledged that experimental 74 evidence had shown that xylose degrades at a faster rate than glucose, but were unable to elucidate the cause of this behavior. Reactions that were conducted in vacuum showed glucose to be more reactive than xylose [38]. This finding indicates the importance of the water structure in the reaction. A possible explanation for the increase reactivity of xylose would be the observed C-3-C-4 bond cleavage that occurred after OH-2 protonation and led to additional degradation products. Qian et al. found in both molecules protonations initiated at the OH-2 led to the most common degradation products, HMF and furfural. The reason for the increased reactivity was caused by high proton affinity at the OH-2.

Investigation into the order of the mechanism was conducted by performing degradation tests at monosaccharide concentrations of 1, 2, and 5% (wt/wt). Table 5 shows the activation energies calculated assuming a first-order reaction. The values do not indicate any significant change correlating to the change in concentration. This result is expected because authors typically model degradation as a pseudo-first-order reaction and achieve good agreement with experimental results [6, 7, 14]. Watanabe et al. [8] suggested that the product slate may shift from monomolecular degradation to condensation products as concentration is increased. Exploration of the product slate in relation to monosaccharide concentration may be useful if a particular degradation product is found to be desirable.

4.4. Conclusion

The use of acid-functionalized mesoporous-silica as a catalyst for hydrolysis of glycosidic bonds provides an optimum concentration of protons that limits the formation of degradation products to a rate lower than occurs in pure water. The pentose sugars arabinose and xylose are much more susceptible to degradation, but they exhibit the same pH-optimum 75 behavior as glucose does. The careful selection of pH with very dilute acid will give optimum yield of monosaccharides from lignocellulosic hydrolysis.

References

1. Bootsma, J. and B. Shanks, Disaccharide Hydrolysis Using Organic-Inorganic Hybrid Mesoporous Silica Catalysts. In Press, 2006. 2. Dhepe, P.L., et al., Hydrolysis of sugars Catalyzed by Water-Tolerant Sulfonated Mesoporous Silicas. Catalysis Letters, 2005. 102: p. 163-169. 3. Gulati, M., et al., Assessment ofEthanol Production Options for Corn Products. Bioresource Technology, 1996. 58: p. 253-264. 4. Bergeron, P., C. Benham, and P. Werdene, Dilute Sulfuric Acid Hydrolysis of Biomass for Ethanol Production. Applied Biochemistry and Biotechnology, 1989. 20- 21: p. 119-134. 5. Bienkowski, P.R., et al., Correlation of Glucose Degradation at 90 to 19(fC in 0.4 to 20% Acid. Chemical Engineering Communications, 1987. 51: p. 179-192. 6. Xiang, Q., Y.Y. Lee, and R.W. Torget, Kinetics of Glucose Decomposition during Dilute-Acid Hydrolysis of Lignocellulosic Biomass. Applied Biochemistry and Biotechnology, 2004.113-116: p. 1127-1138. 7. Bobletter, O., et al., Hydrolysis of Cellobiose in Dilute Sulfuric Acid and Under Hydrothermal Conditions. Journal of Carbohydrate Chemistry, 1986. 5(3): p. 387- 399. 8. Watanabe, M., et al., Glucose Reactions with Acid and Base Catalysts in Hot Compressed Water at 473 K. Carbohydrate Research, 2005. 340: p. 1925-1930. 9. Bhandari, N., D.G. Macdonald, and N.N. Bakhshi, Kinetic Studies of Corn Stover Saccharification Using Sulphuric Acid. Biotechnology and Bioengineering, 1984. 26: p. 320-327. 10. Sun, Y., Hydrolysis of Lignocellulosic Materials for Ethanol Production: A Review. Bioresource Technology, 2002. 83: p. 1-11. 11. Conner, A.H., et al., Kinetic Modeling of the Saccharification of Prehydrolyzed Southern Red Oak, in Cellulose: Structure, Modification and Hydrolysis, R.A. Young and R.M. Rowell, Editors. 1986, Wiley: New York. p. 281-296. 12. Mosier, N.S., et al., Characterization of Dicarboxylic Acids for Cellulose Hydrolysis. Biotechnology Progress, 2001.17: p. 474-480. 13. Mosier, N.S., C.M. Ladisch, and M.R. Ladisch, Characterization of Acid Catalytic Domains for Cellulose Hydrolysis and Glucose Degradation. Biotechnology and Bioengineering, 2002. 79(6): p. 610-618. 14. Saeman, J.F., Kinetics of Wood Saccarification: Hydrolysis of Cellulose and Decomposition of Sugars in Dilute Acid at High Temperature. Industrial Engineering Chemistry Research, 1945. 37(1): p. 43-52. 76

15. Nabarlatz, D., X. Farriol, and D. Montane, Autohydrolysis of Almond Shells for the Production ofXylo-oligosaccharides: Product Characteristics and Reaction Kinetics. Industrial Engineering Chemistry Research, 2005. 44: p. 7746-7755. 16. Nabarlatz, D., X. Farriol, and D. Montane, Kinetic Modeling of the Autohydrolysis of Lignocellulosic Biomass for the Production of Hemicellulose-Derived Oligosaccharides. Industrial Engineering Chemistry Research, 2004. 43: p. 4124- 4131. 17. www.ethanolrfa.org. Removing Ethanol Tariff Not the Answer to High Gas Prices. 2006 May 8, 2006 [cited; Available from: www.ethanolrfa.org. 18. Belyea, R.L., K.D. Rausch, and M.E. Tumbleson, Composition of Corn and Distillers Grains with Solubles from Dry Grind Ethanol Processing. Bioresource Technology, 2004. 94: p. 293-298. 19. Tucker, M.P., et al., Conversion of Distiller's Grain into Fuel Alcohol and a Higher- Value Animal Feed by Dilute-Acid Pretreatment. Applied Biochemistry and Biotechnology, 2004. 113-116: p. 1139-1159. 20. Mosier, N., et al., Features of Promising Technologies for Pretreatment of Lignocellulosic Biomass. Bioresource Technology, 2005. 96: p. 673-686. 21. Cotta, M., et al. Developing Enzyme Systems for Converting Pretreated DDGS to Fermentable Sugars, in Adding Value to DG from Dry Grind Processes. 2005. Indianapolis, In: Purdue University. 22. Lynd, L.R., C.E. Wyman, and T.U. Gerngross, Biocommodity Engineering. Biotechnology Progress, 1999. 15: p. 777-793. 23. Saulnier, L., et al., Cell Wall Polysaccharide Interactions in Maize Bran. Carbohydrate Polymers, 1995. 26: p. 279-287. 24. Abatzoglou, N. and E. Chomet, Acid Hydrolysis ofHemicellulose and Cellulose: Theory and Applications, in Polysaccharides: Structural Diversity and Functional Versatility, S. Dumitriu, Editor. 1998, Marcel Dekker: New York. p. 1007-1046. 25. Esteghlalian, A., et al., Modeling and Optimization of the Dilute-Sulfuric-Acid Pretreatment of Corn Stover, Poplar and Switchgrass. Bioresource Technology, 1997. 59: p. 129-136. 26. Kunihisa, K.S. and H. Ogawa, Acid Hydrolysis of Cellulose in a Differential Scanning Calorimeter. Journal of Thermal Analysis, 1985. 30: p. 49-59. 27. Lee, Y.Y., P. Iyer, and R.W. Torget, Dilute-Acid Hydrolysis of Lignocellulosic Biomass. Advances in Biochemical Engineering/Biotechnology, 1999. 65: p. 93-115. 28. Bootsma, J.A. and B.H. Shanks, Hydrolysis Characteristics of Tissue Fractions Resulting From Mechanical Separation of Corn Stover. Applied Biochemistry and Biotechnology, 2005.125: p. 27-39. 29. Sheehan, J. and M. Himmel, Enzymes. Energy, and the Environment: A Strategic Perspective on the U.S. Department of Energy's Research and Development Activitities for Bioethanol. Biotechnology Progress, 1999. 15: p. 817-827. 30. Margolese, D., et al., Direct Syntheses of Ordered SBA-15 Mesoporous Silica Containing Sulfonic Acid Groups. Chemistry of Materials, 2000. 12: p. 2448-2459. 31. Yang, C.-M., et al., Formation of Cyanide-Functionalized SBA-15 and its Transformation to Carboxy late-Functionalized SBA-15. Physical Chemistry Chemical Physics, 2004. 6: p. 2461-2467. 77

32. Harris, D C., Quantitative Chemical Analysis. 4th ed. 1995, New York: W. H. Freeman. 33. Marshall, W.L. and E.U. Franck, Ion Product of Water Substance, 0-1000 °C, 1- 10,000 Bars New International Formulation and Its Background. Journal of Physical and Chemical Reference Data, 1981.10: p. 295-303. 34. Watanabe, M., et al., Chemical Reactions of CI Compounds in Near-Critical and Supercritical Water. Chemical Reviews, 2004. 104: p. 5803-5821. 35. Robyt, J.F., Essentials of Carbohydrate Chemistry. Springer Advanced Texts in Chemistry, ed. C.R. Cantor. 1998, New York: Springer-Verlag. 36. Qian, X., et al., Ab Initio Molecular Dynamics Simulations of (i-D-Glucose and ft-D- Xylose Degradation Mechanisms in Acidic Aqueous Solution. Carbohydrate Research, 2005.340: p. 2319-2327. 37. Kamiyama, Y. and Y. Sakai, Rate of Hydrolysis ofXylo-Oligosaccharides in Dilute Sulfuric Acid Carbohydrate Research, 1979. 73: p. 151-158. 38. Qian, X., M.R. Nimlos, and D.K. Johnson, Acidic Sugar Degradation Pathways. An Ab Initio Molecular Dynamics Study. Applied Biochemistry and Biotechnology, 2005. 121-124: p. 989-997. 39. Collins, P.M. and R.J. Ferrier, Monosaccharides: Their Chemistry and Their Roles in Natural products. 1995, Chichester: John Wiley. 78

Table 1 Characteristics of acid-functionalized mesoporous silicas 20% 15% 10% Phosphonic Butylcarboxylic Propylsulfonic Acid Acid Acid Median Pore 54 62 70 Diameter (À)

Surface Area 560 765 492 (m2/g)

Pore Volume 0.78 1.27 0.87 (cm3/g)

[H+] (mequil/g) 1.40 1.1 0.78

pKa (gran plot) 4.65 2.67 3.93

pH in Solution 4.90 2.77 4.05 79

Table 2 Kinetic constants for the degradation of arabinose Frequency Factor Activation Energy Treatment (min1) (kJ/mol) Hydrothermolysis 2.30 x 1015 148 (almond shells) Hydrothermolysis 8.92 x1012 125 (corn cobs) 15% Propylsulfonic 5.01 x 1010 154±19

10% Ethylphosphonic 9.80 x 1010 104±23

20% Butylcarboxylic 7.70 x 1010 111± 27 80

Table 3 Kinetic constants for the degradation of xylose Frequency Factor Activation Energy Acid Used (min'1) (kJ/mol) 0.05 M Sulfuric 6.70x10" 134

0.1 M Sulfuric 2.48x10" 134

1.5 M Sulfuric 3.28x10" 134 Hydrothermolysis 4.23x10" 137 (almond shells) Hydrothermolysis 5.63 x 1012 123 (corn cobs) 15% Propylsulfonic 1.84 x1015 150 ±4

10% Ethylphosphonic 1.22x10" 151±16

20% Butylcarboxylic 2.28x10" 130±14 81

Table 4 Kinetic constants for the degradation of glucose Frequency Factor Activation Energy Acid Used (min1) (kJ/mol) 50 mM Sulfuric Acid 5.70 x 1013 137

94 mM Sulfuric Acid 1.47x10" 137

153 mM Sulfuric Acid 4.50x10" 137

50 mM Maleic Acid 1.66 x 1006 72.6±22.5

15% Propylsulfonic 1.03 x 1006 75±6

10% Ethylphosphonic 9.80 x 1005 104±23

20% Butylcarboxylic 1.19 x 1006 79 ±12 82

Table 5 Activation energies for monosaccharide degradation with propylsulfonic catalyst for 1, 2, and 5% concentration

Activation Energy (kJ/mol)

Monosaccharide 1%(wt/wt) 2%(wt/wt) 5%(wt/wt)

Arabinose 149 153 155

Xylose 148 149 154

Glucose 70 78 80 Fig. 1 Organic acid-functionalized groups 84

0.03

0.025 ; <

0.02

i 0.015

CM JaC 0.01

0.005

0.5 1.5 H+Concentration (mM)

Fig. 2 Degradation rate constants at 175 °C for arabinose as a function of proton Concentration (X Nabarlatz et al. [15], • mesoporous silica) 85

0.09

0.08

0.07

0.06

^ 0.05 E 0.04 CMl 0.03 %

0.02

0.01

50 100 150 H+ Concentration (mM)

Fig. 3 Degradation rate constants at 175 °C for xylose as a fonction of proton concentration (X Nabarlatz et al. [15],, • mesoporous silica,• Bhandari et al.[9]) 86

0.06

0.05

^0.04

i 0.03

CM 0.02

0.01

50 100 150 H+ concentration (mM) Fig. 4 Degradation rate constants at 175 °C for glucose as a function of proton concentration (• mesoporous silica, «Mosier et al. [13], A Bhandari et al. [15]) 87

10 9

8 7 CO CD

X

2

1

0 0 0.5 1 1.5 2 H+ Concentration (mM) Fig. 5 Degradation rate constants using mesoporous silica catalyst at 175 °C as a function of proton concentration (X arabinose, A xylose, • glucose) 88

Easily Levoglucosan Hydrolyzable Cellulose

Decomposition Glucose Products

Resistant Cellulose Disaccharides Glucosides

Fig. 6 Conner's model [11] of cellulose hydrolysis and glucose degradation 89

CHAPTER 5

Hydrolysis of Oligosaccharides from Distiller's Dry Grain Using

Organic-Inorganic Hybrid Mesoporous Silica Catalysts

5.1. Introduction

The dramatic increase in demand for ethanol has resulted in a rapid increase in production capacity. New plants and plant expansions currently under construction will increase capacity by 2 billion gallons [1], Growth in the demand for ethanol has been fueled by high petroleum prices and a desire for energy security. Increases in the production of ethanol result in a proportional increase in co-product production. The desire to rapidly increase production has favored the selection of dry grind technology in new plants due to their lower capital costs [2, 3]. The co-product of dry grind plants is distiller's dry grain and it is produced in equal proportion to ethanol. The primary market for distiller's dry grain has been cattle rations. The sudden increase in production of distiller's grain is putting pressure

on prices due to market saturation. Revenue from distiller's grain is important to the

profitability of ethanol production [4],

The hydrolysis of disaccharides and starches with acid-functionalized mesoporous

silica has been demonstrated [5, 6], Heterogeneous catalysts can be separated and reused.

This results in lower processing cost and eliminates the need for neutralization. The

neutralization of homogeneous acids leads to high levels of salts in the remaining product

and causes palatability problems [7].

Pretreatment of lignocellulosic materials with water [8-11], dilute acid [7, 10],

hydrogen peroxide [12], or ammonia [9, 10] releases soluble oligomers that are not directly 90 fermentable. The oligomer streams require further hydrolysis by a chemical catalyst or enzyme [11]. The solubilization of hemicelluloses as oligomers may in fact be desirable because they are less susceptible to degradation than monosaccharides [13, 14].

Distiller's dry grain has a significant hemicellulose content and hot water pretreatment selectively solublizes hemicellulose. The resulting oligosaccharide stream is rich in xylo-oligosaccharides. These pentose sugars are important to the economics of these processes [15, 16].

The complex structure of hemicellulose is difficult to hydrolyze enzymatically.

Hemicellulose is a highly branched heteropolymer having several distinct linkages requiring a cocktail of at least seven types of enzymes to completely hydrolyze hemicellulose into monosaccharides [17]. The assortment of enzymes required drives up the cost of the enzyme system. Acid catalysts can quickly hydrolyze oligosaccharides from a variety of sources [5,

6, 18-24], Acid catalysts are not specific and can catalyze the hydrolysis of all of the glycosidic linkages present. The versatility of chemical catalysts is a significant advantage in the hydrolysis of hemicellulosic materials.

5.2. Experimental

Hydrolyzate liquor was provided by Nathan Mosier and Rick Hendrickson of Purdue

University. Acid hydrolysis of the liquor was conducted to characterize the carbohydrate content. Propylsulfonic functionalized mesoporous silica was synthesized by method previously described [5, 25]. The precursor silane species employed in the synthesis of the catalytic materials, tetraethoxysilane (TEOS) (98%, Aldrich) and (3-mercaptopropyl)- trimethoxysilane (MPTMS) (85%, Acros) were used as received. Trisyl 300 silica gel (Grace Davison), purified zein (Aéras), sulfuric acid (95.7%, Fisher), hydrochloric acid (37.3%,

Fisher), cellobiose (98%, Sigma), glucose (99%, Acros) xylose (Acros), and arabinose

(Sigma) were used as received.

The hydrolysis experiments were conducted in a 300-ml stainless steel batch reactor

(Parr Instruments), which was equipped with a glass liner and an impeller-type mixer. The reactor temperature was controlled using a programmable logic controller (PLC) connected to both a heating jacket and an internal cooling loop. Pretreatment liquor was injected into the reactor after the desired reaction temperature had been reached with a Lab Alliance Series

1 HPLC pump (Scientific Systems).

All products were analyzed with an HPLC system (Agilent 1100) using a Hewlett-

Packard refractive index detector. The mobile phase was ultrapure water with a pump rate of

0.4 ml/min, and the column was a Supelcogel Ca+ (Supelco). The column temperature was set at 85°C. Data were collected and analyzed with Chemstation software (Agilent).

Initial investigation into the effectiveness of acid-functionalized mesoporous-silica was conducted with cellobiose as the model system. A 1% (wt/wt) solution of cellobiose was made with deionized water and 200 mg of propylsulfonic functionalized silica was added.

Reaction temperatures of 145, 160, and 175 °C were used and reaction times were 120 min with samples taken at 0, 30, 60, and 120 min. The effect of protein in solution was investigated by slurring in lwt% of zein into the solution and the experiments were repeated as previously described. The effectiveness of silca gel titration on removing protein inhibition was conducted by slurring in 1 % silica gel after the addition of zein and filtering the solution. The hydrolysis experiment was repeated as previously described. 92

The procedure for the 72% analytical hydrolysis of the hydrolyzate liquor was 0.320 ml of concentrated H2S04 was added to 0.250 g of hydrolyzate solution and the mixture was allowed to set for 3 h at 20 °C (room temperature). The mixture was diluted with 5 ml of distilled water and titrated with 1 M barium hydroxide solution to a neutral pH. The solution stood for 30 min before being filtered with a Buchner funnel. The final mass was recorded and samples were taken for HPLC analysis.

The procedure used for analytical hydrolysis by the Purdue researchers was based on the National Renewable Energy Laboratory procedure LAP-14. The procedure can be summarized as follows: Biomass samples were blended to a 4% sulfuric acid solution. The solution was autoclaved at 121 °C for 1 h. Samples were analyzed on HPLC with the following modified from LAP-14: Aminex HPX-87H column (Biorad) using 0.6 ml flow rate of 5 mM H2S04 mobile phase and a refractive index detector.

5.3. Results and Discussion

Cellobiose hydrolysis was done with the propylsulfonic acid catalyst to prove the concept and discover optimum conditions for use in later experiments with feed material generated by distiller's dry grain pretreatment. As seen in Figure 1, propylsulfonic acid is an effective catalyst for cellobiose hydrolysis at 160 °C. Literature has indicated fouling of catalyst was a problem when using "real" feeds. It has been suggested that protein in the hydrolyzate solution was responsible for the fouling[26], Zein was added to the model system in an effort to reproduce this effect. The loss of catalyst activity can be seen in Figure

1. Silica gel filtration was conducted after the addition of zein to investigate its ability to remove the inhibitory material. Figure 1 shows most of the catalyst activity was restored after the silica gel pretreatment.

Evaluation of the total amount of oligosaccharides in the hydrolyzate liquor was conducted through comparison of monosaccharide content before and after analytical acid hydrolysis. The sugar content of the hydrolyzate as received is shown in Table 1, while the sugar content of the hydrolyzate after acid hydrolysis is shown in Table 2. The first column in Table 2 contains results of the 4% acid hydrolysis conducted at Purdue University and the second column shows the results of the 72% acid hydrolysis conducted at Iowa State

University. The lower values calculated at Iowa State are likely due to the formation of precipitate during shipping and storage of the product. This material was not re-suspended for the hydrolysis. The right columns in both tables show the percentage of the original carbohydrate present in the distiller's grain represented in each component. The overall effectiveness of the hot water pretreatment was 35% solublization of the original carbohydrate and all three types of carbohydrates were equally solubilized. Over one-half of the carbohydrate (52.8%) solubilized is represented in the pentose fractions, which emphasizes their importance. The distribution of the carbohydrates as monosaccharides and oligosaccharides was as follows: glucose was present as 6.7% glucose, 25% cellobiose, and

68.3% higher oligosaccharides; xylose was present as 30.7% xylose, 31.7% xylobiose, and

37.6% higher oligosaccharides; and arabinose was present as 44.9% arabinose and 55.1% oligosaccharides.

Mesoporous-silica-catalyzed hydrolysis was effective at 175 and 160 °C (Figures 2 and 3). The concentration of the carbohydrates is one order of magnitude lower than the original hydrolyzate because the hydrolyzate liquor was diluted 9:1 (water:liquor) for the 94

reaction. The 175 °C hydrolysis condition could achieve nearly complete hydrolysis of the

glucan into glucose and cellobiose at the end of the reaction time. The cellobiose

concentration was beginning to decline due to hydrolysis into glucose; however the majority of cellobiose remained. The 160 °C hydrolysis was considerably less effective in

hydrolyzing glucan; however, the xylan and arabinan hydrolysis was similar to that at 175 °C

with less degradation of the monosaccharides. The 145 °C hydrolysis was not effective with

any of the materials and the results are not shown.

Silica gel filtration was conducted before hydrolysis in an effort to prevent fouling

and deactivation of the catalyst by proteins and minerals in the liquor[26]. The hydrolysis

experiments were repeated following filtration with a 3% body feed of the silica gel using the

same conditions as previously discussed. The results of these experiments are shown in

Figures 4 and 5. The silica pretreatment did enhance the hydrolysis of cellobiose to glucose at 175 °C, but the hydrolysis of the other materials did not appear to be affected. The use of silica at 160 °C did not appear to affect the hydrolysis rate.

To provide a baseline comparison of hydrolysis without catalyst, the experiment was repeated at 175 °C without catalyst. As seen in Figure 6, the hydrolysis rate of the glucans and xylans were much slower without catalyst.

The hydrolysis of lignocellulose is mechanistically complex. The decoupling of kinetics is very difficult because of the heterogeneity of the material. Attempts to understand the kinetics have led to theories that include at least two types of cellulose [27] and two types of hemicellulose [28]. The current study is further complicated by the presence of residual starch [7] in distiller's dry grain. The present material is likely to contain five different materials with distinct first-order hydrolysis kinetics. 95

The prevalence of degradation of monosaccharides was more apparent if the data are presented as percent conversion, based on the total glucan, xylan, and arabinan in the liquor.

Figures 7 and 8 show the data for the hydrolysis at 175 °C and 160 °C recalculated from

Figures 2 and 3, respectively. The conversion rate in the 175 °C experiment appeared to decrease as time proceeds, but the conversion rate in the 160 °C experiment remained constant. The high susceptibility of arabinan to hydrolysis has been noted before [8].

Arabinose yield peaked with the 175 °C hydrolysis after only 15 min of reaction time, but did not peak until after 1 h at 160 °C. The yields of xylose and glucose continued to increase at the end of the 90 minute reaction time in the 160 °C experiment. The xylose yield was highest after 45 min reaction time at 175 °C and the glucose yield appeared to level off. Kim et al. [26] found a much lower activation energy for hydrolysis of cellobiose than degradation of glucose, indicating that the maximum yield for desired products may be achieved at higher temperatures and shorter reaction times. Further examination into the kinetics of degradation and hydrolysis of hemicellulose will be needed to find optimum conditions.

5.4. Conclusion

Acid-functionalized mesoporous-silica was shown to be an effective catalyst for the hydrolysis of oligosaccharide streams from distiller's dry grain pretreatment. The use of silica gel filtration was effective in removing materials causing catalyst fouling. The hydrolysis of arabinans and xylans occured at lower temperatures and an optimum temperature will need to achieve adequate glucan hydrolysis while limiting the degradation of pentose sugars. 96

References

1. www.ethanolrfa.org. Removing Ethanol Tariff Not the Answer to High Gas Prices. 2006 May 8, 2006 [cited; Available from: www.ethanolrfa.org. 2. Rajagopalen, S., et al., Enhancing Profitability of Dry Mill Ethanol Plants. Applied Biochemistry and Biotechnology, 2005.120: p. 37-50. 3. Belyea, R.L., K.D. Rausch, and M.E. Tumbleson, Composition of Corn and Distillers Grains with Solubles from Dry Grind Ethanol Processing. Bioresource Technology, 2004. 94: p. 293-298. 4. Gulati, M., et al., Assessment of Ethanol Production Options for Corn Products. Bioresource Technology, 1996. 58: p. 253-264. 5. Bootsma, J. and B. Shanks, Disaccharide Hydrolysis Using Organic-Inorganic Hybrid Mesoporous Silica Catalysts. In Press, 2006. 6. Dhepe, P.L., et al., Hydrolysis of sugars Catalyzed by Water-Tolerant Sulfonated Mesoporous Silicas. Catalysis Letters, 2005.102: p. 163-169. 7. Tucker, M.P., et al., Conversion of Distiller's Grain into Fuel Alcohol and a Higher- Value Animal Feed by Dilute-Acid Pretreatment. Applied Biochemistry and Biotechnology, 2004.113-116: p. 1139-1159. 8. Bura, R., et al., SO2 -Catalyzed Steam Explosion of Corn Fiber for Ethanol Production. Applied Biochemistry and Biotechnology, 2002. 98-100: p. 60-72. 9. Mosier, N., et al., Features of Promising Technologies for Pretreatment of Lignocellulosic Biomass. Bioresource Technology, 2005. 96: p. 673-686. 10. Wyman, C.E., et al., Coordinated development of leading biomass pretreatment technologies. Bioresource Technology, 2005. 96: p. 1959-1966. 11. Mosier, N.S., et al., Industrial Scale-Up of pH-Controlled Liquid Hot Water Pretreatment of Corn Fiber for Fuel Ethanol Production. Applied Biochemistry and Biotechnology, 2005.125: p. 77-97. 12. Sun, R., et al., Comparative Studies of Hemicelluloses Solubilizaed during the Treatments of Maize Stems with Peroxymonosulfuric Acid, Peroxyformic Acid, Peracetic Acid, and Hydrogen Peroxide. Holzforschung, 2000. 54: p. 349-356. 13. Saeman, J.F., Kinetics of Wood Saccarification: Hydrolysis of Cellulose and Decomposition of Sugars in Dilute Acid at High Temperature. Industrial Engineering Chemistry Research, 1945. 37(1): p. 43-52. 14. Abatzoglou, N., J. Bouchard, and E. Chomet, Dilute Acid Depolymerization of Cellulose in Aqueous Phase: Experimental Evidence of the Significant Presence of Soluble Oligomeric Intermediates. The Canadian Journal of Chemical Engineering, 1986. 64: p. 781-786. 15. Lynd, L.R., C.E. Wyman, and T.U. Gerngross, Biocommodity Engineering. Biotechnology Progress, 1999. 15: p. 777-793. 16. Sun, Y., Hydrolysis of Lignocellulosic Materials for Ethanol Production: A Review. Bioresource Technology, 2002. 83: p. 1-11. 17. Saulnier, L., et al., Cell Wall Polysaccharide Interactions in Maize Bran. Carbohydrate Polymers, 1995. 26: p. 279-287. 18. Bobletter, O., et al., Hydrolysis of Cellobiose in Dilute Sulfuric Acid and Under Hydrothermal Conditions. Journal of Carbohydrate Chemistry, 1986. 5: p. 387-399. 97

19. Abatzoglou, N. and E. Chornet, Acid Hydrolysis of Hemicellulose and Cellulose: Theory and Applications, in Polysaccharides: Structural Diversity and Functional Versatility, S. Dumitriu, Editor. 1998, Marcel Dekker: New York. p. 1007-1046. 20. Esteghlalian, A., et al., Modeling and Optimization of the Dilute-Sulfuric-Acid Pretreatment of Corn Stover, Poplar and Switchgrass. Bioresource Technology, 1997. 59: p. 129-136. 21. Kunihisa, K.S. and H. Ogawa, Acid Hydrolysis of Cellulose in a Differential Scanning Calorimeter. Journal of Thermal Analysis, 1985. 30: p. 49-59. 22. Lee, Y.Y., P. Iyer, and R.W. Torget, Dilute-Acid Hydrolysis of Lignocellulosic Biomass. Advances in Biochemical Engineering/Biotechnology, 1999. 65: p. 93-115. 23. Bootsma, J. A. and B.H. Shanks, Hydrolysis Characteristics of Tissue Fractions Resulting From Mechanical Separation of Corn Stover. Applied Biochemistry and Biotechnology, 2005. 125: p. 27-39. 24. Sheehan, J. and M. Himmel, Enzymes. Energy, and the Environment: A Strategic Perspective on the U.S. Department of Energy's Research and Development Activitities for Bioethanol. Biotechnology Progress, 1999. 15: p. 817-827. 25. Margolese, D., et al., Direct Syntheses of Ordered SBA-15 Mesoporous Silica Containing Sulfonic Acid Groups. Chemistry of Materials, 2000. 12: p. 2448-2459. 26. Kim, Y., et al., Plug-Flow Reactor for Continuous Hydrolysis of Glucans and Xylans from Pretreated Corn Fiber. Energy and Fuels, 2005.19: p. 2189-2200. 27. Conner, A.H., et al., Kinetic Modeling of the Saccharification of Prehydrolyzed Southern Red Oak, in Cellulose: Structure, Modification and Hydrolysis, R.A. Young and R.M. Rowell, Editors. 1986, Wiley: New York. p. 281-296. 28. Belkacemi, K., et al., Phenomenological Kinetics of Complex Systems: Mechanistic Considerations in the Solubilization of Hemicelluloses following Aqueous/Steam Treatments. Industrial Engineering Chemistry Research, 1991. 30: p. 2416-2425. 98

Table 1 Monosaccharide and disaccharide concentration in pretreated hydrolyzate liquor and percent of total carbohydrate Sugar g/L % of Original Cellobiose 3.7 4.1 Xylobiose 3.2 3.5 Glucose 1.0 1.1 Xylose 3.1 3.4 Arabinose 3.1 3.4 99

Table 2 Monosaccharide concentration after 4% acid hydrolysis and 72% acid hydrolysis and percent of total carbohydrate 4% Acid 72% Acid % of Original Sugar g/L g/L Carbohydrate Glucose 15.1 14.0 16.6 Xylose 10.1 8.8 11 Arabinose 7.0 5.5 7.5 100

100

.p

"O

50

25

0 30 60 90 120 Time (min) Fig. 1 Glucose yield from cellobiose hydrolyis with propylsulfonic acid catalyst at 160°C (e pure cellobiose, • zein added, • zein added followed by silica gel filtration) 101

0.9

50.6 10.5 C . . o)0.4 0.3 0.2

90 Time (min) Fig. 2 Monosaccharide and cellobiose concentration during propylsulfonic acid-catalyzed hydrolysis at 175°C (• cellobiose, • glucose, A xylose, X arabinose) 102

1

0.9

0.8

O 0.7 0.6

0.5

0.4

0.2

0.1

0 0 15 30 45 60 75 90 Time (min)

Fig. 3 Monosaccharide and cellobiose concentration during propylsulfonic acid-catalyzed hydrolysis at 160°C (• cellobiose, • glucose, A xylose, X arabinose) 103

0 15 30 45 60 75 90 Time (min) Fig. 4 Monosaccharide and cellobiose concentration during propylsulfonic acid-catalyzed hydrolysis after silica gel filtration at 175°C (• cellobiose, • glucose, A xylose, X arabinose) 104

1

0.9 0.8

.0.7 o0.6

0.5

o0.3 o 0.2

0.1

0 0 15 30 45 60 75 90 Time (minutes) Fig. 5 Monosaccharide and cellobiose concentration during propylsulfonic acid-catalyzed hydrolysis after silica gel filtration at 160°C (* cellobiose, • glucose, A xylose, X arabinose) 105

1 0.9 0.8 0.7

o 0.6 -ê—< CO 0.5 c

0 15 30 45 60 75 90 Time (min) Fig. 6 Monosaccharide and cellobiose concentration during hydrothermolysis at 175°C (• cellobiose, • glucose, A xylose, X arabinose) 106

100 90

80

70

60 50 40 30 20

10

0 0 15 30 45 60 75 90 Time (min) Fig. 7 Conversion to monosaccharide percentage during propylsulfonic-acid catalyzed hydrolysis at 175°C (• glucose, A xylose, X arabinose) 107

100

90

80

40

20

10

0 0 15 30 45 60 75 90 Time (min) Fig. 8 Conversion to monosaccharide percentage during propylsulfonic-acid catalyzed hydrolysis at 160°C (• glucose, A xylose, X arabinose) 108

CHAPTER 6

Summary and Future Investigations

6.1. Summary

The potential for lignocellulosic materials to provide transportation fuels is immense.

It has been conservatively estimated that 100 billion gallons of ethanol could be made annually from waste and energy crops [1]. The knowledge of this potential has lead to extensive research activity in this area, but not yet to a commercial process. The ability to economically hydrolyze the variety of glycosidic bonds present in these materials in high yield is the key to unleashing the energy stored in polysaccharides.

Biological systems have been developed to hydrolyze these materials, but suffer from low reaction rates [2-4], lack of robustness [3, 5, 6], high cost [3, 4, 6, 7], and difficulty dealing with the heterogeneous nature lignocellulose [2, 3, 5, 8, 9], There have been considerable improvements made in all of these areas, but the system has not yet reached commercial viability [5, 6].

Mineral acids have also been studied to accomplish this task [10-19]. The use of acids typically results in the formation of degradation products [10, 20, 21], The understanding of these reactions is not complete [19, 22, 23]. The complex interplay between acid concentration, temperature, and reaction time has been given only superficial treatment, stopping with correlations [10, 13, 24-26], The ability to tailor reaction conditions to provide maximum yields will require a fundamental understanding of both the hydrolysis reactions and the degradation reactions. 109

The preceding chapters have added to the knowledge of both the hydrolysis and degradation reactions that occur in the chemical catalyst system. The use of acid-

functionalized mesoporous silica as a catalyst for lignocellulosic hydrolysis is a new

approach that has led to improved understanding of silica materials, aqueous reaction

conditions, and carbohydrate chemistry. Evidence has been show that these systems offer similar performance to homogenous acid systems for disaccharide hydrolysis as well as the importance of the leveling affect of water and proton mobility. Data has been presented

indicating degradation reactions are slowed through the selection of optimum pH. This

knowledge can be added to current knowledge in related fields to develop an integrated

approach to improving lignocellulosic hydrolysis. The opinion held by many researchers is

that a combination of the chemical and biological treatment systems will likely be the best

approach to lignocelluloic hydrolysis [27-29],

Evidence that protons in solution are responsible for glycosidic bond hydrolysis

directs future research direction. Correlation of hydrolysis rates with hydrogen ion

concentration have shown that hydrogen ions are responsible for cellobiose hydrolysis

independent of the source [24, 30]. Studies on wood hydrolysis by Saeman [10] found that a

100% increase in sulfuric acid concentration resulted in a 153% increase in hydrolysis rate.

The high mobility of the proton in aqueous solution [31-33] suggests that surface character,

acid strength, anion character, and pore size are not important in this system because the

reaction occurs in the bulk of the solution away from the catalyst surface. The primary

characteristic of the catalyst material is the ability to deliver a high concentration of protons

to the bulk solution. The important secondary characteristics of the catalyst are long-term

stability in an aqueous matrix and ease of processing. 110

6.2. Future Investigation

Potential solid-acid-catalyst materials include mesoporous silica [30, 34], cation exchange resins [35-37], zeolites [37], and functionalized carbon [38, 39]. The following paragraphs will briefly discuss some of the advantages and disadvantages of each material.

Mesoporous silica has been used in a variety of reactions with a variety of functional groups [40]. Mesoporous silica has high surface areas, controllable pore sizes, ease of functionalization, and moderate hydrothermal stability. These materials are expensive to produce on a large scale and difficult to utilize in a commercial application due to their very small particle size. The ability to tune a variety of characteristics makes mesoporous silica an excellent material to investigate reaction systems. These characteristics are not important in aqueous systems because proton mobility[32] is so fast that the reaction does not occur in pores or near the surface.

Cation-exchange resins provide high acid concentrations, but suffer from low thermo­ stability [37]. Improvement in the thermo-stability of resins is needed to make them attractive catalysts for hydrolysis reactions.

Zeolites are typically considered inactive at low temperatures due to their hydrophilic nature[37]; however, high-silica zeolites, such as dealuminated mordenite and ZSM-5, are known to be hydrophobic. High-silica zeolites were shown to have moderate activity in the hydrolysis of ethyl acetate [41], and the effective acidity could be controlled by varying the

Si/Al atomic ratio.

Activated carbons can have extremely high surface areas (-1800 m2/g) relatively neutral surfaces, and well developed pore structure [38]. Little research has been done on the use of activated carbon supports to heterogenize homogeneous catalysts for liquid phase Ill processing [42]. Budarin et al. [43] reported the direct chemical modification of mesoporous activated carbon with functionalized silanes to hydrophobize the surface. The success in using this method suggests the use of acid-functionalized silica groups to chemically modify activated carbon materials. The resulting material could potentially deliver a high concentration of protons to solution on an inexpensive and easy to use support material. An alternative method of functionalization reported by Guerreiro et al. [39] is the direct anchoring of 3,6-dioxa-l,8-octanedithiol onto the activated carbon surface. The authors provided no detail, yet claimed their material had higher catalytic activity than a S03H-

MCM-41 material in the transesterification of ethyl benzoate. The Guerreiro et al. material may have advantages by avoiding the use of a silica group; however, more detail concerning their synthesis method is needed.

Achieving selectivity toward desired products and prevention of degradation of monosaccharides will be done by controlling characteristics of the bulk solution. These characteristics include selection of temperature, reaction time, solvent effects, and acid concentrations.

The lower activation energy of glucose degradation compared to cellobiose hydrolysis suggests that higher temperature reactions would favor desired products [30, 44].

The hydrolysis of xylobiose has a higher activation energy [45] than cellobiose, suggesting an even higher optimum temperature. Creative reactor designs, such as a shrinking-bed design, have been used to optimize reaction time [46] and show potential for further improvements. Torget, Kim, and Lee [46] demonstrated an initial hydrolysis rate five times faster with a flowing reactor compared to batch design. The use of small amounts of ethanol

(0.2 wt. %) in the reaction media have improved yields from yellow poplar hydrolysis from 112

55% to 85% [22], suggesting solvent manipulation can significantly improve yields. Acid concentrations have been investigated by several authors [11, 20, 25, 44, 47], showing an apparent pH optimum for glucose selectivity between near 2.2 [11,25].

The realization of commercially viable lignocellulosic hydrolysis will likely require the implementation and optimization of several technologies. The interest in this technology has risen and fallen with the price of petroleum over the last century. The importance of this technology is bigger than cheap energy. The payoff for society will be large in terms of energy independence, pollution reduction, and reinvigoration of rural economies.

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46. Torget, R.W., J.S. Kim, and Y.Y. Lee, Fundamental Aspects of Dilute Acid Hydrolysis/Fractionation Kinetics of Hardwood Carbohydrates. 1. Cellulose Hydrolysis Industrial Engineering Chemistry Research, 2000. 39: p. 2817-2825. 47. Smidsrod, O., A. Haug, and B. Larsen, The Influence ofpH on the Rate of Hydrolysis of Acidic Polysaccharides. Acta Chemica Scandinavica, 1966. 20: p. 1026-1034.