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

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:__________________________________

i

TABLE OF CONTENTS Page

List of figures ............................................................................................ ii

List of tables ............................................................................................ iii

Chapter 1: Introduction ............................................................................ 1

1.1 Lignocellulosic biomass................................................................. 3

1.2 Pretreatment ................................................................................... 6

1.3 Sulfite pulping ................................................................................ 8

1.4 Enzymatic hydrolysis ..................................................................... 8

1.5 Fermentation ................................................................................ 10

1.6 SHF vs. SSF ................................................................................. 12

1.7 Products from a sugar platform .................................................... 13

Chapter 2: Draft paper ........................................................................... 14

Introduction ..................................................................................... 15

Methods .......................................................................................... 16

Results and discussion .................................................................... 20

Conclusion ...................................................................................... 26

Chapter 3: Additional results and discussion ......................................... 28

3.1 Fermentation of SSL using S. cerevisiae ..................................... 28

3.2 Effect of solids loading on enzymatic hydrolysis ........................ 29

3.3 Effect of enzyme loading on hydrolysis ...................................... 30

Chapter 4: Additional conclusions and future work .............................. 32

Acknowledgements ................................................................................. 33

References ............................................................................................... 34

ii

LIST OF FIGURES

Page

1. Bioconversion process schematic………………………………2

2. Experimental design…………………………………………...17

3. Spent sulfite liquor C. guilliermondii fermentation……………22

4. Pulp and sludge hydrolysis….…………………………………23

5. Fermentation of pulp hydrolysates…………………………….24

6. Fermentation of sludge hydrolysates…………………………..25

7. Combined feedstock SSF………………………………………26

8. Spent sulfite liquor S. cerevisiae fermentation………………...28

9. Variable consistency hydrolysis……………………………….29

10. Variable enzyme loading hydrolysis…………………………..31

iii

LIST OF TABLES

Page

1. Pulp and sludge composition…………………………………..20

2. Spent sulfite liquor composition……………………………….21

3. Solid stream result summary…………………………………..27

1

Chapter 1: Introduction

The global impact of human energy consumption has been made increasingly apparent in recent years

by growing rates of atmospheric CO2 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 full-

scale 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.

2

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.

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).

Objective and Outline

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

Lignocellulosic biomass

Pretreatment Hydrolysis Fermentation

Bioproducts (ethanol, xylitol, etc.)

3

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 4.

1.1 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

4

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 100-150

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 100-200. 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.

5

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

6

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 seasonally-

dependent 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 lignin-

degrading microorganisms.

Steam explosion is a method that has been found to be very efficient for woody feedstocks. Biomass

is subjected to high temperature (185-240°C) and pressure (87-500 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 H2SO4 or SO2

to raw biomass before steam explosion has been found to improve pretreatment efficiency (Ramos,

Breuil et al. 1992; Eklund, Galbe et al. 1995).

7

Ammonia fiber expansion (AFEX) pretreatment is a similar process to steam explosion, in which

biomass is combined with ammonia at 100-400 psi and 70-200C 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 H2SO4, 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 RO = t * e(Tr-Tb)/14.75

Where t = residence time (m), Tr = reaction temperature, and Tb = 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

8

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 SO2 at

high temperature and acidic pH. Pulping occurs through the interaction of lignin with SO2, 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

9

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 0.1875 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 end-

product 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

10

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:

C6H12O6 2 C2H5OH + 2 CO2

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.

11

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 201935) 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 C5H10O5 + 12 ADP + 12 Pi + 12 H2O + 3 O2 54 C5H12O5 + 12 ATP + 30 CO2

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).

12

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 50C and a

pH of 4.8, whereas most microbes ferment better at temperatures near 30C 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 end-

product 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 37C) 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).

13

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

14

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.

15

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 energy-

intensive 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. 2007 and Kang et al.

2010. 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. 2008. 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

16

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 SO2 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 ClO2. 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) H2SO4 for 2

h. Samples were then diluted to 4% (w/w) H2SO4, autocalved at 121°C for 1 h, and filtered through

glass fritted crucibles. Filtrate was collected, carbohydrate content analyzed by HPLC, and acid-

17

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-510-

42623 (Sluiter, Hames et al. 2004). Five mL SSL was added to 0.697 mL of 72% (w/w) H2SO4 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.

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.

Hydro

lysis

SS

F

Fe

rme

nta

tio

n

Fe

rme

nta

tio

n

Pulp or Sludge SSL

Bioproducts (e.g. ethanol, xylitol)

add to

SSL H2O or SSL

Sugars

18

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 (NH4)2HPO4 at 2 g/L, Na2SO4 at 0.2 g/L, and NaNO3 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

19

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 201935) 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

20

phase of 5 μM H2SO4 was used at a flow rate of 0.6 mL/min, and a 20 μL injection volume was used

for each sample. Standards of ethanol, xylitol, acetic acid, furfural, and HMF were prepared in

appropriate concentrations. Samples were analyzed by relating to standard curves generated by the

prepared standards of known concentration. Calculation of standard curves and standard deviation

was done in Microsoft Excel.

Results and discussion

Chemical composition of pulp, sludge, and SSL

The chemical composition of pulp and sludge in terms of polysaccharides, lignin, and ash is shown in

Table 1. Both solid streams were mainly comprised of glucan (pulp 88.6%, 70.2%), contained a

small amount of acid insoluble lignin (pulp 3.3%, sludge 12.3%), and very minimal acid soluble

lignin (pulp 1.6%, sludge 0.7% (Table 1). Pulp contained more glucan and less total lignin than

sludge, an expected outcome of the pulping process. Mannan was the next most abundant

polysaccharide (pulp 6.6%, sludge 4.4%) for both streams due to their derivation from mostly

softwoods. As expected, neither stream contained a detectable amount of arabinan or galactan. At

9.9%, sludge contained more ash than pulp, at 0.2%, a result of being partially sourced from boiler

house effluent. It has been observed that similar fiber sludges, derived from Kraft pulping, and

containing as little as 32.3% glucan and as much as 24% total lignin, can still be hydrolyzed into

monomeric sugars for ethanol production (Sjöde, Alriksson et al. 2007).

Table 1. Percent composition of solid streams based on TAPPI method T-222 om-98. Acid insoluble

lignin is shown as AIL, and acid soluble lignin is ASL.

Glucan (%) Xylan (%) Mannan (%) AIL (%) ASL (%) Ash (%)

Pulp 88.6 2.8 6.6 3.3 1.6 0.2

Sludge 70.2 3.1 4.4 12.3 0.7 9.9

The monomeric and oligomeric sugar composition of SSL is shown in Table 2. Due to softwood

hemicellulose being mainly comprised of galactoglucomannans and Arabino-4-O-

methylglucuronoxylans (Fengel and Wegener 1989), the most abundant sugar was mannose (14.0 g/L

monomers), followed by xylose (7.6 g/L monomers). Glucose (5.1 g/L) was the next most abundant

sugar in SSL, followed by minimal amounts of galactose (3.4 g/L) and arabinose (1.2 g/L). All five

sugars were mostly in monomeric form (83%), making SSL a readily-available monosaccharide

21

source. Acetic acid, hydroxymethylfurfural (HMF) and furfurals were also present in SSL at

concentrations of 5.0, 0.11 and 0.16 g/L, respectively (Table 2). An unevaporated SSL stream

containing 31.3 g/L total sugar was used for this study, but this can be concentrated onsite up to 97

g/L total sugar if a higher sugar concentration is desired (Sande 2010).

Table 2. Chemical composition of SSL and oligomer % of total based on NREL LAP TP-510-42623

(Sluiter, Hames et al. 2004). *Acetic acid 5 g/L, HMF 0.11 g/L, furfural 0.16 g/L.

Ara (g/L) Gal (g/L) Glu (g/L) Xyl (g/L) Man (g/L) Total 5-C

sugars (g/L)

Total 6-C

sugars (g/L)

Monomer 1.2 3.4 5.1 7.6 14.0 8.8 22.5

Oligomer 0.11 0.40 0.92 0.04 3.86 0.15 5.18

Monomer

% of total

91.5 98.6 84.7 99.5 78.4 98.4 81.3

Due to its high glucan content and minimal lignin and ash, sulfite pulp, if successfully hydrolyzed,

can be expected to produce a clean, single-sugar stream (glucose). Sludge may produce similar

results, though its higher lignin content is expected to reduce hydrolysis yields as compared to pulp.

SSL, having a higher concentration of both hexoses and pentoses, is already an excellent source of

mixed sugars, most of which are in monomeric form.

Fermentation of SSL

As the high concentration of monomeric sugars in SSL has been demonstrated through its

compositional analysis, we next utilized this mixed sugar stream for fermentation to test its potential

conversion to bioproducts. Figure 3 shows fermentation of SSL utilizing C. guilliermondii, which

ferments hexoses to ethanol and xylose to xylitol. Due to the low concentration of native xylose (7.6

g/L), SSL was spiked with synthetic sugar to demonstrate the full potential of xylose to xylitol

fermentation. Glucose was consumed after 21 h, followed mannose and galactose at 33 h. This

pattern of consumption was consistent with that of a Tembec yeast strain and S. cerevisiae during

mixed synthetic sugar fermentation (Keating, Robinson et al. 2004). Total consumption of

approximately 30 g/L xylose required 120 h. Ethanol and xylitol were produced concurrently.

Ethanol concentration reached 9.5 g/L after 72h, corresponding to 89.5% of the theoretical yield.

This was higher than the ethanol yield achieved with a mixture of synthetic sugars (77%, unpublished

22

data). This may be explained by the presence of inhibitors in SSL, acetic acid in particular, which in

low concentrations have been found to enhance fermentation by C. guilliermondii (Keating,

Panganiban et al. 2006). Xylitol yield was 10.5 g/L after 96 h, or 40.3% of the theoretical yield. This

was lower than that of a mixed synthetic sugar control (47%, unpublished data), an expected result, as

mixed 5- and 6-C sugar fermentation with C. guilliermondii has been observed to produce lower

xylitol yields than single-sugar fermentation (Lee, Sopher et al. 1996). Overall xylitol yields may

also improve by pre-growing yeast in xylose media, and/or fermenting from SSL streams with a

higher xylose:glucose ratio, as a ratio of 5:1 has been found to be ideal when xylitol is the only

product (da Silva and de Almeida Felipe 2006). In addition, 73% of the initial acetic acid was

consumed, demonstrating that C. guilliermondii can concurrently detoxify SSL during fermentation

(Figure 3). HMF and furfural concentrations were initially low, and completely consumed after 3 h.

These observed yields and minimal interference by inhibitory compounds demonstrate the use of SSL

as an excellent feedstock for producing higher-value bioproducts, exemplified by ethanol and xylitol.

0 20 40 60 80 100 120

0

5

10

15

20

25

30

Ara

Gal

Glu

Xyl

Man

AA

EtOH

XOH

Time (hours)

Sugars

, A

A c

oncentr

ation (

g/L

)

0

2

4

6

8

10

12

EtO

H, X

OH

concentra

tion (g

/L)

Figure 3. Fermentation of spent sulfite liquor (SSL), spiked with synthetic xylose,

to ethanol and xylitol using 5 g/L Candida guilliermondii at 30°C.

SHF of pulp and sludge

As pulp and sludge have previously been shown to contain high amounts of glucan, the

hydrolysability of these two solid streams was examined next in order to determine the feasibility of

their use in bioconversion. Preliminary hydrolysis work was done by first varying the solids

89%

40%

23

consistency, then the enzyme loading, to determine the most desirable conditions for SHF, based on

yield. Solids were enzymatically hydrolyzed at 2%, 10%, and 15% (w/v) consistency and 10 FPU/g

cellulose enzyme loading. In both cases, hydrolysis at 10% consistency yielded high glucose

concentrations (87 g/L for pulp, 62 g/L for sludge, not shown) after 24 h. Next, enzymes loadings of

1, 2, 5, 10, and 15 FPU/g cellulose were used to hydrolyze 2% (w/v) consistency slurries of each

solid. In this case, 5 FPU/g cellulose resulted in reasonably high glucose yields (18 g/L for pulp, 14

g/L for sludge) after 24 h without sacrificing enzyme cost. Therefore, SHF experiments were all

performed at 10% (w/v) solids consistency and 5 FPU/g cellulose enzyme loading.

0 20 40 60 80 100

0

10

20

30

40

50

60

70

80

Cellu

lose to g

lucose c

onvers

ion (

%)

Time (hours)

Pulp / H2O

Pulp / SSL

Sludge / H2O

Sludge / SSL

Figure 4. High consistency (10% w/v) hydrolysis of pulp and sludge in both water

and SSL with 5 FPU/g cellulose enzyme loading.

The hydrolysis of pulp and sludge in both water and SSL is show in Figure 4. Hydrolysis in SSL was

done to demonstrate a method of increasing the amount of sugar available for bioproduct conversion.

Hydrolysis of pulp in SSL resulted in 49% glucose conversion (48.5 g/L), compared to hydrolysis of

pulp in water, where 80% of glucose was hydrolyzed (78.8 g/L) (Figure 4, Table 2). This difference

in yield is attributed to enzyme inhibition during mixed feedstock hydrolysis, where increasing

concentrations of SSL during hydrolysis of knot rejects resulted in decreasing hydrolysis yields

(Helle, Petretta et al. 2007). Conversely, hydrolysis of sludge in SSL yielded 43% conversion of

glucose (33.5 g/L), compared to 52.4% (40.8 g/L) for sludge in water (Figure 4, Table 2). In this

case, the addition of a liquid sugar stream appeared to enhance the hydrolysis of sludge, but the same

was not true for pulp. In both cases, with water and with SSL, pulp hydrolyzed more completely than

79 g/L

34 g/L 49 g/L

41 g/L

24

did sludge, an expected result due to the higher lignin and ash content of sludge, which has been

found to inhibit hydrolysis enzymes (Kang, Wang et al. 2010). Sludge hydrolysis was enhanced by

the addition of SSL, however, pulp hydrolyzed more completely in water.

0 5 10 15 20

0

10

20

30

40

50

60

70

Pulp / H2O Glucose

Pulp / SSL Hexose

Pulp / H2O EtOH

Pulp / SSL EtOH

Time (hours)

Sugar

concentr

ation (

g/L

)

0

10

20

30

EtO

H c

oncentra

tion (g

/L)

Figure 5. Fermentation of pulp hydrolysates using 5 g/L Saccharomyces cerevisiae

at 30°C.

Fermentation of hydrolysates was performed to demonstrate the possibility of converting hydrolyzed

sugars to high value products, exemplified by ethanol. Figure 5 shows the fermentation of pulp

hydrolysates by S. cerevisiae, which utilizes hexoses for ethanol production. For pulp in water,

glucose was fully consumed after 6 h. For pulp in SSL, total hexose concentration reached 1.7 g/L

after 6 h, but was never fully exhausted due to incomplete consumption of galactose (Figure 5).

Ethanol yield as a percentage of theoretical yields was 77.8% (28.3 g/L) for pulp in water, and 76.5%

(19.9 g/L) for pulp in SSL, which were similar to that of a concurrently fermented mixed hexose

control (13.0 g/L, 78.2%, not shown). However, a much higher hydrolysis yield for pulp in water

contributed to a significantly higher total SHF process yield and 42% more ethanol, as compared to

pulp in SSL (Table 3). The fermentation of sludge hydrolysates is shown in Figure 6. Glucose was

fully consumed after 4 h for sludge in water, and hexose fully consumed after 6 h for sludge in SSL.

Again, ethanol yields as a percent of the theoretical maximum were similar, 76.2% (10.9 g/L) for

sludge in water, and 73.1% (18.6 g/L) for sludge in SSL, and close to that of the synthetic sugar

control. However, final ethanol concentration was 71% greater for fermentation of sludge in SSL due

to the presence of additional sugars in the liquid stream, from monomers and hydrolysis of some

78%

77%

25

oligomers present in SSL. Fermentation yields were consistent between all four hydrolysates and the

hexose control, demonstrating that fermentation did not account for differences in final ethanol

concentration. Instead, ethanol yield is more likely to be improved by (1) improving cellulose to

glucose conversion during hydrolysis, and (2) increasing the initial availability of sugars by, in some

cases, mixing solid and liquid sugar streams.

0 5 10 15 20

0

10

20

30

40

50

Sludge / H2O Glucose

Sludge / SSL Hexose

Sludge / H2O EtOH

Sludge / SSL EtOH

Time (hours)

Sugar

concentr

ation (

g/L

)

0

5

10

15

20

EtO

H c

oncentra

tion (g

/L)

Figure 6. Fermentation of sludge hydrolysates using 5 g/L S. cerevisiae at 30°C.

SSF of pulp and sludge

In order to evaluate a method of mixed-stream bioconversion other than SHF, SSF was done to

generate mixed-sugar streams from the combination of SSL fortified with pulp and sludge. This data

is shown in Figure 7. SSF of pulp in SSL produced 56.1% (31.9 g/L) of the theoretical ethanol yield,

and sludge in SSL produced 50% (23.7 g/L). Total process yield as a percent of the theoretical

maximum improved dramatically in comparison to SHF of the same mixed streams, from 37.7% for

pulp in SSL and 38.3% for sludge in SSL. Despite the potential benefits of SSF arising from the

reduced effect of end-product inhibition (Moritz and Duff 1996), this result was unexpected, because

fortification of SSL with hydrolysate has been found to decrease fermentation yield (Smith, Cameron

et al. 1997). Since the consistency of S. cerevisiae fermentation of these sugar streams has already

been demonstrated, this increase in ethanol yield can only be attributed to improvements in

hydrolysis.

73%

76%

26

0 10 20 30 40 50 60 70

0

2

4

6

8

10

12

14

Pulp / SSL Hexoses

Sludge / SSL Hexoses

Pulp / SSL EtOH

Sludge / SSL EtOH

Time (hours)

Sugar

concentr

ation (

g/L

)

0

5

10

15

20

25

30

35

EtO

H c

oncentra

tion (g

/L)

Figure 7. Simultaneous saccharification and fermentation (SSF) of SSL fortified

with pulp and sludge using 5 FPU/g cellulose enzyme loading and 5 g/L S.

cerevisiae at 37°C.

Conclusion

Single and mixed sugar streams were generated from sulfite pulp, sludge, and SSL in this study.

SSL, due to being comprised of both hexoses and pentoses in mostly monomeric form, was shown to

be an excellent mixed-sugar stream. Fermentation of SSL with C. guilliermondii produced high

ethanol yields (89.5%), and xylitol yields consistent with that of a synthetic sugar control (40.3%)

(Figure 3). Pulp proved to be an excellent feedstock for single-sugar generation by producing a clean

glucose stream conducive to fermentation with hexose metabolizing organisms. SHF of pulp in water

resulted in the highest overall ethanol yield during fermentation for that feedstock (62.2%, 28.3 g/L)

during fermentation with S. cerevisiae (Table 3). Mixing pulp with SSL during SHF severely

decreased yields (37.7%, 19.9 g/L). Conversely, and unexpectedly, the best utilization of sludge

found in this study, in terms of yield, was as a supplement to the mixed sugar stream available from

SSL. Mixing sludge with SSL during SSF produced the highest ethanol yield (50.0%, 23.7 g/L)

during S. cerevisiae fermentation (Table 3). For mixed feedstocks, SSF improved yields for both

pulp and sludge. Fermentation yields were consistent across all streams, demonstrating that

hydrolysis, not fermentation is the rate-limiting step of SHF and SSF.

56%

50%

27

Table 3. Glucose and ethanol yields shown in concentration (g/L) and percentage of the theoretical

maximum for SHF and SSF. Glucose concentrations for mixed feedstock hydrolysis (pulp or sludge

with SSL) are corrected to exclude the sugars in SSL.

Pulp / H2O Pulp / SSL Sludge / H2O Sludge / SSL

Hydrolysis Cellulose to glucose

conversion (%)

80.0% 49.3% 43.0% 52.4%

Glucose

concentration (g/L)

78.8 48.5 33.5 40.8

Fermentation Hexose to ethanol

yield (%)

77.8% 76.5% 76.2% 73.1%

Ethanol concentration

(g/L)

28.3 19.9 10.9 18.6

SHF Original biomass to

ethanol yield (%)

62.2% 37.7% 32.8% 38.3%

SSF Original biomass to

ethanol yield (%)

56.1% 50.0%

Ethanol concentration

(g/L)

31.9 23.7

Acknowledgements

We are thankful to Kimberly-Clark for providing funding and the feedstocks utilized in this paper.

NSF-IGERT also provided additional funding.

28

Chapter 3: Additional results and discussion

3.1 Fermentation of SSL using S. cerevisiae

0 5 10 15 20 25 30

0

2

4

6

8

10

12

Time (hours)

Concentr

ation (

g/L

)

Gal

Glu

Man

Yeast

Gly

AA

EtOH

0

2

4

6

8

EtO

H c

oncentra

tion (g

/L)

Figure 8. Fermentation of SSL using S. cerevisiae at 30°C. Final ethanol conversion

is shown in % of theoretical yield. Concentration of yeast, glycerol (Gly), acetic

acid (AA) are shown in addition to hexoses and ethanol. Furfural and HMF were

present at insignificant concentrations (<0.1 g/L) and fully consumed after 2 h.

S. cerevisiae fermentation of SSL was done to demonstrate tolerance of the strain to inhibitors present

in SSL. Ethanol concentration was 7.8 g/L after 30 h, representing 88% of the theoretical maximum

(Figure 8). Hexoses became fully consumed in the order of glucose, then mannose, followed by

galactose, a result that has been observed in previous study utilizing the same strain to ferment SSL

(Linden, Peetre et al. 1992). Ethanol yield was higher than that found for a concurrently run mixed

hexose control (10.1 g/L, 78.2%, data not shown), possibly explained by the enhancement of

fermentation by a small concentration of inhibitors (Keating, Panganiban et al. 2006). Yield was

comparable to that of C. guilliermondii (98.5%), though due to the presence of xylose and its

fermentability to xylitol, described in Paper I, utilization of C. guilliermondii for SSL fermentation

may be more beneficial.

88%

29

3.2 Effect of solids loading on enzymatic hydrolysis

(a)

0 10 20 30 40 50

0

20

40

60

80

100

Cellu

lose to g

lucose c

onvers

ion (

%)

Time (hours)

Pulp 2%

Pulp 10%

Pulp 15%

(b)

0 10 20 30 40 50

0

20

40

60

80

100

Cellu

lose to g

lucose c

onvers

ion (

%)

Time (hours)

Sludge 2%

Sludge 10%

Sludge 15%

Figure 9. Pulp (a) and sludge (b) hydrolysis at 10 FPU/g cellulose enzyme loading

with 2%, 10%, and 15% (w/v) solids consistency. Glucose concentration at 24 h is

shown in g/L.

Prior to the hydrolysis reported in Paper I, experiments were conducted to determine the extent to

which high solids consistency would yield sufficient glucose conversions. Subsequent solids

consistency was selected based on this analysis. This was done by performing hydrolysis on a range

21 g/L

87 g/L

89 g/L

15 g/L

62 g/L

82 g/L

30

of solids loadings (2-15% w/v) while keeping enzyme loading at 10 FPU/g cellulose (Figure 9).

Though carried out for 48 h, a short turnover time is favorable in an industrial setting, so conversion

at 24 h was examined to determine the desired solids loading for all subsequent SHF and SSF. As

expected, percent glucose conversion decreased with increased solids loading for both pulp and

sludge. For pulp, percent glucose conversion did not differ between 2% and 10% consistency (both

89%), but dropped significantly to 57% when consistency was increased to 15%. For sludge, percent

conversion between 2% and 10% consistency at 24 h differed by a greater margin (83% and 68%,

respectively), however this also corresponded to a desirable increase in glucose concentration from

14.4 g/L to 61.4 g/L. High consistency hydrolysis has previously been described as a desirable way

of reducing cost, because it produces a highly concentrated sugar streams, and therefore results in a

more concentrated product stream. Taking into account both % glucose conversion and sugar

concentration in the final hydrolysate, a solid consistency of 10% was selected for subsequent the

analyses presented in Paper I.

3.3 Effect of enzyme loading on hydrolysis

Enzyme loading was also selected based on analyses that took place prior to those presented in Paper

I. The effect of enzyme loading on hydrolysis was explored by hydrolyzing pulp and sludge at 2%

consistency using a gradient of enzyme loading of 1-20 FPU/g cellulose (Figure 10). As predicted,

increased enzyme loading had a positive, nonlinear effect on percent glucose conversion. For pulp,

doubling the amount of enzyme from 5 to 10 FPU/g cellulose resulted in only a 10% gain in glucose

conversion (79% to 89%). Comparatively, reducing to 2 FPU/g cellulose enzyme loading resulted in

a significant drop in percent conversion (46%). Similar results were found for sludge, where glucose

conversion at 2, 5, and 10 FPU/g cellulose for sludge was 49%, 76%, and 83%, respectively after 24 h

(Fig. 3b). As previously described, reducing enzyme loading to the extent that glucose conversion is

not significantly effected is a favorable industrial practice that reduces cost. Based on this analysis,

an enzyme loading of 5 FPU/g cellulose was selected for subsequent SHF and SSF.

31

(a)

0 10 20 30 40 50

0

20

40

60

80

100

Cellu

lose to g

lucose c

onvers

ion (

%)

Time (hours)

Pulp 20 FPU

Pulp 10 FPU

Pulp 5 FPU

Pulp 2 FPU

Pulp 1 FPU

(b)

0 10 20 30 40 50

0

20

40

60

80

100

Cellu

lose to g

lucose c

onvers

ion (

%)

Time (hours)

Sludge 20 FPU

Sludge 10 FPU

Sludge 5 FPU

Sludge 2 FPU

Sludge 1 FPU

Figure 10. Pulp (a) and sludge (b) hydrolysis at 2% consistency with 1, 2, 5, 10, and

20 FPU/g cellulose enzyme loading. Glucose concentration at 24 h is shown in g/L.

21 g/L

21 g/L

18 g/L

11 g/L

7 g/L

15 g/L

15 g/L

14 g/L

10 g/L

6 g/L

32

Chapter 4: Additional conclusions and future work

The production of single and mixed sugar streams from pulp, sludge, and SSL has been demonstrated

in this thesis and the enclosed draft paper. Fermentation of SSL with S. cerevisiae produced ethanol

yields equivalent to that of C. guilliermondii, presented in the draft paper (88%, and 89.5%,

respectively). However, the prospect of exploiting xylose, present in SSL at 5.1 g/L, makes the use of

C. guilliermondii or a similar pentose-metabolizing microorganism a more appealing option due to

the potential of making two products from SSL. From the hydrolysis results presented in Chapter 3,

10% (w/v) consistency and 5 FPU/g cellulose enzyme loading was determined to be the most

desirable parameters for the high consistency, low enzyme loading hydrolysis results presented in the

draft paper. However, this determination was based on laboratory-scale yield alone, and the

economic viability of these conditions for a full-scale industrial operation must be evaluated prior to

implementation. For the purposes of this thesis and draft paper, these hydrolysis parameters were

sufficient.

From the draft paper presented in Chapter 2, it was concluded that SSL is a readily-available source

of mixed 5- and 6-C sugars, which are mostly in monomeric form. Pulp proved to a be desirable

feedstock for a single sugar stream (glucose) due to its 88.6% glucan content, 80% of which was

hydrolyzed in water. Mixing of pulp with SSL yielded low cellulose to ethanol conversions for both

SHF (37.7%), and SSF (56.1%). Sludge hydrolyzed more completely when combined with SSL

during SSF, yielding 50% total cellulose to ethanol conversion.

Future work

The objective of generating single- and mixed-sugar streams from sulfite pulp, sludge, and SSL has

been achieved in this thesis. However, the full potential of bioproduct conversion during the

fermentation step has not been entirely explored. First, growth of yeast on SSL prior to fermentation

can be explored as a means of better adapting the yeast to inhibitory compounds within SSL. In

addition to the potential of boosting yields, this reduces the cost spent on synthetic sugars for the

propagation of yeast. Secondly, production of other biochemicals via fermentation with a wider

variety of sugar-metabolizing microorganisms can be explored. On an industrial scale, flexibility in

the choice of microorganism would allow the bioconversion process to adjust according to the market

value of the variety of bioproducts that can be made. Finally, techno-economic and life-cycle

analyses of the proposed bioconversion processes of pulping streams to bioproducts would benefit

decisions regarding industrial-scale operation. In particular, comparison of the processes utilized here

33

with dedicated lignocellulose-to-ethanol facilities would be valuable. Continued research along these

lines would allow the biochemicals industry to tap into a currently unused source of sugars for

bioproduct conversion.

Acknowledgements

This thesis is the result of 2 ½ years of work in the School of Forest Resources at the University of

Washington. I give my sincere thanks to my advisor and committee chair, Renata Bura, and

committee members Rick Gustafson and Bill McKean for their guidance through the duration of my

research and thesis writing. I would also like to thank Doug Asbe, Wally Sande, and Jeff Ross at

Kimberly-Clark for their collaboration on this project and for providing the feedstocks. I’m also

grateful to NSF-IGERT for providing funding through the duration of my graduate work.

34

References

Abatzoglou, N., E. Chornet, et al. (1992). "Phenomenological kinetics of complex systems: the

development of a generalized severity parameter and its application to lignocellulosics

fractionation." Chemical Engineering Science 47(5): 1109-1122.

Akin, D. E. (2007). Grass Lignocellulose. Applied Biochemistry and Biotecnology. J. R. Mielenz, K.

T. Klasson, W. S. Adney and J. D. McMillan, Humana Press: 3-15.

Alfani, F., A. Gallifuoco, et al. (2000). "Comparison of SHF and SSF processes for the bioconversion

of steam-exploded wheat straw." Journal of Industrial Microbiology &amp; Biotechnology

25(4): 184-192.

Alizadeh, H., F. Teymouri, et al. (2005). "Pretreatment of switchgrass by ammonia fiber explosion

(AFEX)." Applied Biochemistry and Biotechnology 124(1): 1133-1141.

Atalla, R. H. and D. L. VanderHart (1999). "The role of solid state 13C NMR spectroscopy in studies

of the nature of native celluloses." Solid State Nuclear Magnetic Resonance 15(1): 1-19.

Banerjee, N., R. Bhatnagar, et al. (1981). "Development of resistance in Saccharomyces cerevisiae

against inhibitory effects of Browning reaction products." Enzyme and Microbial Technology

3(1): 24-28.

Barbosa, M. F. S., M. B. de Medeiros, et al. (1988). "Screening of yeasts for production of xylitol

fromd-xylose and some factors which affect xylitol yield in Candida guilliermondii." Journal

of Industrial Microbiology &amp; Biotechnology 3(4): 241-251.

Bergeron, P. (1996). Environmental impacts of bioethanol. Handbook on ethanol: Production and

utilization. C. E. Wyman, Taylor & Francis: 90-93.

Blanchette, R. A. (1991). "Delignification by Wood-Decay Fungi." Annual Review of

Phytopathology 29(1): 381-403.

Brown, R. M. (2004). "Cellulose structure and biosynthesis: What is in store for the 21st century?"

Journal of Polymer Science Part A: Polymer Chemistry 42(3): 487-495.

Brown, S. W., S. G. Oliver, et al. (1981). "Ethanol inhibition of yeast growth and fermentation:

Differences in the magnitude and complexity of the effect." Applied Microbiology and

Biotechnology 11(3): 151-155.

Canadell, J. G., C. Le Quéré, et al. (2007). "Contributions to accelerating atmospheric CO2 growth

from economic activity, carbon intensity, and efficiency of natural sinks." Proceedings of the

National Academy of Sciences 104(47): 18866-18870.

35

Cantarella, M., L. Cantarella, et al. (2004). "Comparison of different detoxification methods for

steam-exploded poplar wood as a substrate for the bioproduction of ethanol in SHF and SSF."

Process Biochemistry 39(11): 1533-1542.

Cara, C., M. Moya, et al. (2007). "Influence of solid loading on enzymatic hydrolysis of steam

exploded or liquid hot water pretreated olive tree biomass." Process Biochemistry 42(6):

1003-1009.

Clark, J. and F. Deswarte, Eds. (2008). Introduction to chemicals from biomass. Renewable

resources. West Sussex, Wiley.

da Silva, D. D. V. and M. d. G. de Almeida Felipe (2006). "Effect of glucose:xylose ratio on xylose

reductase and xylitol dehydrogenase activities from Candida guilliermondii in sugarcane

bagasse hydrolysate." Journal of Chemical Technology & Biotechnology 81(7): 1294-1300.

Decker, S., W. Adney, et al. (2003). "Automated filter paper assay for determination of cellulase

activity." Applied Biochemistry and Biotechnology 107(1): 689-703.

Delgenes, J. P., R. Moletta, et al. (1996). "Effects of lignocellulose degradation products on ethanol

fermentations of glucose and xylose by Saccharomyces cerevisiae, Zymomonas mobilis,

Pichia stipitis, and Candida shehatae." Enzyme and Microbial Technology 19(3): 220-225.

DeLong, E. A. (1977). Method of rendering lignin separable from cellulose and hemicellulose in

lignocellulosic material and the product so produced. C. P. a. D. Limited. Canada.

Ding, S.-Y. and M. E. Himmel (2006). "The Maize Primary Cell Wall Microfibril:  A New Model

Derived from Direct Visualization." Journal of Agricultural and Food Chemistry 54(3): 597-

606.

Ding, S.-Y. and M. E. Himmel (2008). Anatomy and ultrastructure of maize cell walls: An example

of energy plants. Biomass recalcitrance: Deconstructing the plant cell wall for bioenergy. M.

E. Himmel, Wiley-Balckwell: 38-60.

Eklund, R., M. Galbe, et al. (1995). "The influence of SO2 and H2SO4 impregnation of willow prior

to steam pretreatment." Bioresource Technology 52(3): 225-229.

Eriksson, T., J. Karlsson, et al. (2002). "A model explaining declining rate in hydrolysis of

lignocellulose substrates with cellobiohydrolase I (Cel7A) and endoglucanase I (Cel7B) of

Trichoderma reesei." Applied Biochemistry and Biotechnology 101(1): 41-60.

Fan, L., Y.-H. Lee, et al. (1982). The nature of lignocellulosics and their pretreatments for enzymatic

hydrolysis. Microbial Reactions, Springer Berlin / Heidelberg. 23: 157-187.

Felby, C., L. Thygesen, et al. (2008). "Cellulose–water interactions during enzymatic hydrolysis as

studied by time domain NMR." Cellulose 15(5): 703-710.

36

Fengel, D. and G. Wegener (1989). Wood: Chemistry, ultrastructure, reactions. Berlin, New York,

Walter de Gruyter.

Galbe, M. and G. Zacchi (2002). "A review of the production of ethanol from softwood." Applied

Microbiology and Biotechnology 59(6): 618-628.

Gellerstedt, G. (1973). The reactions of lignin during sulphite pulping. Stockholm.

Georgi, T., D. Rittmann, et al. (2005). "Lysine and glutamate production by Corynebacterium

glutamicum on glucose, fructose and sucrose: Roles of malic enzyme and fructose-1,6-

bisphosphatase." Metabolic Engineering 7(4): 291-301.

Ghose, T. K. (1987). "Measurement of cellulase ativities." Pure & Applied Chemistry 59(2): 257-268.

Glennie, D. W. (1971). Reactions in sulfite pulping. Lignins: Occurrence, formation, structure, and

reactions. K. V. Sarkanen and C. H. Ludwig. New York, Wiley: 597-637.

Gnansounou, E. and A. Dauriat (2005). Energy balance of bioethanol: a synthesis.

Gregg, D. J. and J. N. Saddler (1996). "Factors affecting cellulose hydrolysis and the potential of

enzyme recycle to enhance the efficiency of an integrated wood to ethanol process."

Biotechnology and Bioengineering 51(4): 375-383.

Hassinger, W., G. Sauer, et al. (1981). "The effects of equal caloric amounts of xylitol, sucrose and

starch on insulin requirements and blood glucose levels in insulin-dependent diabetics."

Diabetologia 21(1): 37-40.

Helle, S. S., R. A. Petretta, et al. (2007). "Fortifying spent sulfite pulping liquor with hydrolyzed

reject knots." Enzyme and Microbial Technology 41(1-2): 44-50.

Holtzapple, M., M. Cognata, et al. (1990). "Inhibition of Trichoderma reesei cellulase by sugars and

solvents." Biotechnology and Bioengineering 36(3): 275-287.

Hsu, T.-A. (1996). Pretreatment of biomass. Handbook on bioethanol: Production and utilization. C.

E. Wyman. Washington, Taylor & Francis: 179-212.

Jarvis, M. (2003). "Chemistry: Cellulose stacks up." Nature 426: 611-612.

Jørgensen, H., J. Vibe-Pedersen, et al. (2007). "Liquefaction of lignocellulose at high-solids

concentrations." Biotechnology and Bioengineering 96(5): 862-870.

Kai, Y. (1991). Chemistry of extractives. Wood and cellulosic chemistry. D. N.-S. Hon and N.

Shiraishi, CRC Press.

Kang, L., W. Wang, et al. (2010). "Bioconversion of Kraft Paper Mill Sludges to Ethanol by SSF and

SSCF." Applied Biochemistry and Biotechnology 161(1): 53-66.

Keating, J. D., C. Panganiban, et al. (2006). "Tolerance and adaptation of ethanologenic yeasts to

lignocellulosic inhibitory compounds." Biotechnology and Bioengineering 93(6): 1196-1206.

37

Keating, J. D., J. Robinson, et al. (2004). "An ethanologenic yeast exhibiting unusual metabolism in

the fermentation of lignocellulosic hexose sugars." Journal of Industrial Microbiology &amp;

Biotechnology 31(5): 235-244.

Keeney, D. R. and T. H. DeLuca (1992). "Biomass as an energy source for the midwestern U.S."

American Journal of Alternative Agriculture 7(03): 137-144.

Klemm, D., B. Heublein, et al. (2005). "Cellulose: Fascinating Biopolymer and Sustainable Raw

Material." Angewandte Chemie International Edition 44(22): 3358-3393.

Lee, H., C. R. Sopher, et al. (1996). "Induction of xylose reductase and xylitol dehydrogenase

activities on mixed sugars in Candida guilliermondii." Journal of Chemical Technology &

Biotechnology 65(4): 375-379.

Linden, T., J. Peetre, et al. (1992). "Isolation and characterization of acetic acid-tolerant galactose-

fermenting strains of Saccharomyces cerevisiae from a spent sulfite liquor fermentation

plant." Appl. Environ. Microbiol. 58(5): 1661-1669.

Lora, J. H. and S. Aziz (1985). "Organosolv pulping: a versatile approach to wood refining." Journal

Name: Tappi; (United States); Journal Volume: 68:8: Medium: X; Size: Pages: 94-97.

Lorenz, D., D. Morris, et al. (1995). How much energy does it take to make a gallon of ethanol?

Minneapolis, MN, Institute for Local Self-Reliance.

Mason, W. H. (1933). Process and apparatus for disintegration of material. United States Patent,

MASONITE CORP.

Meyrial, V., J. P. Delgenes, et al. (1991). "Xylitol production from D-xylose by&lt;i&gt;Candida

guillermondii&lt;/i&gt;: Fermentation behaviour." Biotechnology Letters 13(4): 281-286.

Millard, C. S., Y. P. Chao, et al. Enhanced production of succinic acid by overexpression of

phosphoenolpyruvate carboxylase in Escherichia coli.

Moritz, J. and S. Duff (1996). "Ethanol production from spent sulfite liquor fortified by hydrolysis of

pulp mill primary clarifier sludge." Applied Biochemistry and Biotechnology 57-58(1): 689-

698.

Mosier, N., C. Wyman, et al. (2005). "Features of promising technologies for pretreatment of

lignocellulosic biomass." Bioresource Technology 96(6): 673-686.

Nguyen, Q. A. and J. N. Saddler (1991). "An integrated model for the technical and economic

evaluation of an enzymatic biomass conversion process." Bioresource Technology 35(3):

275-282.

Ohara, H., K. Hiyama, et al. (1992). "Non-competitive product inhibition in lactic acid fermentation

from glucose." Applied Microbiology and Biotechnology 36(6): 773-776.

38

Ohlsson, K. A. (2004). "Carbonation of wood ash recycled to a forest soil as measured by isotope

ratio mass spectrometry." Soil Science Society of America Journal 64(6): 2155-2161.

Oleszkiewicz, J. A. and V. K. Sharma (1990). "Stimulation and inhibition of anaerobic processes by

heavy metals--A review." Biological Wastes 31(1): 45-67.

Palmqvist, E. and B. Hahn-Hägerdal (2000). "Fermentation of lignocellulosic hydrolysates. II:

inhibitors and mechanisms of inhibition." Bioresource Technology 74(1): 25-33.

Pampulha, M. E. and M. C. Loureiro-Dias (1990). "Activity of glycolytic enzymes of Saccharomyces

cerevisiae in the presence of acetic acid." Applied Microbiology and Biotechnology 34(3):

375-380.

Pan, X. (2008). "Role of Functional Groups in Lignin Inhibition of Enzymatic Hydrolysis of

Cellulose to Glucose." Journal of Biobased Materials and Bioenergy 2: 25-32.

Pimentel, D. (2003). "Ethanol Fuels: Energy Balance, Economics, and Environmental Impacts Are

Negative." Natural Resources Research 12(2): 127-134.

Pointing, S. B., A. L. Pelling, et al. (2005). "Screening of basidiomycetes and xylariaceous fungi for

lignin peroxidase and laccase gene-specific sequences." Mycological Research 109(1): 115-

124.

Prakasham, R. S., R. S. Rao, et al. (2009). "Current trends in biotechnological production of xylitol

and future prospects." Current Trends in Biotechnology and Pharmacy 3(1): 8-36.

Ramos, L., C. Breuil, et al. (1992). "Comparison of steam pretreatment of eucalyptus, aspen, and

spruce wood chips and their enzymatic hydrolysis." Applied Biochemistry and Biotechnology

34-35(1): 37-48.

Ramos, L. P., C. Breuil, et al. (1993). "The use of enzyme recycling and the influence of sugar

accumulation on cellulose hydrolysis by Trichoderma cellulases." Enzyme and microbial

technology. 15(1): 19.

Sanchez, B. and J. Bautista (1988). "Effects of furfural and 5-hydroxymethylfurfural on the

fermentation of Saccharomyces cerevisiae and biomass production from Candida

guilliermondii." Enzyme and Microbial Technology 10(5): 315-318.

Sande, W. (2010). Personal communication. L. Lai. Everett, WA.

Sedlak, M. and N. Ho (2004). "Production of ethanol from cellulosic biomass hydrolysates using

genetically engineered &lt;i&gt;saccharomyces&lt;/i&gt; yeast capable of cofermenting

glucose and xylose." Applied Biochemistry and Biotechnology 114(1): 403-416.

39

Shen, J. and F. A. Agblevor (2008). "Optimization of enzyme loading and hydrolytic time in the

hydrolysis of mixtures of cotton gin waste and recycled paper sludge for the maximum profit

rate." Biochemical Engineering Journal 41(3): 241-250.

Sjöde, A., B. Alriksson, et al. (2007). The Potential in Bioethanol Production From Waste Fiber

Sludges in Pulp Mill-Based Biorefineries. Applied Biochemistry and Biotecnology. J. R.

Mielenz, K. T. Klasson, W. S. Adney and J. D. McMillan, Humana Press: 327-337.

Sluiter, A., B. Hames, et al. (2004). Determination of sugars, byproducts, and degradation products in

liquid fraction process samples. Golden, CO, NREL. NREL/TP-510-42623.

Smith, M. T., D. R. Cameron, et al. (1997). "Comparison of industrial yeast strains for fermentation

of spent sulphite pulping liquor fortified with wood hydrolysate." Journal of Industrial

Microbiology &amp; Biotechnology 18(1): 18-21.

Taherzadeh, M. J., C. Niklasson, et al. (1997). "Acetic acid--friend or foe in anaerobic batch

conversion of glucose to ethanol by Saccharomyces cerevisiae?" Chemical Engineering

Science 52(15): 2653-2659.

Taniguchi, M., H. Suzuki, et al. (2005). "Evaluation of pretreatment with Pleurotus ostreatus for

enzymatic hydrolysis of rice straw." Journal of Bioscience and Bioengineering 100(6): 637-

643.

TAPPI (1998). TAPPI standard methods, T222 om-98. TAPPI Journal, TAPPI Press.

Tengborg, C., M. Galbe, et al. (2001). "Influence of enzyme loading and physical parameteres on the

enzymatic hydrolysis of steam-pretreated softwood." Biotechnology Progress 17(1): 110-117.

Tyson, K. S. (1993). Fuel cycle evaluations of biomass-ethanol and reformulated gasoline. Volume 1.

Other Information: PBD: Nov 1993: Medium: ED; Size: 130 p.

Väljamäe, P., K. Kipper, et al. (2003). "Synergistic cellulose hydrolysis can be described in terms of

fractal-like kinetics." Biotechnology and Bioengineering 84(2): 254-257.

von Sivers, M. and G. Zacchi (1995). "A techno-economical comparison of three processes for the

production of ethanol from pine." Bioresource Technology 51(1): 43-52.

Werpy, T., G. Petersen, et al. (2004). Top value added chemicals from biomass, Volume I: Results

from screening for potential candidates from sugars and synthesis gas. T. Werpy and G.

Petersen, NREL, Pacific Northwest National Laboratory.

Willke, T. and K. D. Vorlop (2001). "Biotechnological production of itaconic acid." Applied

Microbiology and Biotechnology 56(3): 289-295.

40

Wingren, A., M. Galbe, et al. (2003). "Techno-Economic Evaluation of Producing Ethanol from

Softwood: Comparison of SSF and SHF and Identification of Bottlenecks." Biotechnology

Progress 19(4): 1109-1117.

Wiselogel, A., S. Tyson, et al. (1996). Biomass feedstock resources and composition. Handbook on

bioethanol: Production and utilization. C. E. Wyman, Taylor & Francis: 106-117.

Xiao, Z., X. Zhang, et al. (2004). "Effects of Sugar Inhibition on Cellulases and &beta;-Glucosidase

During Enzymatic Hydrolysis of Softwood Substrates." Applied Biochemistry and

Biotechnology 115(1/3): 1115-1126.

Yazdani, S. S. and R. Gonzalez (2007). "Anaerobic fermentation of glycerol: a path to economic

viability for the biofuels industry." Current Opinion in Biotechnology 18(3): 213-219.

Yu, J., J. Zhang, et al. (2009). "Combinations of mild physical or chemical pretreatment with

biological pretreatment for enzymatic hydrolysis of rice hull." Bioresource Technology

100(2): 903-908.

Zeng, A. P., A. Ross, et al. (1994). "Multiple product inhibition and growth modeling of clostridium

butyricum and klebsiella pneumoniae in glycerol fermentation." Biotechnology and

Bioengineering 44(8): 902-911.

Zhang, X., W. Qin, et al. (2009). "High consistency enzymatic hydrolysis of hardwood substrates."

Bioresource Technology 100(23): 5890-5897.