Bioproducts from sulfite pulping: Bioconversion of sugar streams from pulp, sludge, and spent sulfite liquor. Lisa X. Lai.

Transcription

1 Bioproducts from sulfite pulping: Bioconversion of sugar streams from pulp, sludge, and spent sulfite liquor Lisa X. Lai A thesis submitted in partial fulfillment of the requirements for the degree of Master of Science University of Washington 2010 Program Authorized to Offer Degree: School of Forest Resources

2 University of Washington Graduate School This is to certify that I have examined this copy of a master s thesis by Lisa X. Lai and have found that it is complete and satisfactory in all respects, and that any and all revisions required by the final examining committee have been made. Committee Members: Renata Bura Rick Gustafson William McKean Date:

3 TABLE OF CONTENTS Page List of figures... ii List of tables... iii Chapter 1: Introduction Lignocellulosic biomass Pretreatment Sulfite pulping Enzymatic hydrolysis Fermentation SHF vs. SSF Products from a sugar platform Chapter 2: Draft paper Introduction Methods Results and discussion Conclusion Chapter 3: Additional results and discussion Fermentation of SSL using S. cerevisiae Effect of solids loading on enzymatic hydrolysis Effect of enzyme loading on hydrolysis Chapter 4: Additional conclusions and future work Acknowledgements References i

4 LIST OF FIGURES Page 1. Bioconversion process schematic 2 2. Experimental design Spent sulfite liquor C. guilliermondii fermentation Pulp and sludge hydrolysis Fermentation of pulp hydrolysates Fermentation of sludge hydrolysates Combined feedstock SSF Spent sulfite liquor S. cerevisiae fermentation Variable consistency hydrolysis Variable enzyme loading hydrolysis..31 ii

5 LIST OF TABLES Page 1. Pulp and sludge composition Spent sulfite liquor composition Solid stream result summary..27 iii

6 Chapter 1: Introduction The global impact of human energy consumption has been made increasingly apparent in recent years by growing rates of atmospheric CO 2 accumulation, likely caused by a rise in anthropogenic burning of fossil fuels (Canadell, Le Quéré et al. 2007). It has been shown that mitigation of these effects may be achieved through displacement of petroleum-based transportation fuels with a biofuel producing less harmful emissions, such as ethanol (Tyson 1993; von Sivers and Zacchi 1995; Bergeron 1996; Galbe and Zacchi 2002). In addition to environmental benefits, ethanol addresses concerns about the exhaustion of available fossil fuels, energy security, and ease of adaptation to the current fuel infrastructure in the US. Though ethanol is overwhelmingly produced from sugar cane in Brazil, and corn in North America, lignocellulosic biomass, such as agricultural and wood residue, is a widely available and largely untapped feedstock (Wiselogel, Tyson et al. 1996). In addition, ethanol production from lignocellulosics, commonly called bioethanol, has been shown to have a more favorable energy ratio than that originating from starch or sugar sources (Keeney and DeLuca 1992; Lorenz, Morris et al. 1995; Gnansounou and Dauriat 2005). Major process steps for the traditional bioconversion scheme of lignocellulosic biomass to bioethanol are well known and illustrated in Figure 1. First, lignocellulosic biomass is pretreated to break up its rigid structure and prepare it for the next step, hydrolysis. During enzymatic hydrolysis, sugars are released from the pretreated biomass. Fermentation utilizes microorganisms to convert these sugars to bioproducts, for example ethanol and xylitol. Though laboratory-scale bioethanol production has been increasingly studied in recent years, no fullscale commercial plants exist in the US today. One reason is that challenges exist in each step of the bioconversion scheme, which will be discussed in greater detail in subsequent chapters. First, an ideal biomass is difficult to find, as it must be inexpensive, readily-available in large quantities, and consistent in chemical composition. Pretreatment requires optimization of severity, as the ideal degree of biomass fractionation is often limited by the formation of fermentation inhibitors. Hydrolysis necessitates consideration for the cost of enzymes, potentially requiring sacrifices in sugar yield. Finally, fermentation requires selection of an ideal microorganism for the specific operation at hand, making considerations for inhibitor tolerance and possible fermentation of co-products alongside of bioethanol. 1

7 Sugar streams from sulfite pulping, described in greater detail later, were studied in this thesis as a means of mitigating the problems associated with the biomass acquisition and pretreatment steps described above. Spent sulfite liquor (SSL), pulp, and sludge, the streams studied in this thesis, are concurrently produced with bleached sulfite pulp at a rapid rate, thus meeting the criteria of being inexpensive and available in large quantities. In addition, due to the majority of fractionation and delignification already being accomplished by the sulfite pulping process, the streams studied here do not require pretreatment. Lignocellulosic biomass Pretreatment Hydrolysis Fermentation Bioproducts (ethanol, xylitol, etc.) Objective and Outline Figure 1. Simplified process schematic for the conversion of lignocellulosic biomass to bioproducts. This process flow illustrates the separate hydrolysis and fermentation (SHF) process, however hydrolysis and fermentation can alternatively be combined into one step, or a process known as simultaneous saccharification and fermentation (SSF). The main objective of the work presented in this thesis is to examine the potential of producing single and mixed sugars from sulfite pulping streams to use for the production of bioethanol and other valuable bioproducts. This was accomplished first through the fermentation of SSL with Candida guilliermondii, a microorganism capable of fermentation mixed 5-and 6-carbon sugars. Next, single sugars streams were produced from the separate hydrolysis and fermentation (SHF) of pulp and sludge in water. Finally, mixed sugars streams were produced from SHF and simultaneous saccharification and fermentation (SSF) of pulp and sludge mixed with SSL. Success of this process would create the potential to exploit a new feedstock for bioethanol conversion, as well as address some of the challenges associated with bioconversion described above. Several background sections are presented in order to put this research into context. Section 1.1 provides a background about biomass chemistry. Section 1.2 discusses pretreatment methods and 2

8 limitations. Section 1.3 discusses sulfite pulping as a means of pretreatment. Section 1.4 describes enzymatic hydrolysis. Section 1.5 provides an overview of fermentation in terms of the microorganisms used in this study and their associated products. Section 1.6 compares separate hydrolysis and fermentation (SHF) and simultaneous saccharification and fermentation (SSF) bioconversion processes. Finally, Section 1.7 contains an overview of the most promising biochemical building blocks that can be made from biomass. Chapter 2 contains a draft paper that will be submitted for publication regarding the use of sulfite pulping streams as a sugar platform from which to produce higher-value bioproducts. Additional results and discussion not presented in Paper I are shown in Chapter 3, followed by additional conclusions and future work in Chapter Lignocellulosic biomass Feedstocks for ethanol production can be categorized into sucrose-, starch-, and lignocellulosic-based materials. Sucrose-based materials are readily fermentable to ethanol and other products, while starch-based feedstocks can easily be converted to sugars via enzymatic hydrolysis. Lignocellulosic feedstocks represent great production value, but due to their chemistry, are the most challenging to utilize for bioconversion. Lignocellulosics encompass a broad range of materials, including woody biomass, herbaceous plants (switchgrass and giant reed), and agricultural residues (corn stover and wheat straw). Common among all lignocellulosic biomass types are their three major components of cellulose, hemicellulose, and lignin, and minor components of extractives and ash. These constituent compounds are present in varying quantities among biomass types, depending on their species of origin. Cellulose Lignocellulosic feedstocks are derived from plant biomass, whose defining characteristic at the cellular level is the presence of cell walls comprised primarily of cellulose. The most abundant natural polysaccharide, cellulose is the target of bioethanol conversion processes. Cellulose is comprised of D-glucose units linked by β-1,4 glycosidic bonds. A dimer of two linked glucose units is referred to as cellobiose. It is because of this linkage that each individual cellulose molecule forms a linear polymer. Bioconversion necessitates the release of these individual glucose units for microbial fermentation to bioproducts. The degree of polymerization (DP) of cellulose, or number of monomeric units in an oligomeric molecule, varies by source. Sulfite pulp, one of the feedstocks used in this study, averages a DP of 1255, while softwood tree species, the primary source of the sulfite pulp used in this study, average a DP of 8000 (Fengel and Wegener 1989). Cellulose 3

9 molecules with a DP less than 8 are considered water-soluble, while at higher DPs, they have a greater affinity for one another than for water (Brown 2004). In native cellulose, individual molecules form rigid microfibrils in which they are aligned with reducing ends oriented in the same direction. This structure is stabilized by rigidly arranged intra- and intermolecular hydrogen bonds, thus giving native cellulose a highly crystalline conformation that is difficult to degrade (Klemm, Heublein et al. 2005). Amorphous (non-crystalline) regions have also been found to comprise 10-50% of native cellulose (Fan, Lee et al. 1982), but the influence of crystallinity on cellulose degradability is still being investigated (Atalla and VanderHart 1999; Jarvis 2003; Ding and Himmel 2006). The rigid structure of cellulose is resistant to chemical and mechanical alteration, thus posing a challenge for the bioconversion process (Ding and Himmel 2008). Hemicellulose Similar to cellulose, hemicellulose is also a polysaccharide, but its constituent sugars are both hexoses (6-C sugars; D-glucose, D-mannose, and D-galactose) and pentoses (5-C sugars; D-xylose and L- arabinose). With a few exceptions, pentoses are generally not readily fermentable by naturally occurring yeasts. Hemicellulose has a much lower DP, and therefore lower molecular weight, than cellulose, and is often a branched, rather than linear polymer. Hemicellulose composition and DP varies widely across species. Softwood hemicellulose is majorly comprised of galactoglucomannans, composed of a backbone of β-1,4 linked mannose and glucose units in roughly a 3:1 ratio. Acetyl groups occur on the C2 or C3 carbon of roughly every third backbone unit. The glucomannan backbone is branched with α-1,6 linked galactose side chains occurring on glucose backbone units. Softwood galactoglucomannans constitute about 20% of dry wood by mass, and ranges from in DP (Fengel and Wegener 1989). Softwoods also contain 10-15% xylans, composed of a β-1,2 linked xylose backbone. One in 10 xylose units has a branched α-1,4 linked arabinose unit, and one in five has a α-1,2 linked 4-O-methylglucuronic acid unit (Fengel and Wegener 1989). In contrast, hardwood hemicellulose is mainly comprised of xylans (15-30%) characterized by a β-1,4 linked xylose backbone, where many of its xylose units are acetylated at their C2 or C3 carbons. Side chains of α -1,2 linked 4-O-methylglucuronic acid units occur at every 6-11 xylose units. Hardwood xylan has a DP of Mannans occur in only 3-5% of hardwood by mass, and are characterized by β -1,4 linked backbone of mannose and glucose units in roughly a 2:1 ratio with no side chains. Native hemicellulose is embedded within the cell wall, binding to cellulose and components, such as structural proteins. The heterogeneity of hemicellulose makes it non-crystalline, and easier to hydrolyze. 4

10 Lignin The second most abundant organic substance within plant biomass is lignin, a highly complex polymer consisting of phenolic compounds. Lignin composition again varies greatly between species, but its three constituent phenolic ring structures are ρ-hydroxyphenyl, guaiacyl (containing one methoxyl group), and syringyl (containing two methoxyl groups), which are bound to one another through a complex network of carbon-carbon and ether bonds. Softwood lignin is comprised overwhelmingly by guaiacyl units. Common lignin linkages include β-o-4, α-o-4, 4-O-5, 5-5, β-β, β-5, and β-1, all of which, with the exception of α-o-4, are formed through free radical coupling of precursor ring structures (Fengel and Wegener 1989). Roughly 50% of softwood lignin is β-o-4 linked, a reactive linkage under alkaline pulping conditions. The next most common linkages for softwood lignin are β-5, comprising 9-12%, and 5-5, comprising 10-11%. All other linkages each comprise less than 8% of softwood lignin (Fengel and Wegener 1989). Like glue, lignin binds the cell wall components together, giving lignocellulosic biomass its structural integrity, while also contributing to its flexibility (Pan 2008). The presence of lignin poses one of the biggest challenges to bioconversion, as its removal is necessary in order to allow hydrolysis enzymes access to cellulose and its constituent sugars (Akin 2007; Pan 2008). Extractives and ash Though present in very small quantities, typically below 5%, organic extractives can still greatly affect certain properties of biomass, including its color, odor, and density (Kai 1991). In nature, extractives perform the functions of protection from diseases and parasites, attraction of pollinators and seed dispersers, and food storage. Softwood extractives consist mainly of terpenes, fats, waxes, and phenolics (Fengel and Wegener 1989). Ash is composed of the inorganic compounds remaining after complete combustion of biomass. Though comprising only about 1% of wood biomass, ash is composed of Ca, K, and Na oxides that are essential for plant growth (Ohlsson 2004). Ideal feedstock Though lignocellulosic bioconversion has been increasingly studied in recent years, an ideal feedstock has yet to be found. Pretreatment and other steps in the bioconversion process must be optimized in a manner that is dependent on feedstock composition, so the wide compositional variability in biomass is problematic. Also, a desirable feedstock should be high in cellulose content, while low in lignin content to maximize the ease and efficiency of bioconversion. Ideal feedstocks 5

11 must also be free of non-native contaminants that may interfere with hydrolysis or fermentation, rendering many industrial residues (e.g. wood and paper waste) undesirable. There are also economic and social considerations. Biomass can account for 25-40% of the cost of bioethanol production (Nguyen and Saddler 1991; Gregg and Saddler 1996; Galbe and Zacchi 2002; Wingren, Galbe et al. 2003). An inexpensive feedstock with year-round availability is ideal, making, for instance, agricultural grasses less desirable due to their high cultivation costs and seasonallydependent growth rates. Also, the acquisition of an ideal biomass should not be in competition with other industries, such as agriculture or wood products manufacturing. This topic has become more relevant with the advent of recent studies regarding the downfalls of corn ethanol (Pimentel 2003). An ideal feedstock exhibiting all of these characteristics has proven to be difficult to find. 1.2 Pretreatment As previously described, the crystallinity of cellulose, its particle size, available surface area, and the presence of lignin make raw lignocellulosic biomass resistant to direct saccharification by hydrolysis enzymes. Therefore, a pretreatment step is needed in order to alter biomass rigid structure (Figure 1). During pretreatment, the surface area of biomass is increased, particle size decreased, and, to some degree, the crystallinity of cellulose is decreased (Mosier, Wyman et al. 2005). The cell wall matrix is disrupted, accompanied by partial removal of lignin and hemicellulose. Mechanical, chemical, and biological methods, or any combination of these, are utilized in pretreatment. Mechanical methods involve milling or grinding, often in combination with a thermal treatment such a steaming. Chemical pretreatment involves the addition of acid or alkali solvents that result in partial degradation of lignin or hemicellulose. Biological pretreatment subjects the biomass to lignindegrading microorganisms. Steam explosion is a method that has been found to be very efficient for woody feedstocks. Biomass is subjected to high temperature ( C) and pressure ( psi) for a period of 10 s to 10 m, followed by a rapid release of pressure, which is said to cause an explosion of biomass into its fractionated form (Mason 1933; DeLong 1977). Optimal conditions for temperature, pressure, and time are dependent on feedstock composition. The addition of an acid catalyst such as H 2 SO 4 or SO 2 to raw biomass before steam explosion has been found to improve pretreatment efficiency (Ramos, Breuil et al. 1992; Eklund, Galbe et al. 1995). 6

12 Ammonia fiber expansion (AFEX) pretreatment is a similar process to steam explosion, in which biomass is combined with ammonia at psi and C before a rapid release of pressure. Yields of up to 90% of the theoretical ethanol yield have been observed using AFEX pretreatment in switchgrass (Alizadeh, Teymouri et al. 2005), however, while effective on agricultural residues, this method has not been found to work with softwoods (Hsu 1996). Organosolv pulping has emerged as a pretreatment method for ethanol production. In this process, biomass is combined with ethanol, water, and H 2 SO 4, and cooked under high temperature and pressure. The resultant slurry is filtered, and solids are hydrolyzed, while lignin is precipitated from the filtrate. The residual liquid fraction, containing hydrolyzed sugars, can be concentrated and also fermented (Lora and Aziz 1985). Biological pretreatment is carried out through the delignification of biomass by white rot fungi, mostly belonging to the phylum Basidiomycotina (Blanchette 1991). Lignin degradation by these microorganisms occurs via the enzyme activity of lignin peroxidase, manganese peroxidase, and laccase, though some cellulose degradation is frequently observed due to non-specific enzyme binding (Pointing, Pelling et al. 2005). The degree of preferential lignin consumption can vary widely between organisms. For example, cellulose loss of 65% has been observed for Trametes versicolor, while for Perenniporia medulla-panis, 73% delignification has been achieved without any loss of cellulose (Blanchette 1991). Due to a slow reaction rate, the largest drawback of solely using biological pretreatment is the long residence time that is required, up to 60 days, relative to that of steam explosion (Taniguchi, Suzuki et al. 2005; Yu, Zhang et al. 2009). Microbial delignification has more prominently been studied as a pretreatment method to use in conjunction with organosolv pulping, where it has been found to decrease energy consumption by up to 68% (Blanchette 1991). The severity factor, or the degree to which biomass is fractionated during pretreatment, is dependent on conditions such as temperature and residence time, and can be represented by the following formula: Log R O = t * e (T r-t b )/14.75 Where t = residence time (m), T r = reaction temperature, and T b = reference temperature (100 C) (Abatzoglou, Chornet et al. 1992). Too low of a severity can lead to incomplete fractionation of biomass, causing low hydrolysis yields in the next step. Higher severity pretreatments result in more complete delignification and a higher degree of polysaccharide hydrolysis, which are desirable 7

13 reactions. However, unfavorable reactions also occur during pretreatment. Hexose sugars are degraded to hydroxymethylfurfural (HMF), and pentoses to furfural. Further degradation of HMF leads to the formation of formic and levulinic acids (Fengel and Wegener 1989). Acetic acid is produced through the cleavage of acetyl groups in hemicellulose. These compounds are inhibitory to fermentation, and, at high enough concentrations, toxic to yeast (Delgenes, Moletta et al. 1996). Therefore, pretreatment optimization is essential to achieve efficient bioconversion. In addition, pretreatment can account for a large portion of the energy and/or chemical input needed in any bioconversion process. For this reason, and the need for optimization described above, pretreatment is often seen as the limiting step of bioconversion. 1.3 Sulfite pulping The feedstocks utilized here for bioconversion were sugar streams produced from ammonia-based acid sulfite pulping. This process involves the mixing of raw, mainly softwood chips with SO 2 at high temperature and acidic ph. Pulping occurs through the interaction of lignin with SO 2, whereby lignin is solubilized through the addition of hydrophillic sulfonate groups. Sulfonation mainly occurs on the α, and sometime γ carbon of lignin groups (Gellerstedt 1973). In addition, carbohydrates are cleaved from lignin-carbohydrate-lignin linkages, reducing lignin s molecular weight. In hemicellulose, acetyl groups are cleaved, as well as α-1, 6 galactosidic linkages (Glennie 1971). Therefore, the resulting spent sulfite liquor (SSL), as it is commonly referred, is high in monomeric sugar and sulfonated lignin (lignosulfonate) content. The resulting pulp is mostly delignified, and high in glucose content. Sludge, which is made up of pulping fines and rejects, is similar to pulp in sugar composition, but contains a high amount of residual lignin. These three streams were obtained for this study from collaborators at Kimberly-Clark in Everett, WA. Utilization of these as feedstocks for bioconversion makes pretreatment obsolete, as a high degree of solids delignification can be expected from the sulfite cooking process, and sugars in the liquid stream are expected to be mostly in monomeric form. Also, sulfite pulping provides the benefit of a central location where sugar streams are already being generated at no additional cost. 1.4 Enzymatic hydrolysis Saccharification of cellulose into its monomeric glucose components is achieved by the action of a group of enzymes collectively referred to as cellulases. These enzymes are synthesized by various naturally-occurring bacteria and fungi, the most prominent of these being Trichoderma reesei, an 8

14 aerobic mesophilic fungus. Its cellulase enzymes are classified into three groups: endoglucanases (EG), cellobiohydrolases (CBH), and β-glucosidase (βg), each associated with a specific action (Ghose 1987). EG attacks internal bonds within cellulose, exposing free ends and disrupting its crystalline structure. Starting from these exposed free ends, CBH travels along the length of the cellulose chain, cleaving cellobiose units, which are subsequently hydrolyzed into two glucose units by βg. One explanation for this system of synergistic enzymatic activity is limitation of end-product inhibition (Eriksson, Karlsson et al. 2002; Väljamäe, Kipper et al. 2003). T. reesei is cultured industrially and its enzymes are separated and purified for use in large-scale hydrolysis. Two user-controlled factors affecting hydrolysis rate are enzyme loading and solids consistency. Since enzyme quantity cannot be directly measured, enzyme activity for EG and CBH is measured in terms of filter paper units (FPU) per volume. The quantity of enzyme activity required to produce 2.0 mg of cellobiose from 50 mg of filter paper (essentially pure cellulose), or a 4% conversion, after 1 hr at 50 C and 4.8 ph (the ideal conditions for cellulase activity) is defined as FPU (Decker, Adney et al. 2003). The effect of enzyme loading on glucose yield is not linear. For example, a 50% increase in enzyme loading would result in less than a 50% increase in glucose yield (Ghose 1987). Enzyme loading mainly affects the initial rate of hydrolysis, and therefore, the residence time required to reach maximum glucose yield. Therefore, a change in enzyme loading may not have a proportionate effect on final glucose yield itself. Even at very high enzyme loading, the complete hydrolysis of cellulose is often difficult to achieve due to the presence of residual lignin, and endproduct inhibition (Tengborg, Galbe et al. 2001). Both cellobiose and glucose act as noncompetitive inhibitors of T. reesei cellulases (Holtzapple, Cognata et al. 1990; Xiao, Zhang et al. 2004). Ethanol is also inhibitory, but to a far lesser extent (Holtzapple, Cognata et al. 1990). Accumulation of these compounds during hydrolysis is disadvantageous. Glucose has been found to be less inhibitory than cellobiose, so a proportionately higher loading of βg is typically used during enzymatic hydrolysis to limit inhibition, compared to EG or CBH. Enzyme cost occupies a significant portion of the total cost for bioconversion, so minimizing enzyme loading to the extent that it is not detrimental to glucose yield is favorable (Shen and Agblevor 2008). Solids consistency is described as dry weight of biomass (substrate) divided by the volume of total liquid in a hydrolysis reaction, expressed as a percentage. Enzyme kinetic curves for cellulose hydrolysis typically exhibit a biphasal shape, with an initial logarithmic phase followed by an asymptotic phase as maximum glucose conversion is approached (Ramos, Breuil et al. 1993). When expressed in terms of percent glucose conversion from cellulose, increasing solids consistency is 9

15 typically found to reduce the initial conversion rate. Glucose percent conversion is typically found to decrease, but final glucose concentration may improve due to greater substrate availability (Cara, Moya et al. 2007). High consistency hydrolysis represents a promising source of cost reduction. A previous assessment has shown that total cost can be reduced by nearly 20% when solids consistency is increased from 5% to 8% for softwood (Wingren, Galbe et al. 2003). However, it has been supposed that bioconversion using solids consistencies of greater than 8% encounters three key barriers. First, there is a reduction in the amount of free water available for enzyme transport due to hydrogen bonding with the released sugars (Felby, Thygesen et al. 2008). Secondly, higher consistencies can introduce a high concentration of lignin, a fermentation inhibitor (Delgenes, Moletta et al. 1996). Finally, increased sugar production can cause severe end-product inhibition (Holtzapple, Cognata et al. 1990). Solids loading of up to 40% has been reported in wheat straw, resulting in a decrease in percent glucose conversion to one third that of 2% consistency hydrolysis (Jørgensen, Vibe-Pedersen et al. 2007). However, a more promising result has been published for hardwoods, where 20% consistency hydrolysis resulted in 158 g/l glucose (Zhang, Qin et al. 2009). Low enzyme loading and high consistency hydrolysis represent great potential in terms of cost reduction for bioconversion technologies. 1.5 Fermentation Microbial fermenters include a variety of bacteria and fungi. The well-studied Saccharomyces cerevisiae, known commonly as baker s or brewer s yeast, is one such organism utilized in this report. Originally isolated from the skin of grapes, it has become a model eukaryotic organism in biological research due to its size, ease of genetic manipulation, and high economic value. Like many organisms, S. cerevisiae metabolizes glucose, and other 6-carbon sugars, via the Embden-Meyerhof- Parnas glycolytic pathway, mainly producing ethanol. Genetically modified S. cerevisiae such as 424A (LNH-ST) can also be made to metabolize 5-carbon sugars (Sedlak and Ho 2004). The overall reaction of hexose fermentation to ethanol is: C 6 H 12 O 6 2 C 2 H 5 OH + 2 CO 2 The mass balance of this process indicates that the maximum possible yield of ethanol is 51% of the mass of starting glucose. The strain of S. cerevisiae utilized in this study (ATCC 96581) was isolated from spent sulfite liquor, and had previously demonstrated tolerance of 8 g/l of acetic acid with superior galactose fermentation to a commercially available S. cerevisiae (Linden, Peetre et al. 1992). Yields of about 75% of the theoretical maximum ethanol were achieved for SSL at ph 6, but other S. 10

16 cerevisiae strains have shown near 100% yield from synthetic sugars (Keating, Panganiban et al. 2006). Similar to other ethanologenic yeasts, the strain exhibits preferential consumption of glucose first, followed by mannose, then galactose (Linden, Peetre et al. 1992). A second organism, Candida guilliermondii, (ATCC ) was also utilized in this study to demonstrate cofermentation of xylitol alongside of ethanol. Xylitol is primarily used as a sweetener, and due to being sugar alcohol, does not impact insulin levels when ingested, and contains 36% fewer calories than sucrose (Hassinger, Sauer et al. 1981). C. guilliermondii converts xylose to xylitol via the Xylose Reductase-Xylitol Dehydrogenase (XR-XDH) pathway, represented by the net equation: 60 C 5 H 10 O ADP + 12 P i + 12 H 2 O + 3 O 2 54 C 5 H 12 O ATP + 30 CO 2 The theoretical yield of xylitol is 91% of the starting xylose mass (Barbosa, de Medeiros et al. 1988). C. guilliermondii also concurrently ferments hexoses to ethanol. Because of this unique ability, the composition of media used to propagate the yeast prior to fermentation has an effect on its productivity, an phenomenon that has been explained by enzyme induction (Lee, Sopher et al. 1996). C. guilliermondii pre-grown in xylose media has previously been found to produce a significantly higher xylitol yield than yeast grown in glucose alone or in mixed media. By pre-growing yeast on xylose, yields of approximately 60% of the theoretical maximum for xylitol were found during fermentation of hydrolysate containing both glucose and xylose (da Silva and de Almeida Felipe 2006). This yield has been found to further improve when xylose is the only sugar in the fermentation media, to about 80% (Barbosa, de Medeiros et al. 1988). Fermentation inhibitors released from biomass during pretreatment include of a variety of compounds that are categorized into three groups. Weak acids (acetic acid) and furan derivatives (furfural and HMF) are sugar degradation products, while phenolic compounds result from the degradation of lignin during pretreatment. Inhibitors have been topic of great interest, but their mechanisms of inhibition are still a matter of investigation (Palmqvist and Hahn-Hägerdal 2000). Though seemingly contradictory, it has been reported by several studies that small quantities of inhibitors can enhance fermentation yields (Banerjee, Bhatnagar et al. 1981; Sanchez and Bautista 1988; Pampulha and Loureiro-Dias 1990; Taherzadeh, Niklasson et al. 1997). In particular, ethanologenic yeasts have been reported to tolerate up to 10 g/l acetic acid, 3 g/l HMF, and 1.6 g/l furfural with no significant effect on ethanol yield, and even improved yields in some cases (Keating, Panganiban et al. 2006). 11

17 Fermentation products, such as ethanol (Brown, Oliver et al. 1981), xylitol (Meyrial, Delgenes et al. 1991), lactic acid (Ohara, Hiyama et al. 1992), and glycerol (Zeng, Ross et al. 1994), can furthermore act as inhibitors by limiting yeast growth and product formation. Metals released from reactors and equipment during industrial bioconversion processes have also been found to be inhibitory (Oleszkiewicz and Sharma 1990). Due to the variation in their fermentation products and inhibitor tolerance, different microorganisms can yield dissimilar results under the same conditions. Therefore, the appropriate microorganisms must be selected in each individual case of fermentation, with regard to the sugars and inhibitors that are present, as well as desired product(s). 1.6 SHF vs. SSF In SHF, hydrolysis and fermentation are carried out as completely separate steps, whereby enzymes are added and saccharification is executed to completion, only after which, yeasts are added. The main advantage of this process is it allows for each step to be performed at its optimum conditions for temperature and ph. As previously mentioned, cellulase enzymes are maximally active at 50 C and a ph of 4.8, whereas most microbes ferment better at temperatures near 30 C and ph near 6.0. However, a major disadvantage of SHF is that the accumulation of glucose and cellobiose during hydrolysis can lead to end-product inhibition, as described earlier. To maintain acceptable ethanol yields, the hydrolysis step in SHF must often be carried out at low solid loadings, resulting in a relatively dilute ethanol stream. Economically, this is not ideal, as it can increase costs for downstream processing, namely for distillation (Wingren, Galbe et al. 2003). In SSF, hydrolysis and fermentation take place concurrently, thereby reducing the possibility of endproduct inhibition. As described previously, ethanol is far less inhibitory to cellulases than are the sugars released during hydrolysis, 1/16 as inhibitory as cellobiose according to one study (Holtzapple, Cognata et al. 1990). However, the main disadvantage of SSF is that it takes place under compromised conditions of temperature (around 37 C) and ph (around 5.5), which can have an effect on total yield. Generally, studies comparing SSF and SHF have illustrated that determination of the better method is dependent on a number of factors. Several studies have confirmed that SHF produces higher overall yields, while SSF requires less time (Alfani, Gallifuoco et al. 2000; Cantarella, Cantarella et al. 2004). An ideal operating process would be flexible to allow for either method to be used in accordance to feedstock availability and the desired product(s). 12

18 1.7 Products from a sugar platform A number of building blocks for high value bioproducts can be made using the platform of microbial conversion from sugars. Of the 12 most promising of these reported by the U.S. Department of Energy, seven can currently be produced using known biological pathways with microorganisms. The remaining five are presently produced using chemical pathways (Werpy, Petersen et al. 2004). Four carbon 1,4-diacids, including succinic fumaric, and malic acids, are synthesized through microbial fermentation via overexpression of Krebs Cycle pathways associated with C4 diacid formation, primarily with Escherichia coli (Millard, Chao et al.). Chemical reduction of these building blocks produces derivatives that can be used to make solvents, water soluble polymers, and fibers such as lycra. 3-Hydroxypropionic acid (3-HPA) can be produced via microbial fermentation, though the pathway is not known, nor is its chemical conversion pathway. Sonora fiber can be derived from 1,3-propane diol, the product of 3-HPA reduction. Acrylates are formed by 3-HPA dehydration, which are used to make super absorbent polymers. Aspartic acid is produced microbially through fermentation or enzymatic action on oxaloacetate in the Krebs cycle, or chemically from the animation of fumaric acid. Amine butanediol can be produced through its chemical reduction, aspartic anhydride through its dehydration, or polyaspartic from its polymerization. Glutamic acid is made through microbial fermentation of glucose, chiefly by Corynebacterium glutamicum (Georgi, Rittmann et al. 2005). Reduction of glutamic acid produces diol, diacid, and aminodiol derivatives, which can be made into monomers for polyesters and polyamides. Itaconic acid can be produced through fermentation by aerobic fungi, and is primarily used as a copolymer and polymer precursor, namely for acrylic or methacrylic acid (Willke and Vorlop 2001). Glycerol is produced through the transesterification of oils or from anaerobic microbial fermentation of sugars (Yazdani and Gonzalez 2007). It can be oxidized to generate glyceric acid, or directly polymerized to produce polyesters and polyols, utilized in polyurethane resins. Propylene glycol and 1,3-propanediol can also be produced from glycerol through its hydrogenolysis. Sugar alcohols, including xylitol and arabinitol, are made through microbial fermentation of xylose and arabinose, respectively, though its current primary production through the 13

19 chemical pathway of hydrogenation of those sugars. These building blocks may be used directly as sweetners, or oxidized to produce sugar acids (Prakasham, Rao et al. 2009). Though not produced microbially, levulinic acid can be generated from the degradation of HMF during pretreatment. It is currently produced through the chemical pathway of acid catalyzed dehydration of hexose sugars. Levulinic acid can be reduced to make derivatives for fuel oxygenates and solvents, oxidized to acetyl acrylates or acetic-acrylic succinic acids, used as copolymers. In addition, condensation of levilinic acid produced diphenolic acid, used in polycarbonate synthesis. This thesis presents xylitol and ethanol fermentation as examples of biochemical production, but a wide variety of bioproducts can potentially be made from the sugars available in sulfite pulping streams. Chapter 2: Draft paper Biochemical production from sulfite pulping sugar streams Abstract The production of single- and mixed-sugar streams and their conversion to bioproducts were studied using sulfite pulping streams as feedstocks. Sulfite pulp, sludge, and spent sulfite liquor (SSL) were utilized because they are concurrently generated alongside of bleached pulp, and because the pulping process renders pretreatment unnecessary. SSL, comprised of mostly monomeric hexose and pentose sugars, was directly fermented to ethanol and xylitol with Candida guilliermondii. Single-sugar streams were generated through hydrolysis of pulp and sludge in buffered water, followed by fermentation to ethanol with Saccharomyces cerevisiae. Mixed-sugar streams were generated through both SHF and SSF of pulp and sludge in SSL using S. cerevisiae. Direct fermentation of SSL to ethanol and xylitol produced yields consistent with that of a synthetic sugar control (89.5%, 40.3%, respectively). The best utilization of pulp was determined to be as a single-sugar stream, derived from SHF in water, which yielded a cellulose to ethanol conversion of 62.2% of the theoretical maximum (28.3 g/l). Sludge produced the highest yield when mixed with SSL during SSF (50.0%, 23.7 g/l). Sulfite pulp, sludge, and SSL currently represent untapped industrial resources for the production of single- and mixed-sugar streams from which high-value bioproducts can be made. 14

20 Introduction The biochemical industry currently lacks an abundant sugar source from which to make higher value products. An ideal sugar source must be inexpensive, readily available year-round, and relatively pure in composition. Using biological conversion, these sugars can be fermented to valuable products such as ethanol, xylitol, arabitol, succinic acid, and lactic acid (Clark and Deswarte 2008). Lignocellulosic biomass is a widely abundant sugar source, but requires expensive and energyintensive pretreatment, and rarely is consistent in composition. Large scale biomass-to-bioproduct plants do not currently exist in the US. Coupling biochemical production with an existing industry, such as sulfite pulping, creates the potential to use sugar streams already being generated by the pulping process, and simultaneously eliminates the need for pretreatment, as the resultant streams are mostly delignified. We explored the potential of converting sulfite pulping streams to sugar sources from which higher value bioproducts can be made. The three sulfite pulping streams examined in this study were pulp, primary clarifier sludge, and spent sulfite liquor (SSL). The hydrolysability and fermentability to ethanol of Kraft mill sludges has already been demonstrated by Sjöde et al and Kang et al The fermentability of SSL to ethanol using Rhizopus oryzae and Saccharomyces cerevisiae, respectively, has been exemplified by Taherzadeh et al. 1997, and Helle et al However, aside from ethanol, these materials can alternatively be converted to sugar streams, from which a host of biochemical can be produced using the model of a sugar platform (Clark and Deswarte 2008). Without bioconversion, sludge and SSL can require costly treatment techniques prior to disposal or burning, so their conversion to higher value products is both desirable and economically beneficial. Pulp is itself a highly valued product, so its conversion to biochemicals is less economically desirable. However, conversion to a very high value bioproduct may be desirable in the future if there is an excess of sulfite pulp on the market. In addition, reconfiguration of the traditional bioconversion scheme using these three pulping streams has not been well studied. The objective of this paper is to examine the potential of generating single and mixed sugars streams from sulfite pulp, sludge, and SSL for bioproduct conversion via SHF and SSF. We explored high consistency, low enzyme loading SHF of each stream separately, but we also analyzed SHF and SSF of combined feedstocks, SSL fortified with either of the two solid streams. This offers the potential to produce highly concentrated sugar streams without using large quantities of enzyme, as would be the case in traditional, single-feedstock enzymatic hydrolysis. Ethanol and 15

21 xylitol were produced to demonstrate examples of biological conversion, but many other bioproducts are possible. Methods Raw materials Ammonia-based sulfite pulping at Kimberly-Clark in Everett, WA produces woodpulp for direct sale or conversion to tissue products. Byproducts include primary clarifier sludge, and spent sulfite liquor. Pulp is produced at a nominal rate of rate of 500 air dry metric tons per day (admt/d). Under the right conditions, up to 90 admt/d could be provided for hydrolysis (Sande 2010). Current practice at Kimberly-Clark is to dry, press, and store pulp when it is not being used for paper production. Sludge is produced at a rate of 45 dry short tons/day, and SSL at 500 short tons dry solids/admt pulp at 14% solids (Sande 2010). Sludge is dewatered and burned as hog fuel onsite, and SSL is evaporated and burned to recover SO 2 and heat. Collaborators at Kimberly-Clark provided us with the pulp, sludge and SSL used in this study. All materials were derived from primarily softwoods. The mill also produces a hardwood grade, so pulp and sludge contained a small, unknown amount of hardwood fiber. Pulp was collected from the mill s pre-bleach washers and had not been treated with ClO 2. Sludge was collected from primary clarifiers before introduction to aerobic bacteria, and contained a mixture of pulping fines and rejects, tissue mill sludge, and boiler house effluent. SSL was taken directly from brown stock washers at 14% solids and had not been evaporated. Solids were washed with ten times their mass in water, and stored at -20 C until use. Moisture content was 77.3% for pulp, and 75% for sludge. SSL was stored at 4 C. Compositional analysis Insoluble carbohydrates and lignin TAPPI method T-222 om-98 (TAPPI 1998) was used to gravimetrically analyze insoluble lignin, and photometrically analyze soluble lignin. Carbohydrate content was measured using HPLC. Dried samples of 0.2 g were ground to 40-mesh size and combined with 3.0 ml of 72% (w/w) H 2 SO 4 for 2 h. Samples were then diluted to 4% (w/w) H 2 SO 4, autocalved at 121 C for 1 h, and filtered through glass fritted crucibles. Filtrate was collected, carbohydrate content analyzed by HPLC, and acid- 16

22 Fermentation SSF Fermentation Hydrolysis insoluble lignin content calculated by measuring UV at 205 nm. Oven-dried crucibles were weighed to determine acid insoluble lignin content. Soluble carbohydrates Soluble monomeric and oligomeric carbohydrate content was determined using NREL LAP TP (Sluiter, Hames et al. 2004). Five ml SSL was added to ml of 72% (w/w) H 2 SO 4 and filled to a 20 ml total volume with water. Samples were autoclaved at 121 C for 1 h and analyzed by HPLC to determine total sugar content. Monomeric sugars were analyzed on the raw SSL, and oligomeric sugar was calculated as the difference between total and monomeric sugar content. Oligomeric standards containing a range of arabinose, galactose, glucose, xylose, and mannose concentrations were treated in the same manner as samples, and a sugar degradation factor was applied to oligomeric sugar calculations. SSL Pulp or Sludge add to H 2 O or SSL SSL Sugars Bioproducts (e.g. ethanol, xylitol) Figure 2. Experimental design of converting pulp, sludge, and SSL to bioproducts via fermentation, SHF, and SSF. In SHF, solid streams are combined with either water or SSL. In SSF, solid streams are combined with SSL and converted in one step. 17

23 Enzymatic hydrolysis and fermentation We explored process designs by which pulp, sludge, and SSL could be converted to a sugar platform from which biochemicals could be produced. The resulting schematic of converting pulp and sludge to ethanol, and SSL to ethanol and xylitol, is shown in Figure 2. Separate hydrolysis and fermentation (SHF) was used to convert the three streams separately, and simultaneous saccharification and fermentation (SSF) was used to convert SSL fortified with each of the two solid streams in one step. SSL was fermented to ethanol and xylitol. Saccharification Hydrolysis was carried out in 125 ml (50 ml reaction volume) Erlenmeyer flasks in triplicate on washed solid materials. Solids were enzymatically hydrolyzed at 10% (w/v) consistency in both water and SSL with ph adjusted to 4.8. Flasks were incubated at 50 C and 150 rpm in an orbital shaker (New Brunswick). Enzymes added were cellulase at 5 FPU/g cellulose (Spezyme, Genencor, Palo Alto, CA) and β-glucosidase at 10 CBU/g cellulose (Novozymes 188, Bagsverd, Denmark). For controls, the same amount of enzyme added to pulp and sludge flasks was respectively added to flasks containing plain SSL. Samples of 1 ml volume were taken periodically over 48 h, boiled at 100 C for 5 min to denature enzymes, and stored at -20 C until HPLC analysis. Fermentation (Saccharomyces cerevisiae) Prior to fermentation, Saccharomyces cerevisiae (ATCC 96581) isolated from spent sulfite liquor (Linden, Peetre et al. 1992) was streaked onto YPD agar plates and allowed to grow for 48 h. Sterile liquid media containing 10 g/l each of glucose, yeast extract, and peptone was inoculated with one colony from the plate. Cells were grown for a total of 48 h at 30 C and 150 rpm in an orbital shaker, with fresh media replaced at 24 h. Cells were then spun down, washed twice in water, and resuspended in a small volume of 0.5% NaCl. Cell concentration was determined by comparing optical density of the cell suspension at 600 nm to a calibration curve. After completion of hydrolysis, the remaining liquid hydrolysate was boiled at 100 C to denature enzymes and vacuum filtered through filter paper. The resulting filtrate was collected and nutrients were added in the form of (NH 4 ) 2 HPO 4 at 2 g/l, Na 2 SO 4 at 0.2 g/l, and NaNO 3 at 2 g/l. Hydrolyzed SSL, and a solution of 10 g/l each of glucose, galactose, and mannose in water were also treated in this manner to use as controls. The ph was adjusted to 6.0 with 50% w/v NaOH and S. cerevisiae 18

24 was added at a concentration of 5 g/l. Fermentation was done in 50 ml volume in 125 ml Erlenmeyer flasks, incubated at 30 C and 150 rpm for 72 h in an orbital shaker. Samples of 1 ml volume were taken periodically and centrifuged at 10,000 rpm for 5 min. Supernatant was collected through 0.22 μm syringe filters and stored at -20 C, while pellets were washed and optical density measured at 600 nm to determine cell concentration. Fermentation (Candida guilliermondii) Fermentation was also performed on hydrolyzed SSL spiked with synthetic xylose up to a 30 g/l total concentration using C. guilliermondii (ATCC ) to demonstrate xylitol production. This was done in an identical manner as fermentation with S. cerevisiae except that the nutrients added in this case were 5 g/l urea, 1.7 g/l yeast nitrogen base, and 1 g/l yeast extract. Two controls containing 30 g/l each of glucose and xylose were handled in the same manner. SSF SSL containing the same nutrient concentrations as in SHF was adjusted to ph 5.5 with 50% w/v NaOH. The solution was fortified with pulp or sludge at 10% consistency, and enzyme and S. cerevisiae were added at 5 FPU/g cellulose (10 CBU/g β-glucosidase) and 5 g/l, respectively. Samples were run in triplicate in 100 ml volume at 37 C in an orbital shaker for 72 h. A control containing 10 g/l each of galactose, glucose, and mannose was prepared and analyzed identically. Analysis of sugars and ethanol Monosaccharides were quantified using a Dionex (Sunnyvale, CA) HPLC (ICS-3000) system equipped with an AS50 autosampler, ED50 electrochemical detector, GP50 gradient pump, and anion exchange column (Dionex, CarboPac PA1). Deionized water at 1 ml/min was used as an eluent, with a postcolumn addition of 0.2 M NaOH. The injection volume was 5 μl. Standards were prepared containing arabinose, galactose, glucose, xylose, and mannose in an appropriate range of concentrations that fully encompassed the range found respectively in the samples. Samples were filtered through 0.22 µm syringe filters and an internal standard, fucose, was added at 0.2 g/l. Ethanol, xylitol, acetic acid, furfural, and hydroxymethylfurfural (HMF) were measured using a Shimadzu (Columbia, MD) HPLC system equipped with a RID-10A differential refractometric detector, LC-20AD solvent delivery module, SIL-20AHT autosampler, and a Rezex RHM monosaccharide H+ anion exchange column (Phenomenex, Torrance, CA). An isocratic mobile 19